Abstract
Membrane resident proteins are a common feature of biology yet many of these proteins are not integral to the membrane. These peripheral membrane proteins are often bound to the membrane by the addition of fatty acyl chains to the protein. This modification, known as S-acylation or palmitoylation, promotes very strong membrane association but is also reversible allowing for a high degree of control over membrane association. Many S-acylated proteins are resident in sterol, sphingolipid and saturated-lipid enriched microdomains indicating an important role for S-acylation in protein partitioning within membranes. This review summarises the current knowledge of S-acylation in plants. S-acylated proteins play a wide variety of roles in plants and affect Ca2+ signalling, K+ movement, stress signalling, small and heterotrimeric G-protein membrane association and partitioning, tubulin function as well as pathogenesis. Although the study of S-acylation is in its infancy in plants this review illustrates that S-acylation is extremely important for plant function and that there are many unexplored aspects of S-acylation in plants. A full summary of the techniques and methods available to study S-acylation in plants is also presented.
Introduction
One of the hallmarks of eukaryotic cells is the compartmentalization and segregation of different functions by membranes. Signal transduction both at and across membranes is a common feature of eukaryotic signalling. Using membranes to hold the components required for signalling increases the likelihood of interactions occurring as the space for interaction is reduced from the 3-dimensional cell to the effectively 2-dimensional membrane. This reasoning explains why many signalling complexes are formed at membranes even though they may not be transducing a signal across the membrane. Many of these complexes are regulated by altering the affinity of individual subunits for membranes leading to formation or dissolution of the complex. Proteins associating with and acting within a single membrane compartment may partition between domains of varying lipid composition which may have very different properties. These differing compartments have come to be known as membrane microdomains and include the sterol and sphingolipid enriched fractions of membranes characterized by non-ionic detergent insolubility (detergent resistant membranes – DRMs, also known as lipid rafts). There is a wealth of literature on these topics which has been comprehensively reviewed Citation[1], Citation[2]. Some proteins involved in complex formation at membranes might be integral to the membrane while others, despite their sequences indicating that they are soluble, will be found associated with membranes. Anchoring soluble proteins to membranes for the purpose of complex formation can be achieved by a number of mechanisms. Direct protein-protein interactions provide the simplest mechanism but linking a lipid group to proteins, such as S-acyl, myristoyl, farnesyl or geranylgeranyl moieties, can also promote association with membranes. The added lipid group allows membrane association by intercalation of the lipid moiety into the hydrophobic core of the membrane. Combinations of different lipid modifications provide differing strengths of membrane association Citation[3], Citation[4] and provide different attributes to the complex. Lipidation state can also change as a direct result of a complex being activated or deactivated leading to enhanced or decreased solubility of the protein in a specific membrane Citation[5]. These modifications are often accompanied by stretches of basic amino acids within the protein that interact with negatively charged membrane lipid head groups Citation[6]. The study of S-acylation in plants is still relatively undeveloped but many exciting and important aspects of plant function have been shown to be reliant upon S-acylation Citation[5], Citation[7–13].
S-acylation
N-myristoylation, the addition of 14 carbon myristoyl groups to N-terminal glycine residues, and prenylation, the addition of farnesyl or geranylgeranyl groups to CaaX C-terminal motifs in proteins are well understood and predictable hydrophobic modifications Citation[14–18]. In contrast S-acylation is very poorly understood. S-acylation involves the addition of fatty acids such as palmitate or stearate to the thiol group of specific cysteine residues within a target protein via a thioester bond. S-acylation is catalyzed by DHHC domain containing Protein S-Acyl Transferases (PATs) Citation[19], Citation[20] and frequently appears to require prior membrane association of the protein either through prenylation, myristoylation, protein interactions, polybasic domains or transmembrane spans Citation[5], Citation[9], Citation[21]. With the exception of transmembrane domains these groups and domains generally do not promote long residence times on membranes Citation[3], Citation[4] but may allow interaction with, and sampling of, cellular membranes until S-acylation occurs Citation[9]. Modification with a palmitoyl acyl group provides an affinity approximately 100×stronger than a farnesyl group and 10×stronger than a myristoyl group Citation[3], Citation[4]. Prenylated proteins are frequently processed further by RCE1/Ste24 like enzymes that remove the aaX motif to expose the prenylated cysteine at the C-terminus Citation[22]. The prenylated protein can then be further modified by methylation of the cysteine carboxyl group by isoprenyl cysteine methyl transferase (ICMT) Citation[23]. The action of ICMT is reversible and can alter the strength of membrane association considerably with a methylated prenyl cysteine providing 10- to 40-fold stronger membrane association than non-methylated prenylated cysteines Citation[4]. Despite this increase in hydrophobicity the bulky branched nature of prenyl groups generally excludes them from tightly packed or highly ordered lipid domains that S-acyl groups are able to promote association with. Longer and more saturated S-acyl chains provide even stronger association with the membrane, hence the 14 carbon myristoyl group provides a weaker affinity than the 16 carbon or greater acyl groups generally added through S-acylation Citation[3], Citation[4]. The strength of association provided by S-acylation makes it unlikely that S-acylated proteins will partition into the aqueous environment but they can still move through the lipid environment and sample different lipid domains Citation[3], Citation[4], Citation[24]. PATs are integral membrane proteins and this may explain why their substrates generally need to associate with membranes prior to S-acylation Citation[9]. No consensus sequence for S-acylation has been described despite only specific cysteines being modified in any given S-acylated protein. This indicates that specificity determinants are present, possibly via protein-protein interactions, but also may be a direct effect of an individual cysteine's chemistry. S-acylation requires a free thiolate group, and basic or aromatic residues close to the cysteine residue reduce the pKa making thiolate formation more likely. Basic or aromatic residues that promote S-acylation may be in cis or, potentially more interestingly, in trans, either in a PAT or in another interacting protein Citation[25].
S-acyl transferases in plants
Numerous DHHC domain S-acyltransferases are present in plant genome databases. Arabidopsis contains 23 DHHC PATs (AtPATs) that are divergent in sequence and only show strong homology within the DHHC domain Citation[7]. Only one plant PAT, TIP1 of Arabidopsis, has been characterized in the literature. Tip1-plants display a pleiotropic phenotype with defects in cell expansion, root hair and pollen tube growth and have an overall dwarf phenotype Citation[7], Citation[26], Citation[27]. ESTs from Viridiplantae member species indicate that DHHC PATs are found in both vascular and non-vascular plants as well as green algae (). The subcellular localization of plant PATs is unknown with the exception of TIP1 which localises predominantly to the Golgi Citation[28]. Arabidopsis PATs show developmental, tissue type, stress and environmental dependant expression indicating that AtPATs likely have discrete roles within the plant (BAR, http://www.bar.utoronto.ca/Citation[29], Hemsley, P. unpublished work).
Table I. Plants contain varying numbers of PATs. Data was obtained from TIGR/JCVI (www.tigr.org) and Gene Index (http://compbio.dfci.harvard.edu/tgi/) sequencing projects and represents current numbers based on EST data and hence may not represent the final number for a given species.
S-acylation is important as it is the only reversible lipid modification of proteins. The reversal of S-acylation is catalyzed by acyl protein thioesterases (PPTases) and is important for the cycling on and off membranes or in and out of microdomains that has been observed in many S-acylated proteins and hence aids in the regulation of S-acylated protein activity Citation[30]. Arabidopsis contains 7 putative PPTases but no data is available for them except for the NaCl stress responsive expression of At5g47330 Citation[31].
Functions of known S-acylated proteins in plants
S-acylation and GTPases – heterotrimeric G proteins
Arabidopsis contains one canonical Gα (GPA1 Citation[32]), one Gβ (AGB1 Citation[33]) and two Gγ (AGG1 Citation[34] and AGG2 Citation[35]) subunits. Only the GPA1and AGG2 subunits of the heterotrimer have been reported to be S-acylated based on subcellular localization of fluorescent protein fusions with putative S-acylation sites mutated Citation[11], Citation[12] or direct biochemical assay Citation[8]. Arabidopsis plants lacking GPA1 show defects in cell division Citation[36] and germination Citation[37] whereas AGG2 mutants show defects in gravitropism, germination and basipetal auxin transport Citation[38]. The role of S-acylation in heterotrimeric G-protein signalling is not clear but may involve ensuring that the component parts are at the correct membrane, or control association with receptors or effectors in different lipid microdomains.
GPA1 contains an N-terminal myristoylation sequence with an adjacent cysteine that is S-acylated. GPA1 S-acylation is not required for binding to either the Gβγ1 (AGB1/AGG1) or Gβγ2 (AGB1/AGG2) heterodimers or trafficking of the heterotrimer to the plasma membrane Citation[11]. It would appear however that S-acylation acts to localise GPA1 to the PM where it interacts with the Gβγ dimer and probably helps to stabilise the newly formed heterotrimer Citation[11]. Further roles may be found for S-acylation of GPA1 in regulation of signal transduction in a similar manner to Gα subunits in mammals Citation[39–41].
AGG1 and AGG2 are both prenylated and rely on dimerization with AGB1 in the case of AGG1 Citation[11] and S-acylation in the case of AGG2 Citation[12] to traffic to the plasma membrane. AGG2 is S-acylated in-planta at the Golgi before delivery to the plasma membrane Citation[11], Citation[12]. The role of S-acylation in AGG2 function beyond acting as a membrane targeting signal is unknown but may act to restrict AGG2 signalling to the PM or cycle AGG2 in and out of PM sub domains in response to stimuli. AGG1 is distributed between the plasma membrane and the Golgi and is thought to cycle between the two but not been shown to be S-acylated Citation[12].
These studies indicate that the lipid modifications of the component heterotrimer parts, when compared to animals and fungi, contribute to the unique nature of heterotrimeric G-protein signalling in plants Citation[38], Citation[42].
Type I ROPs
Type I ROPs (Rho of plants) are small, plant-unique G-proteins involved in many aspects of cellular signalling and development including polar growth, regulation of actin dynamics Citation[43], Citation[44], stomatal opening Citation[45] and reactive oxygen signalling Citation[46]. Arabidopsis contains 8 Type I ROPs (ROPs 1–8) all of which are predicted to be prenylated and recent work also indicates that they are S-acylated as a consequence of activation Citation[5]. Arabidopsis ROP6 was used to demonstrate that GDP bound (inactive) ROP6 is not acylated and is associated with the Triton X-100 detergent soluble fraction of the membrane. GTP bound (active) ROP6 is S-acylated and associated with the Triton X-100 insoluble membrane fraction (frequently interpreted as the sterol and sphingolipid enriched microdomain fraction of the membrane) where signalling complex formation is thought to occur. In support of this, AtROP6 with a C156S mutation at the presumed S-acylation site did not partition into DRMs irrespective of activation state and was not modified by acyl groups in GC-MS analysis. More importantly ROP6 was shown to cycle between soluble and insoluble fractions of the membrane in response to activation and inactivation cycles. This partitioning between membrane microdomains is though to lend further regulation to signalling complex formation by controlling association with different subsets of proteins resident in different membrane microdomains. The movement in response to a change in nucleotide binding state may be due to altered protein-protein interactions or membrane solubility of the protein once S-acylated. AtROP6 movement may also be affected by changes in conformation leading to altered lipid group penetrance or alignment within the membrane in a similar way to that predicted for H-Ras (reviewed Citation[47]). These experiments were also performed with a reportedly general anti-ROP antibody indicating that this cycling may apply to all ROPs Citation[5] although the specificity of the antibody was not tested with respect to every ROP. Interestingly the major acyl lipid attached to AtROP6 identified by GC-MS was stearic acid (C18) rather than the commonly assumed palmitic acid (C16). Whether this indicates that plants preferentially use stearic acid for S-acylation is currently unknown Citation[5].
S-acylation is catalyzed by PATs but these observations do not address which AtPAT is responsible for ROP6 S-acylation and where it resides. Due to the intimate nature of ROP6 GTP/GDP and S-acylation states it seems likely that ROP6 is acylated at the membrane where ROP activation and GTP loading occurs. Given that all ROP6 signal detected by over-expression of GFP fusions is at the PM Citation[5], ROP6 is likely to be S-acylated at the PM.
The function of S-acylation in small G-protein regulation may be extended to interactions with GDI proteins. GDI proteins are regulators of small GTPases function that sequester small GTPases from the membrane to the cytoplasm. GDIs bind to small GTPases by protein-protein contacts and insertion of the GTPase prenyl group into a hydrophobic pocket. Given the nature of small G-protein and GDI interaction it seems possible that S-acylation could act to inhibit GDI binding to small GTPases by stearicaly disrupting the interaction. S-acylation may also provide sufficiently strong membrane association that GDI proteins are unable to remove them from the membrane. Work on mammalian Rho-family small GTPases supports these view as the S-acylated RhoB and TC10 proteins were resistant to GDI induced redistribution to the cytoplasm and inhibition of S-acylation allowed GDI1 to bind to RhoB and TC10 Citation[48].
Type II ROPs
Plant type II ROPs, of which there are 3 in Arabidopsis (ROPs 9, 10 and 11), appear to be rare amongst small GTPases as, unlike Type I ROPs and many other small G-proteins, they are not prenylated and are tethered to the membrane solely by S-acylation and a polybasic domain Citation[10], Citation[21]. Type II ROPs all appear to have 2 or more S-acylated cysteines following a polybasic domain at the C-terminus Citation[10], Citation[21]. The best studied type II ROPs, AtROP9 and AtROP10, are involved in the regulation of ABA signalling Citation[49]. AtROP9 and AtROP10 contain 3 and 2 S-acylatable cysteines in their respective C-terminals. AtROPs 9 and 10 are tightly associated with the PM and no signal is detected in cytoplasmic fractions by GFP fusion fluorescence or by immunoblot. AtROP10 requires both cysteines to be acylated for membrane attachment as GFP-AtROP10 fusions lacking either or both cysteines were found in the cytoplasmic fraction of the cell (Figure 1) Citation[10]. ROP10 lacking the polybasic domain is also cytoplasmic Citation[21] and may indicate that the polybasic domain is required for acylation to occur and is consistent with the commonly observed requirement for membrane association prior to S-acylation. Mutation of glycine residues adjacent to the acylatable cysteines of AtROP10 resulted in a reduction, but not abolition, of membrane association. Substitution of glycine for another amino acid may act to stearicaly hinder the acylation reaction or destabilise thiolate formation. Also often found adjacent to the acylatable cysteines of type II ROPs are one to two serine residues and one to two hydrophobic residues Citation[21]. The hydrophobic nature of these latter residues coupled with the small size and non-polar nature of the glycine residues may act to push the acyl group through the charged head groups of the membrane further into the bilayer. This may provide stronger anchorage and protect the thioester bond from being cleaved by PPTases thereby increasing residence time on the membrane. Interactions between these hydrophobic amino acids and the lipid tails of the membrane may provide further strength to the association Citation[47]. The proximal serine residues might also play an important role. Phosphorylation of serine residues increases the negative charge of the residue leading to repulsive forces between the negatively charged cytosolic membrane face and phosphoserine residues. This has been previously shown to decrease affinity for the membrane Citation[50] and may be involved in altering membrane affinity of type II ROPs leading to altered membrane partitioning during activation cycles.
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09 July 2009Figure 1. ROP10 requires S-acylation for localization to the plasma membrane. Deletion of either or both S-acylated cysteines in the ROP10 C-terminus leads to ROP10 being found largely in the cytoplasm and nucleoplasm indicating that ROP10 requires dual S-acylation for stable membrane association. (A) GFP-ROP10; (B) GFP-ROP10 C199S; (C) GFP-ROP10 C205S; (D) GFP-ROP10 C199,205S. Scale bar represents 10 µm.
Figure 1. ROP10 requires S-acylation for localization to the plasma membrane. Deletion of either or both S-acylated cysteines in the ROP10 C-terminus leads to ROP10 being found largely in the cytoplasm and nucleoplasm indicating that ROP10 requires dual S-acylation for stable membrane association. (A) GFP-ROP10; (B) GFP-ROP10 C199S; (C) GFP-ROP10 C205S; (D) GFP-ROP10 C199,205S. Scale bar represents 10 µm.
AtRABF1
AtRABF1, previously known as ARA6, is a plant specific Rab5 GTPase homologue that is N-myristoylated and S-acylated rather than the more usual dual geranylgeranylation found on Rab proteins Citation[51]. AtRABF1 localises to a specific subset of vesicles, most likely a prevacuolar compartment (PVC), and S-acylation is essential for PVC localization with solely myristoylated AtRABF1 remaining largely on the ER. The lack of prenyl groups on AtRABF1 also prevents interaction with AtRAB GDI1 which is thought to sequester inactive RAB proteins in the cytosol. AtRABF1 is observed in the cytosol which may indicate that there is another AtRAB GDI capable of binding AtRABF1. AtRABF1 might also dissociate from membranes in its non-S-acylated state making S-acylation essential for stable membrane association and removing the need for a GDI protein. Other unknown factors act through the C-terminus to restrict AtRABF1 solely to the PVC as GFP fused to the N-terminal 36 amino acids containing the S-acylation motif accumulated on the PM and punctate structures whereas a combination of the N-terminal 34 and C-terminal 8 amino acids are able to direct the fusion to endosomes Citation[51]. AtRABF1 also partially co-localizes with the sterol reactive dye filipin on the PVC Citation[52]. This raises the possibility that N-myristoylation and S-acylation of AtRABF1 promote association with sterol enriched areas of the PM and facilitate their endocytosis. AtRABF1 might be involved in the endocytosis of sterols and sterol enriched microdomain associated proteins.
S-acylation and Ca2+ signalling
CBLs
CBLs (Calcineurin B-Like proteins) are calcium binding proteins that bind to CBL-interacting protein kinases (CIPKs) in a calcium dependant manner. Each of the 10 CBLs in Arabidopsis interacts with specific CIPKs allowing for a structured interpretation of calcium signalling. CBL/CIPK complexes decode Ca2+ signals involved in a range of processes including K+ transport Citation[53], salt tolerance Citation[54] and ABA signalling Citation[55]. Recent work Citation[9] has shown that CBL1 is myristoylated and S-acylated and associates with the plasma membrane. Solely myristoylated CBL1 is found on ER, nuclear and plasma membranes whereas CBL1 lacking both lipid modifications is predominantly cytoplasmic (Figure 2). These observations indicate that S-acylation determines the plasma membrane localization of CBL1 whereas myristoylation allows for the sampling of membranes or retention at the ER until it encounters the CBL1 S-acyltransferase. As a large proportion of solely myristoylated CBL1 is associated with the ER it is tempting to speculate that that the S-acyltransferase for CBL1 also resides in the ER. Interestingly dual lipid modification of CBL1 is also essential for plasma membrane localization of the CBL1/CIPK1 dimer. CBL1 lipidation mutants still form a dimer with CIPK1 but are distributed in a largely cytoplasmic manner. The localizations of full length CBL1 fusions can be mimicked with the N-terminal 12 amino acids indicating that these residues contain the targeting signals Citation[9].
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09 July 2009Figure 2. CBL1 is N-myristoylated and S-acylated and both modifications are required for membrane association. (A) CBL1-YFP is found at the plasma membrane. (B) CBL1-YFP G2A mutants lacking the N-myristoylated glycine are found in the cytoplasm and nucleoplasm. (C) CBL1-YFP C3S mutants lacking the S-acylated cysteine are found associated with most cellular membranes with a small amount in the cytoplasm and nucleoplasm. This suggests that CBL1 S-acylation requires prior N-myristoylation. Scale bar represents 10 µm.
Figure 2. CBL1 is N-myristoylated and S-acylated and both modifications are required for membrane association. (A) CBL1-YFP is found at the plasma membrane. (B) CBL1-YFP G2A mutants lacking the N-myristoylated glycine are found in the cytoplasm and nucleoplasm. (C) CBL1-YFP C3S mutants lacking the S-acylated cysteine are found associated with most cellular membranes with a small amount in the cytoplasm and nucleoplasm. This suggests that CBL1 S-acylation requires prior N-myristoylation. Scale bar represents 10 µm.
CPKs
Calcium dependent protein kinases (CPKs) are calcium sensing kinases and many of them have been linked to stress and ABA signalling Citation[56]. Many CPKs are N-myristoylated and also contain cysteine residues nearby which may be S-acylated Citation[57]. The rice CPK OsCPK2 is the only CPK to have been shown biochemically to be modified by myristate and palmitate and that S-acylation depends upon prior myristoylation Citation[13]. Mutagenesis data obtained from the Tomato CPK LeCRK also supports the idea that myristoylation and S-acylation aid in membrane association. LeCRK contains 2 cysteine residues at positions four and five. WT LeCRK:GFP is observed at the plasma membrane while LeCRK1:GFP with a mutated myristoylation site is cytoplasmic and LeCRK1:GFP lacking cysteines 4 and 5 is observed on all cellular membranes. This indicates a role for S-acylation in specification of LeCRK plasma membrane localization and that myristoylation of LeCRK1 is required for S-acylation of cysteines 4 and 5 Citation[58]. A similar situation is observed in Medicago truncatula with MtCPK3 involved in root nodule formation, Citation[59] and Solanum tuberosum with StCDPK1 involved in tuber formation Citation[60].
Arabidopsis contains 34 CPKs of which 27 have an N-myristoylation site followed by one to two cysteines which may be potential S-acylation sites. A study of the subcellular localizations of AtCPKs indicated that those with one (AtCPK9, 16, 21, 28) or two (AtCPK7, 8) cysteine residues adjacent to a myristoylated glycine may bind to the plasma membrane Citation[61]. AtCPK1 and AtCPK2 have one cysteine following the myristoylation site and localise to peroxisomes Citation[61] and the ER Citation[62] respectively. The solely myristoylated AtCPK3 associates with membranes on a transient basis Citation[61]. This indicates that although myristoylation and S-acylation do aid in anchoring CPKs to membranes there is still no way to accurately determine which membranes are targeted based on lipidation state. Specific membrane localization may be achieved by association with other proteins or determined by other factors such as basic or phosphorylated residues.
S-acylation and pathogenesis
Bacterial type III effector proteins
Pathogenic bacteria are capable of secreting proteins using the type III secretory pathway to interact with and modulate a host cells response to the pathogen. Interestingly, many of these secreted bacterial effectors are modified by N-myristoylation and S-acylation in the host cell and these modifications are required for activity. Pseudomonas syringae avrRpm1 is both N-myristoylated and S-acylated by the host cell and localises to the plasma membrane. Deletion of either the N-myristoylation site or S-acylation site impaired virulence of avrRpm1 with loss of N-myristoylation resulting in greater impairment. This is in agreement with the view that S-acylation requires prior membrane association, in this case by N-myristoylation. It is unknown whether S-acylation acts solely as a membrane anchor or if it also plays a regulatory role in avrRpm1 function. The authors report that the P. syringae avrB, avrPphB, avrC and avrPto Type III effectors also contain sequences likely to be N-myristoylated and S-acylated Citation[63].
XopE1 and XopE2, members of the HopX2 effector family, and XopJ, an avrRxv family effector, from Xanthomonas campestris have also been reported to be N-myristoylated and S-acylated and localize to the PM. Sequence analysis of both Pseudomonas spp. and Xanthamonas spp. genomes indicates that many type III effectors are potentially N-myristoylated and S-acylated Citation[64]. These data are interesting as they represent another host-pathogen interaction whereby host processes are subverted to aid the pathogen.
RIN4
RIN4 is involved in defending against Pseudomonas syringae attack by acting as a foreign protein monitoring system. Pathogen derived proteins affect the state or stability of RIN4 leading to a cellular response. RIN4 is C-terminally acylated at 3 cysteines and anchored to the PM Citation[65]. Membrane localization of RIN4 is required for interaction with the membrane bound P. syringae type III effectors avrRpm1 and avrB which are themselves S-acylated. avrRpm1 and avrB act to phosphorylate RIN4 leading to the activation of their cognate R protein RPM1 and increased resistance to P. syringae.
S-acylation of RIN4 is also important as mutant non-acylatable forms of RIN4 are degraded by the proteasome indicating that S-acylation acts to protect RIN4 from degradation in the non-P. syringae challenged state. The cytosolic P. syringae type III effector protease AvrRpt2 cleaves RIN4 leaving a short C-terminal S-acylated portion of RIN4 on the membrane while the remaining, now cytosolic, portion is degraded by the proteasome Citation[65] and repression of RPS2 activation is relieved leading to increased defence responses against P. syringaeCitation[66–68]. Plants lacking RIN4 are embryo lethal due to constitutive activation of RPS2 and cell death Citation[68]. The rapid degradation of non-acylated RIN4 would also be expected to be embryo lethal. Thus S-acylation is important for both aspects of RIN4 based defence responses.
Viral proteins
East African Cassava Mosaic Cameroon Virus (EACMCV) AC4 protein is involved in suppressing the systemic post transcriptional gene silencing (PTGS) signal in response to virus invasion. AC4 is membrane localized and contains a myristoylation motif adjacent to a potential S-acylation site. Mutation of the myristoylated glycine (G2A) leads to a loss of AC4-GFP localization at the plasma membrane whereas mutation of the potential S-acylation site (C3A) leads to a localization reminiscent of the ER. Functionally, AC4 G2A mutants do not induce EACMCV like symptoms nor suppress systemic PTGS. AC4 C3A mutants showed reduced systemic PTGS suppression and later onset of viral symptoms compared to WT AC4. This indicates that membrane localization and S-acylation of AC4 is required for full virus infection and suppression of gene silencing Citation[69].
Other S-acylated proteins in plants
Tubulin
α-tubulin has long been known to be S-acylated and the major site for S-acylation has been identified in the yeast α-tubulin TUB1 protein as Cys377Citation[70]. Plant α-tubulin subunits contain an analogous residue making S-acylation of this residue a distinct possibility. Recently α-tubulin subunits from Arabidopsis were shown to be S-acylated but no specific cysteine residue was identified Citation[8]. The exact role of S-acylation in α-tubulin function is unknown but appears to aid in spindle positioning and astral microtubule turnover in budding yeast during cell division Citation[70]. S-acylation also reduces the polymerization competence of α-tubulin monomers with progressively less S-acylated α-tubulin being incorporated into microtubules over repeated polymerization/depolymerization cycles in vitroCitation[71]. This indicates that either α-tubulin is S-acylated after polymerization or is incorporated at a reduced rate compared to non-S-acylated α-tubulin. It should be noted however that this may be due to the hydrophobic acyl groups being poorly soluble in the assay buffer used leading to aggregation of S-acylated α-tubulin subunits and rendering them non-competent for polymerization. Most S-acylated tubulin is found in the membrane fraction of cells but a small proportion is found in the cytosolic fraction of the cell indicating that the role of tubulin S-acylation may not be solely to interact with membranes Citation[72]. An interesting possibility is that S-acylation of tubulin could act to bind cortical microtubules to membranes and aid in maintaining their alignment which may in turn affect cell wall deposition. An untested theory is that a bulky hydrophobic group such as an acyl chain could help or hinder microtubule bundling and may even help nucleation of new microtubules at membranes. Finally S-acylated microtubules may help to link and maintain the positions of membrane compartments within the cell which may be important for directional growth and cellular organization.
Chloroplast photosystem II protein D1
Chloroplast photosystem II (PSII) protein D1 encoded by the plastid psbA gene is an essential part of the PSII complex and acts in the electron transport chain. D1 is rapidly S-acylated in a light dependant manner in the stromal lamellae. D1 is then de-acylated in the granal lamellae after incorporation into PSII Citation[73]. This is of interest as no DHHC PATs are encoded in organellar genomes in any organism indicating that there may be a non-DHHC prokaryotic protein S-acyltransferase or that a DHHC S-acyl transferase is imported into the chloroplast. Alternatively D1 may undergo auto-S-acylation as the chloroplast is one of the main sites for lipid synthesis in plants Citation[74] and S-acylation has been shown to be auto-catalytic under extremely high acyl-CoA concentrations Citation[71].
Indirect consequences of protein acylation
An effect of acylation that may be independent of the previously published or described functions of S-acylated proteins has been reported in mammalian filipodia formation Citation[75]. The authors report that filipodia could be induced by the 12–16 amino acids containing the S-acylation sites of the functionally unrelated proteins GAP-43 and paralemmin fused to GFP but not by those of PSD-95 or PSD-93. All four S-acylated regions are similar, containing basic amino acids and two S-acylated cysteines. Mutation of the GAP-43 and paralemmin S-acylated cysteines prevented filipodial formation indicating that S-acylation is required for the observed effects Citation[75]. These results demonstrate that S-acylation may be directly responsible for some of the observed biological effects of some S-acylated proteins. These effects of S-acylation are possibly due to stabilization of membrane microdomains responsible for organising signalling complexes during filipodial expansion. Another explanation is that changes in membrane tension are created due to intercalation of the acyl moieties of S-acylated proteins. Filipodial expansion is an actin dependent process and it has been postulated that actin polymerization is inhibited by the tension of the membrane Citation[76]. A reduction in membrane tension has been shown to allow actin polymerization to occur Citation[76] and the theory behind it dubbed the Brownian ratchet model Citation[77].
It remains to be seen whether this mechanism acts in plants but root hairs, pollen tubes and to a lesser extent many other plant cell types exhibit polarized growth. Root hairs and pollen tubes are much finer projections than filipodia and like all plant cells are constrained by the cell wall which may prevent the membrane movement necessary for the Brownian-ratchet theory. On the other hand filipodial expansion is dependant upon ARF and CDC42 (RHO) GTPase control of actin dynamics Citation[75] and the plant GTPase ROP2 affects actin dynamics and is reliant upon ARF GTPases for localization to sites of polarized growth in root hairs Citation[43], Citation[78] demonstrating that the cellular machinery required is present.
Approaches for studying S-acylation in plants
Predictive approaches
Despite the lack of a motif for S-acylation two web-based applications are available to predict S-acylation sites in proteins. CSS-palm Citation[79] and NBA-palm Citation[80] are probably a good starting point for those wishing to identify residues for point mutagenesis studies although there are no published validations of their predictions.
Mutational analysis
Mutational analysis often provides the first evidence for protein S-acylation and can be used in conjunction with biochemical and microscopy approaches to determine the effects of S-acylation on membrane association. Candidate cysteine residues are typically mutated to either serine or alanine.
Inhibition of S-acylation
The palmitate analogue 2-bromopalmitate (2-bromohexadecanoic acid) has proved to be a potent inhibitor of S-acylation but care should be taken in the interpretation of data as it also inhibits N-myristoylation, albeit to a lesser extent Citation[81]. 2-bromopalmitate has been successfully used in plants to look at both phenotypes and subcellular localization changes of fluorescent fusion proteins either through direct application onto plate grown plants Citation[8] or by infiltration into leaves Citation[10], Citation[12]. Two other inhibitors have been used in mammals and yeast but are untried in plants. Cerulenin is thought to directly inhibit PAT activity but also binds β-ketoacyl acyl-carrier protein synthase and HMG-CoA synthase and inhibits fatty acid and sterol synthesis respectively Citation[82]. Tunicamycin has been used as an N-glycosylation inhibitor but has also shown activity as an S-acylation inhibitor Citation[83].
Alteration of lipid content of cells
Although largely restricted to cell culture experiments changing the lipid content of a cell allows for different acyl groups to be incorporated into proteins and may alter their partitioning into membrane microdomains Citation[81], Citation[84]. This technique may be useful for uncoupling effects caused by association of proteins with membranes from those caused by sterol and sphingolipid enriched microdomain partitioning but probably also affects the overall composition of the membrane so results should be treated with caution.
In-vivo labelling using radioactive fatty acids
In-vivo labelling with 3H-palmitic acid has long been the main method for determining the S-acylation state of proteins. This is a relatively simple, if costly and time consuming, procedure for cells grown in culture but is limited in its application to plants where the ability to work on intact organs in the context of a whole plant is one of the major advantages of plant biology. Hydroxylamine cleavage of the 3H-palmitate from the protein is frequently used to demonstrate the incorporation of the label via a thioester bond indicative of S-acylation. Failure to cleave indicates that secondary metabolites have likely been incorporated. For this reason 3H-palmitate labelling is usually done for a short period of time after inhibition of cellular fatty acid synthesis. 125I labelled 16-iodo-hexadecanoic acid (IC16) has also been used successfully and is rapidly visualized compared to 3H-palmitate but requires synthesis and is much more hazardous to work with Citation[85].
Biotin switch assay for S-acylation
The development Citation[86] and subsequent refinement for use with plants Citation[8] of the biotin switch assay allows for faster and more sensitive assay of protein S-acylation that is particularly suited to whole organism biology. This approach also has the advantage that no external interference occurs, such as addition of radiolabelled palmitate, before measuring acylation state. The biotin switch method for assaying S-acylation, although an indirect approach, does not require special equipment, large quantities of material or the use of radioactive sources and can be performed in any laboratory familiar with Western blotting making it very useful for rapidly determining if a protein is S-acylated Citation[8].
GC-MS
The GC-MS techniques used to directly analyse the composition of ROP6 Citation[5] and CBL1 Citation[9] covalently bound lipid groups allows the quantitative identification of the lipid groups added. The high resolution of this technique and non-invasive nature before assaying is offset by it being ultimately more costly, time consuming and requiring more material and specialist equipment than the biotin switch assay. This method is ideally suited to a detailed analysis of an S-acylated protein. Further development of MS/MS techniques would hopefully allow direct identification of acylated peptides without the need for mutagenesis or biochemical assay.
Microscopy techniques
Direct analysis of fluorescent fusion protein membrane association has proven very useful in the analysis of S-acylated proteins. More advanced techniques such as FRAP have been used to look at delivery and recycling of proteins between the PM and Golgi. FRAP has also been used to discriminate lateral diffusion of S-acylated H-Ras from cytoplasmic exchange and thereby measure strength of association of various S-acylated forms of H-Ras and assess the contribution each acyl group makes to the overall affinity of H-Ras for membranes Citation[87], Citation[88]. FLIM-FRET and immunogold electron microscopy have also been used to examine H-Ras S-acylation dependant clustering with Raf1 Citation[89].
Concluding remarks
The study of S-acylation in plants has lagged behind that of the mammalian and yeast systems for many years. Despite this, data on S-acylated proteins in plants is coming to light and much of that information is summarized in this review. The data presented indicates that there are unique S-acylated proteins in plants as well as those conserved with other eukaryotic systems. S-acylation is recognized as being a major factor in many aspects of plant biology but is likely to be involved in many more. Due to the difficulties in determining the S-acylation state of proteins based on sequence, proteomics approaches may yield valuable and interesting data at a faster rate than would otherwise be possible using a traditional ‘protein at a time’ approach. These methods are also more likely to identify novel S-acylated proteins as well as identify S-acylation sites for already characterized proteins. Plants are also well poised to investigate the role of S-acylation in a multi-cellular context which has so far not been addressed in any organism.
Acknowledgements
Apologies are offered to those colleagues whose work was not cited due to space constraints. Shaul Yalovsky and Jörg Kudla kindly provided data for Figures 1 and 2. I would like to thank Angharad Jones for critical reading of this manuscript.
Declaration of interest: The author reports no conflicts of interest. The author alone is responsible for the content and writing of the paper.
| Species | Common name | PATs |
|---|---|---|
| Dicots | ||
| Aquilegia formosa x pubescens | Aquilegia | 13 |
| Arabidopsis thaliana | Thale Cress | 23 |
| Beta vulgaris | Beet | 2 |
| Brassica napus | Oil seed rape | 24 |
| Citrus clementina | Clementine | 9 |
| Citrus sinensis | Orange | 12 |
| Coffea canephora | Coffee | 4 |
| Euphorrbia esula | Leafy Spurge | 6 |
| Glycine max | Soy bean | 12 |
| Gossypium sp. | Cotton | 26 |
| Helianthus annuus | Sunflower | 7 |
| Ipomea nil | Morning glory | 5 |
| Lactuca sativa | Lettuce | 7 |
| Lactuca serriola | Prickly lettuce | 5 |
| Lotus japonicus | Lotus | 1 |
| Lycopersicon esculentum | Tomato | 10 |
| Malus x domestica | Apple | 8 |
| Medicago trunctula | Barrel medick | 9 |
| Nicotiana benthamiana | Tobacco | 2 |
| Nicotiana tobaccum | Tobacco | 5 |
| Petunia vulgaris | Petunia | 1 |
| Populus sp. | Poplar | 9 |
| Prunus persica | Peach | 1 |
| Saccharum officinarum | Sugar cane | 20 |
| Solanum tuberosum | Potato | 19 |
| Vitis vinifera | Grape | 3 |
| Monocots | ||
| Alium cepa | Onion | 5 |
| Hordeum vulgare | Barley | 9 |
| Oryza sativa cv. japonica | Rice | 51 |
| Sorghum bicolour | Sorghum | 3 |
| Triticum aestivum | Wheat | 16 |
| Zea mays | Maize | 36 |
| Gymnosperms | ||
| Picea sp. | Spruce | 5 |
| Pinus sp. | Pine | 11 |
| Bryophytes | ||
| Physcomitrella patens | Moss | 20 |
| Green algae | ||
| Ostreococcus tauri | Ostreococcus | 10 |
References
- Harding AS, Hancock JF. Using plasma membrane nanoclusters to build better signaling circuits. Trends Cell Biol 2008; 18: 364–371 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Hancock JF. Lipid rafts: contentious only from simplistic standpoints. Nat Rev Mol Cell Biol 2006; 7: 456–462 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Shahinian S, Silvius JR. Doubly-lipid-modified protein sequence motifs exhibit long-lived anchorage to lipid bilayer membranes. Biochemistry 1995; 34: 3813–3822 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Silvius JR, l'Heureux F. Fluorimetric evaluation of the affinities of isoprenylated peptides for lipid bilayers. Biochemistry 1994; 33: 3014–3022 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Sorek N, Poraty L, Sternberg H, Bar E, Lewinsohn E, Yalovsky S. Activation status-coupled transient S acylation determines membrane partitioning of a plant Rho-related GTPase. Mol Cell Biol 2007; 27: 2144–2154 [Taylor & Francis Online], [Web of Science ®], [Google Scholar]
- Lorenz CD, Faraudo J, Travesset A. Hydrogen bonding and binding of polybasic residues with negatively charged mixed lipid monolayers. Langmuir 2008; 24: 1654–1658 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Hemsley PA, Kemp AC, Grierson CS. The Tip Growth Defective1 S-acyl transferase regulates plant cell growth in Arabidopsis. Plant Cell 2005; 17: 2554–2563 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Hemsley PA, Taylor L, Grierson CS. Assaying protein palmitoylation in plants. Plant Methods 2008; 4: 2 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Batistic O, Sorek N, Schultke S, Yalovsky S, Kudla J. Dual fatty acyl modification determines the localization and plasma membrane targeting of CBL/CIPK Ca2+ signaling complexes in Arabidopsis. Plant Cell 2008; 20: 1346–1362 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Lavy M, Bracha-Drori K, Sternberg H, Yalovsky S. A cell-specific, prenylation-independent mechanism regulates targeting of type II RACs. Plant Cell 2002; 14: 2431–2450 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Adjobo-Hermans MJ, Goedhart J, Gadella TW, Jr. Plant G protein heterotrimers require dual lipidation motifs of Galpha and Ggamma and do not dissociate upon activation. J Cell Sci 2006; 119: 5087–5097 [Crossref], [PubMed], [Google Scholar]
- Zeng Q, Wang X, Running MP. Dual lipid modification of Arabidopsis Ggamma-subunits is required for efficient plasma membrane targeting. Plant Physiol 2007; 143: 1119–1131 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Martin ML, Busconi L. Membrane localization of a rice calcium-dependent protein kinase (CDPK) is mediated by myristoylation and palmitoylation. Plant J 2000; 24: 429–435 [Crossref], [PubMed], [Google Scholar]
- Running MP, Lavy M, Sternberg H, Galichet A, Gruissem W, Hake S, Ori N, Yalovsky S. Enlarged meristems and delayed growth in plp mutants result from lack of CaaX prenyltransferases. Proc Natl Acad Sci USA 2004; 101: 7815–7820 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Martinez A, Traverso JA, Valot B, Ferro M, Espagne C, Ephritikhine G, Zivy M, Giglione C, Meinnel T. Extent of N-terminal modifications in cytosolic proteins from eukaryotes. Proteomics 2008; 8: 2809–2831 [Crossref], [PubMed], [Google Scholar]
- Austin RS, Provart NJ, Cutler SR. C-terminal motif prediction in eukaryotic proteomes using comparative genomics and statistical over-representation across protein families. BMC Genomics 2007; 8: 191 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Maurer-Stroh S, Koranda M, Benetka W, Schneider G, Sirota FL, Eisenhaber F. Towards complete sets of farnesylated and geranylgeranylated proteins. PLoS Comput Biol 2007; 3: e66 [Crossref], [PubMed], [Google Scholar]
- Marmagne A, Ferro M, Meinnel T, Bruley C, Kuhn L, Garin J, Barbier-Brygoo H, Ephritikhine G. A high content in lipid-modified peripheral proteins and integral receptor kinases features in the arabidopsis plasma membrane proteome. Mol Cell Proteomics 2007; 6: 1980–1996 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Lobo S, Greentree WK, Linder ME, Deschenes RJ. Identification of a Ras palmitoyltransferase in Saccharomyces cerevisiae. J Biol Chem 2002; 277: 41268–41273 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Roth AF, Feng Y, Chen L, Davis NG. The yeast DHHC cysteine-rich domain protein Akr1p is a palmitoyl transferase. J Cell Biol 2002; 159: 23–28 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Lavy M, Yalovsky S. Association of Arabidopsis type-II ROPs with the plasma membrane requires a conserved C-terminal sequence motif and a proximal polybasic domain. Plant J 2006; 46: 934–947 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Boyartchuk VL, Ashby MN, Rine J. Modulation of Ras and a-factor function by carboxyl-terminal proteolysis. Science 1997; 275: 1796–1800 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Hrycyna CA, Sapperstein SK, Clarke S, Michaelis S. The Saccharomyces cerevisiae STE14 gene encodes a methyltransferase that mediates C-terminal methylation of a-factor and RAS proteins. Embo J 1991; 10: 1699–1709 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Peitzsch RM, McLaughlin S. Binding of acylated peptides and fatty acids to phospholipid vesicles: pertinence to myristoylated proteins. Biochemistry 1993; 32: 10436–10443 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Dietrich LE, Ungermann C. On the mechanism of protein palmitoylation. EMBO Rep 2004; 5: 1053–1057 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Ryan E, Grierson CS, Cavell A, Steer M, Dolan L. TIP1 is required for both tip growth and non-tip growth in Arabidopsis. New Phytol 1998; 138: 49–58 [Crossref], [Google Scholar]
- Schiefelbein J, Galway M, Masucci J, Ford S. Pollen tube and root-hair tip growth is disrupted in a mutant of Arabidopsis thaliana. Plant Physiol 1993; 103: 979–985 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Dunkley TP, Watson R, Griffin JL, Dupree P, Lilley KS. Localization of organelle proteins by isotope tagging (LOPIT). Mol Cell Proteomics 2004; 3: 1128–1134 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Toufighi K, Brady SM, Austin R, Ly E, Provart NJ. The botany array resource: e-northerns, expression angling, and promoter analyses. Plant J 2005; 43: 153–163 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Lu JY, Hofmann SL. Inefficient cleavage of palmitoyl-protein thioesterase (PPT) substrates by aminothiols: implications for treatment of infantile neuronal ceroid lipofuscinosis. J Inherit Metab Dis 2006; 29: 119–126 [Crossref], [PubMed], [Google Scholar]
- Gong Z, Koiwa H, Cushman MA, Ray A, Bufford D, Kore-eda S, Matsumoto TK, Zhu J, Cushman JC, Bressan RA, Hasegawa PM. Genes that are uniquely stress regulated in salt overly sensitive (sos) mutants. Plant Physiol 2001; 126: 363–375 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Ma H, Yanofsky MF, Meyerowitz EM. Molecular cloning and characterization of GPA1, a G protein alpha subunit gene from Arabidopsis thaliana. Proc Natl Acad Sci USA 1990; 87: 3821–3825 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Lease KA, Wen J, Li J, Doke JT, Liscum E, Walker JC. A mutant Arabidopsis heterotrimeric G-protein beta subunit affects leaf, flower, and fruit development. Plant Cell 2001; 13: 2631–2641 [PubMed], [Web of Science ®], [Google Scholar]
- Mason MG, Botella JR. Completing the heterotrimer: isolation and characterization of an Arabidopsis thaliana G protein gamma-subunit cDNA. Proc Natl Acad Sci USA 2000; 97: 14784–14788 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Mason MG, Botella JR. Isolation of a novel G-protein gamma-subunit from Arabidopsis thaliana and its interaction with Gbeta. Biochim Biophys Acta 2001; 1520: 147–153 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Ullah H, Chen JG, Young JC, Im KH, Sussman MR, Jones AM. Modulation of cell proliferation by heterotrimeric G protein in Arabidopsis. Science 2001; 292: 2066–2069 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Ullah H, Chen JG, Wang S, Jones AM. Role of a heterotrimeric G protein in regulation of Arabidopsis seed germination. Plant Physiol 2002; 129: 897–907 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Trusov Y, Rookes JE, Tilbrook K, Chakravorty D, Mason MG, Anderson D, Chen JG, Jones AM, Botella JR. Heterotrimeric G protein gamma subunits provide functional selectivity in Gbetagamma dimer signaling in Arabidopsis. Plant Cell 2007; 19: 1235–1250 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Pedone KH, Hepler JR. The importance of N-terminal polycysteine and polybasic sequences for G14alpha and G16alpha palmitoylation, plasma membrane localization, and signaling function. J Biol Chem 2007; 282: 25199–25212 [Crossref], [PubMed], [Google Scholar]
- Wedegaertner PB, Chu DH, Wilson PT, Levis MJ, Bourne HR. Palmitoylation is required for signaling functions and membrane attachment of Gq alpha and Gs alpha. J Biol Chem 1993; 268: 25001–25008 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Hughes TE, Zhang H, Logothetis DE, Berlot CH. Visualization of a functional Galpha q-green fluorescent protein fusion in living cells. Association with the plasma membrane is disrupted by mutational activation and by elimination of palmitoylation sites, but not be activation mediated by receptors or AlF4. J Biol Chem 2001; 276: 4227–4235 [PubMed], [Web of Science ®], [Google Scholar]
- Wu Y, Xu X, Li S, Liu T, Ma L, Shang Z. Heterotrimeric G-protein participation in Arabidopsis pollen germination through modulation of a plasmamembrane hyperpolarization-activated Ca2 + -permeable channel. New Phytol 2007; 176: 550–559 [Crossref], [PubMed], [Google Scholar]
- Jones MA, Shen JJ, Fu Y, Li H, Yang Z, Grierson CS. The Arabidopsis Rop2 GTPase is a positive regulator of both root hair initiation and tip growth. Plant Cell 2002; 14: 763–776 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Fu Y, Gu Y, Zheng Z, Wasteneys G, Yang Z. Arabidopsis interdigitating cell growth requires two antagonistic pathways with opposing action on cell morphogenesis. Cell 2005; 120: 687–700 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Jeon BW, Hwang JU, Hwang Y, Song WY, Fu Y, Gu Y, Bao F, Cho D, Kwak JM, Yang Z, Lee Y. The Arabidopsis small G protein ROP2 is activated by light in guard cells and inhibits light-induced stomatal opening. Plant Cell 2008; 20: 75–87 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Foreman J, Demidchik V, Bothwell JH, Mylona P, Miedema H, Torres MA, Linstead P, Costa S, Brownlee C, Jones JD, Davies JM, Dolan L. Reactive oxygen species produced by NADPH oxidase regulate plant cell growth. Nature 2003; 422: 442–446 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Abankwa D, Gorfe AA, Hancock JF. Ras nanoclusters: molecular structure and assembly. Semin Cell Dev Biol 2007; 18: 599–607 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Michaelson D, Silletti J, Murphy G, D'Eustachio P, Rush M, Philips MR. Differential localization of Rho GTPases in live cells: regulation by hypervariable regions and RhoGDI binding. J Cell Biol 2001; 152: 111–126 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Zheng ZL, Nafisi M, Tam A, Li H, Crowell DN, Chary SN, Schroeder JI, Shen J, Yang Z. Plasma membrane-associated ROP10 small GTPase is a specific negative regulator of abscisic acid responses in Arabidopsis. Plant Cell 2002; 14: 2787–2797 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- McLaughlin S, Aderem A. The myristoyl-electrostatic switch: a modulator of reversible protein-membrane interactions. Trends Biochem Sci 1995; 20: 272–276 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Ueda T, Yamaguchi M, Uchimiya H, Nakano A. Ara6, a plant-unique novel type Rab GTPase, functions in the endocytic pathway of Arabidopsis thaliana. Embo J 2001; 20: 4730–4741 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Grebe M, Friml J, Swarup R, Ljung K, Sandberg G, Terlou M, Palme K, Bennett MJ, Scheres B. Cell polarity signaling in Arabidopsis involves a BFA-sensitive auxin influx pathway. Curr Biol 2002; 12: 329–334 [Crossref], [PubMed], [Google Scholar]
- Li L, Kim BG, Cheong YH, Pandey GK, Luan S. A Ca(2)+ signaling pathway regulates a K(+) channel for low-K response in Arabidopsis. Proc Natl Acad Sci USA 2006; 103: 12625–12630 [Crossref], [PubMed], [Google Scholar]
- Cheong YH, Kim KN, Pandey GK, Gupta R, Grant JJ, Luan S. CBL1, a calcium sensor that differentially regulates salt, drought, and cold responses in Arabidopsis. Plant Cell 2003; 15: 1833–1845 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Pandey GK, Cheong YH, Kim KN, Grant JJ, Li L, Hung W, D'Angelo C, Weinl S, Kudla J, Luan S. The calcium sensor calcineurin B-like 9 modulates abscisic acid sensitivity and biosynthesis in Arabidopsis. Plant Cell 2004; 16: 1912–1924 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Zhu SY, Yu XC, Wang XJ, Zhao R, Li Y, Fan RC, Shang Y, Du SY, Wang XF, Wu FQ, Xu YH, Zhang XY, Zhang DP. Two calcium-dependent protein kinases, CPK4 and CPK11, regulate abscisic acid signal transduction in Arabidopsis. Plant Cell 2007; 19: 3019–3036 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Hrabak EM, Chan CW, Gribskov M, Harper JF, Choi JH, Halford N, Kudla J, Luan S, Nimmo HG, Sussman MR, Thomas M, Walker-Simmons K, Zhu JK, Harmon AC. The Arabidopsis CDPK-SnRK superfamily of protein kinases. Plant Physiol 2003; 132: 666–680 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Leclercq J, Ranty B, Sanchez-Ballesta MT, Li Z, Jones B, Jauneau A, Pech JC, Latche A, Ranjeva R, Bouzayen M. Molecular and biochemical characterization of LeCRK1, a ripening-associated tomato CDPK-related kinase. J Exp Bot 2005; 56: 25–35 [PubMed], [Google Scholar]
- Gargantini PR, Gonzalez-Rizzo S, Chinchilla D, Raices M, Giammaria V, Ulloa RM, Frugier F, Crespi MD. A CDPK isoform participates in the regulation of nodule number in Medicago truncatula. Plant J 2006; 48: 843–856 [Crossref], [PubMed], [Google Scholar]
- Raices M, Gargantini PR, Chinchilla D, Crespi M, Tellez-Inon MT, Ulloa RM. Regulation of CDPK isoforms during tuber development. Plant Mol Biol 2003; 52: 1011–1024 [Crossref], [PubMed], [Google Scholar]
- Dammann C, Ichida A, Hong B, Romanowsky SM, Hrabak EM, Harmon AC, Pickard BG, Harper JF. Subcellular targeting of nine calcium-dependent protein kinase isoforms from Arabidopsis. Plant Physiol 2003; 132: 1840–1848 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Lu SX, Hrabak EM. An Arabidopsis calcium-dependent protein kinase is associated with the endoplasmic reticulum. Plant Physiol 2002; 128: 1008–1021 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Nimchuk Z, Marois E, Kjemtrup S, Leister RT, Katagiri F, Dangl JL. Eukaryotic fatty acylation drives plasma membrane targeting and enhances function of several type III effector proteins from Pseudomonas syringae. Cell 2000; 101: 353–363 [Crossref], [PubMed], [Google Scholar]
- Thieme F, Szczesny R, Urban A, Kirchner O, Hause G, Bonas U. New type III effectors from Xanthomonas campestris pv. vesicatoria trigger plant reactions dependent on a conserved N-myristoylation motif. Mol Plant Microbe Interact 2007; 20: 1250–1261 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Kim HS, Desveaux D, Singer AU, Patel P, Sondek J, Dangl JL. The Pseudomonas syringae effector AvrRpt2 cleaves its C-terminally acylated target, RIN4, from Arabidopsis membranes to block RPM1 activation. Proc Natl Acad Sci USA 2005; 102: 6496–6501 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Axtell MJ, Chisholm ST, Dahlbeck D, Staskawicz BJ. Genetic and molecular evidence that the Pseudomonas syringae type III effector protein AvrRpt2 is a cysteine protease. Mol Microbiol 2003; 49: 1537–1546 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Axtell MJ, Staskawicz BJ. Initiation of RPS2-specified disease resistance in Arabidopsis is coupled to the AvrRpt2-directed elimination of RIN4. Cell 2003; 112: 369–377 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Mackey D, Belkhadir Y, Alonso JM, Ecker JR, Dangl JL. Arabidopsis RIN4 is a target of the type III virulence effector AvrRpt2 and modulates RPS2-mediated resistance. Cell 2003; 112: 379–389 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Fondong VN, Reddy RV, Lu C, Hankoua B, Felton C, Czymmek K, Achenjang F. The consensus N-myristoylation motif of a geminivirus AC4 protein is required for membrane binding and pathogenicity. Mol Plant Microbe Interact 2007; 20: 380–391 [Crossref], [PubMed], [Google Scholar]
- Caron JM, Vega LR, Fleming J, Bishop R, Solomon F. Single site alpha-tubulin mutation affects astral microtubules and nuclear positioning during anaphase in Saccharomyces cerevisiae: possible role for palmitoylation of alpha-tubulin. Mol Biol Cell 2001; 12: 2672–2687 [Crossref], [PubMed], [Google Scholar]
- Wolff J, Zambito AM, Britto PJ, Knipling L. Autopalmitoylation of tubulin. Protein Sci 2000; 9: 1357–1364 [Crossref], [PubMed], [Google Scholar]
- Zambito AM, Wolff J. Palmitoylation of tubulin. Biochem Biophys Res Commun 1997; 239: 650–654 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Mattoo AK, Edelman M. Intramembrane translocation and posttranslational palmitoylation of the chloroplast 32-kDa herbicide-binding protein. Proc Natl Acad Sci USA 1987; 84: 1497–1501 [Crossref], [PubMed], [Google Scholar]
- Joyard J, Stumpf PK. Synthesis of long-chain Acyl-CoA in chloroplast envelope membranes. Plant Physiol 1981; 67: 250–256 [Crossref], [PubMed], [Google Scholar]
- Gauthier-Campbell C, Bredt DS, Murphy TH, El-Husseini AED. Regulation of dendritic branching and filopodia formation in hippocampal neurons by specific acylated protein motifs. Molec Biol Cell 2004; 15: 2205–2217 [Crossref], [PubMed], [Google Scholar]
- Raucher D, Sheetz MP. Cell spreading and lamellipodial extension rate is regulated by membrane tension. J Cell Biol 2000; 148: 127–136 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Peskin CS, Odell GM, Oster GF. Cellular motions and yhermal fluctuations – the Brownian ratchet. Biophys J 1993; 65: 316–324 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Xu J, Scheres B. Dissection of Arabidopsis ADP-RIBOSYLATION FACTOR 1 function in epidermal cell polarity. Plant Cell 2005; 17: 525–536 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Zhou F, Xue Y, Yao X, Xu Y. CSS-Palm: palmitoylation site prediction with a clustering and scoring strategy (CSS). Bioinformatics 2006; 22: 894–896 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Xue Y, Chen H, Jin C, Sun Z, Yao X. NBA-Palm: prediction of palmitoylation site implemented in Naive Bayes algorithm. BMC Bioinformatics 2006; 7: 458 [Crossref], [PubMed], [Google Scholar]
- Webb Y, Hermida-Matsumoto L, Resh MD. Inhibition of protein palmitoylation, raft localization, and T cell signaling by 2-bromopalmitate and polyunsaturated fatty acids. J Biol Chem 2000; 275: 261–270 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Lawrence DS, Zilfou JT, Smith CD. Structure-activity studies of cerulenin analogues as protein palmitoylation inhibitors. J Med Chem 1999; 42: 4932–4941 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Patterson SI, Skene JH. Inhibition of dynamic protein palmitoylation in intact cells with tunicamycin. Methods Enzymol 1995; 250: 284–300 [Crossref], [PubMed], [Google Scholar]
- Liang X, Nazarian A, Erdjument-Bromage H, Bornmann W, Tempst P, Resh MD. Heterogeneous fatty acylation of Src family kinases with polyunsaturated fatty acids regulates raft localization and signal transduction. J Biol Chem 2001; 276: 30987–30994 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Berthiaume L, Peseckis SM, Resh MD. Synthesis and use of iodo-fatty acid analogs. Meth Enzymol 1995; 250: 454–466 [Crossref], [PubMed], [Google Scholar]
- Drisdel RC, Green WN. Labeling and quantifying sites of protein palmitoylation. Biotechniques 2004; 36: 276–285 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Kenworthy AK. Fluorescence-based methods to image palmitoylated proteins. Methods 2006; 40: 198–205 [Crossref], [PubMed], [Google Scholar]
- Goodwin JS, Kenworthy AK. Photobleaching approaches to investigate diffusional mobility and trafficking of Ras in living cells. Methods 2005; 37: 154–164 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]
- Tian T, Harding A, Inder K, Plowman S, Parton RG, Hancock JF. Plasma membrane nanoswitches generate high-fidelity Ras signal transduction. Nat Cell Biol 2007; 9: 905–914 [Crossref], [PubMed], [Web of Science ®], [Google Scholar]