Plants exposed to hyperosmotic stress undergo changes in membrane dynamics and lipid composition to maintain cellular integrity and avoid membrane leakage. Various plant species respond to hyperosmotic stress with transient increases in PtdIns(4,5)P2; however, the physiological role of such increases is unresolved. The plasma membrane represents the outermost barrier between the symplast of plant cells and its apoplastic surroundings. In the present study, the spatio-temporal dynamics of stress-induced changes in phosphoinositides were analysed in subcellular fractions of Arabidopsis leaves to delineate possible physiological roles. Unlabelled lipids were separated by TLC and quantified by gas-chromatographic detection of associated fatty acids. Transient PtdIns(4,5)P2 increases upon exposure to hyperosmotic stress were detected first in enriched plasmamembrane fractions, however, at later time points, PtdIns(4,5)P2 was increased in the endomembrane fractions of the corresponding two-phase systems. When major endomembranes were enriched from rosette leaves prior to hyperosmotic stress and during stimulation for 60 min, no stress-induced increases in the levels of PtdIns(4,5)P2 were found in fractions enriched for endoplasmic reticulum, nuclei or plastidial membranes. Instead, increased PtdIns(4,5)P2 was found in CCVs (clathrin-coated vesicles), which proliferated several-fold in mass within 60 min of hyperosmotic stress, according to the abundance of CCV-associated proteins and lipids. Monitoring the subcellular distribution of fluorescence-tagged reporters for clathrin and PtdIns(4,5)P2 during transient co-expression in onion epidermal cells indicates rapid stress-induced co-localization of clathrin with PtdIns(4,5)P2 at the plasma membrane. The results indicate that PtdIns(4,5)P2 may act in stress-induced formation of CCVs in plant cells, highlighting the evolutionary conservation of the phosphoinositide system between organismic kingdoms.

INTRODUCTION

PIs (phosphoinositides) are central regulators of physiological processes in eukaryotic cells. PtdIns(4,5)P2 has emerged as a multifunctional regulatory player in various biological model systems studied. By acting as a lipid ligand for various alternative protein partners, PtdIns(4,5)P2 can regulate the localization or activity of specific target proteins, and the ensuing effects on physiology in various eukaryotic organisms have been extensively reviewed previously [13]. Besides many other processes, vesicle trafficking in eukaryotic cells is influenced by PtdIns(4,5)P2–protein interactions, which are required for the assembly of the protein machinery for vesicle fusion or for budding [4,5], as well as by the particular biophysical properties of the inverse conical lipid, exerting curvature strain on the membrane [6,7]. Whereas the presence of PtdIns(4,5)P2 in plants has been demonstrated and changes in PtdIns(4,5)P2 levels during various stress responses have been reported [812], the particular adaptational processes regulated in plants by PtdIns(4,5)P2 remain unclear, in part because the stress-activated lipid kinase(s) producing PIs remain unidentified. Although increases in PtdIns(4,5)P2 after hyperosmotic stress have been proposed to support increases in Ins(1,4,5)P3 [9,13], correlating with increases in cytosolic [Ca2+] [12], no functional evidence has been provided to link these metabolite changes with the plant adaptational response. So far, it is not clear whether intact PtdIns(4,5)P2, Ins(1,4,5)P3 or other derived metabolites represent physiological stress signals.

The subcellular distribution of PIs in a eukaryotic cell is a key factor in determining the regulatory effects that will be exerted by the lipids [14]. PIs and the activities of their biosynthetic enzymes have been found to be associated with different subcellular structures of plant cells [3,15,16], suggesting that different regulatory functions of PIs are exerted in different compartments. Previous work performed on plants provides evidence that PtdIns(4,5)P2 and other PIs are organized into distinct lipid pools [8,9,13,17] that may be independently regulated. The subcellular location of stress-induced increases of PtdIns(4,5)P2 in plant cells may thus give important insights into what physiological processes may be controlled by PtdIns(4,5)P2.

In the present study, hyperosmotic stress was used as a well-characterized stimulus eliciting changes in PtdIns(4,5)P2 levels in plants [8,9,11,12]. The exact physiological roles of stress-induced PtdIns(4,5)P2 are unknown, however, but its functions will probably relate to stress adaptation. The most immediate threats posed by hyperosmotic stress are osmotic efflux of water and loss of turgor pressure [18], leading to changes in the cellular ultrastructure [19] and membrane leakage [20]. In order to maintain cellular integrity under hyperosmotic conditions, a number of membrane-rearrangement processes have been described in biological and artificial model systems that are thought to minimize membrane leakage [20]. A process which immediately follows osmotic water loss is bulk-flow endocytosis, by which the excess membrane area is internalized into endomembrane vesicles to reduce the overall cell-surface area and regain turgor pressure [18]. It has been suggested that PtdIns(3)P is involved in endomembrane trafficking following bulk-flow endocytosis in plant cells [21]. Although the PI system may thus be involved in plasma-membrane-to-endomembrane trafficking in plants, a role for PtdIns(4,5)P2 in related processes has so far not been demonstrated. Because it is known from other eukaryotic model systems that PtdIns(4,5)P2 takes part in vesicle fusion or vesicle-budding processes at the plasma membrane, for instance during synaptic exocytosis for neurotransmitter release and subsequent endocytotic membrane recycling events [4,5,7], it was one working hypothesis of the present study that intact PtdIns(4,5)P2 may play a role in the formation of endocytotic plasma-membrane vesicles.

In the present paper, we show that PtdIns(4,5)P2 formed as a result of hyperosmotic stress is associated first with plasma-membrane-enriched fractions of stressed Arabidopsis plants and is increased in the endomembrane fraction at later time points of stimulation. Increased PtdIns(4,5)P2 levels are shown to associate with CCVs (clathrin-coated vesicles), whereas no increase in the levels of PtdIns(4,5)P2 was found in the ER (endoplasmic reticulum), nuclei or plastidial membranes. Stress-induced co-localization of PtdIns(4,5)P2 with clathrin is visualized in onion epidermal cells transiently co-expressing fluorescent reporters for PtdIns(4,5)P2 and clathrin, suggesting a role for PtdIns(4,5)P2 in the recruitment of coat proteins to the plasma membrane during vesicle budding in plant cells.

EXPERIMENTAL

Plant growth conditions and stress treatments

All experiments were performed with Arabidopsis thaliana ecotype Columbia 0 (col-0). Plants were grown on soil under exposure to 140 μmol photons·m−2·s−1 of light in an 8 h light/16 h dark regime. Seeds for plants destined for stress treatments were surface-sterilized by incubation for 5 min at 22 °C in 75% (v/v) ethanol, followed by incubation for 20 min at 22 °C in 6% (w/v) sodium hypochorite in 0.1% Triton X-100 and washed five times in sterile distilled water. Seeds were vernalized at 4 °C for 2 days, followed by culture in sealed jars on 0.5% Murashige and Skoog medium with modified vitamins (Duchefa) containing 1% sucrose and 0.25% Gelrite (Carl Roth). After 14 days, plants were transferred into hydroponic cultures in liquid medium as described previously [22]. Hydroponic cultures were exposed to 140 μmol photons·m−2·s−1 of light in an 8 h light/16 h dark regime and continuously aerated. Plants (8–10 weeks old) were treated by adding NaCl to final concentrations of 0.4 M and 0.8 M respectively to the hydroponic medium. Rosette leaves were harvested before treatment and after various periods of stimulation, as indicated in the Results section, and immediately frozen in liquid nitrogen. Care was taken to perform experiments over the same time period within the light/dark regime, and not to cross the light/dark transition.

Enrichment of subcellular fractions

Plasma membrane and bulk endomembranes

Plant material (50–100 g) was snap-frozen in liquid nitrogen and ground to a fine powder using a mortar and pestle. The resulting powder was resuspended in ice-cold 30 mM Tris/HCl (pH 7.4) containing 200 mM sucrose, 14 mM 2-mercaptoethanol, 2 mM DTT (dithiothreitol), 3 mM EDTA, 3 mM EGTA, 1 mM PMSF, 1.5% (w/v) PVPP (polyvinylpolypyrrolidone) and 10 μg·ml−1 leupeptin and subjected to 20 strokes in a glass dounce homogenizer on ice. The resulting extract was cleared by centrifugation at 3700 g for 15 min at 4 °C. The supernatant was centrifuged at 14500 rev./min for 45 min at 4 °C (Sorvall SS34 rotor) to pellet total cellular membranes. The total membrane pellet was used for plasma-membrane preparation by aqueous two-phase partitioning on 6.3% (w/w) PEG [poly(ethylene glycol)]/dextran polymer gradients as described previously [10].

ER

Plant material (50–100 g) was macerated with a razor blade and resuspended to a final concentration of 1 g·ml−1 in ice-cold buffer A [40 mM Hepes (pH 7.5) containing 400 mM sucrose, 10 mM KCl, 3 mM MgCl2, 1 mM DTT and 0.1 mM EDTA]. The homogenate was filtered through one layer of Miracloth (Calbiochem) and cleared by centrifugation at 6000 g for 20 min at 4 °C. From the supernatant, ER membranes were enriched as described previously [23].

Nuclei

Plant material (50–100 g) was snap-frozen in liquid nitrogen and ground to a fine powder using a mortar and pestle. The resulting powder was resuspended to a final concentration of 1 g·ml−1 in ice-cold buffer B {25 mM Mes/KOH (pH 6.2), 5.5 M glycerol, 600 mM sorbitol, 10 mM 2-mercaptoethanol, 5 mM EDTA, 0.5 mM DEDTC (diethyldithiocarbamate), 0.5 mM AEBSF [4-(2-aminoethyl)benzenesulfonyl fluoride], 0.5 mM spermidine, 0.2 mM spermine, 0.05% Triton X-100, 0.1 mg·ml−1 delipidated BSA (Sigma) and 0.1 mg·ml−1 PVP-40 (polyvinylpyrrolidone-40)} and incubated for 30 s at 22 °C. The suspension was sequentially filtered through nylon meshes of decreasing pore size (250 μm-, 80 μm- and 20 μm-pore-size). The filtrate was collected and used for the enrichment of nuclei using Percoll gradients as described previously [24].

F-actin

Plant material (50–100 g) was snap-frozen in liquid nitrogen and ground to a fine powder using a mortar and pestle. The powder was resuspended at a final concentration of 1 g·ml−1 in cold 5 mM Hepes/KOH (pH 7.5) containing 10 mM magensium acetate, 2 mM EGTA, 1 mM PMSF and 0.5% (w/v) PTE (polyoxyethylene tridecyl ether) and used for the enrichment of filamentous actin as described previously [25].

Plastids

Plant material (50 g) was homogenized for 3×5 s using a polytron homogenizer (PCU-2, Type 10203500; Kinematic) in 50 ml of ice-cold 20 mM Hepes/KOH (pH 8.0) containing 0.3 M sorbitol, 5 mM MgCl2, 5 mM EGTA, 5 mM EDTA and 10 mM NaHCO3 in a 150 ml beaker. The homogenate was filtered through a double layer of Miracloth. The debris retained in the Miracloth was returned to the beaker with 20 ml of fresh homogenization buffer and the homogenization was repeated. Plastid enrichment was carried out on the filtrate as described previously [24].

CCVs

Plant material (50–100 g) was homogenized in 1 g·ml−1 of ice-cold buffer A using a polytron homogenizer (PCU-2, Type 10203500; Kinematic). The homogenate was filtered through three layers of Miracloth and was cleared by centrifugation at 8000 g for 50 min at 4 °C. CCVs were enriched as described previously [24].

The lipid phosphatase inhibitor (3S)[1,1-difluoro-3,4-bis-(oleoyloxy)butyl] phosphonate (Echelon) was included in all buffers at a final concentration of 3 μM.

Marker tests

Biochemical tests for the enrichment of particular subcellular fractions were performed as described previously: concanavalin A binding [26]; β1,3-glucan synthetase and NADH cytochrome C reductase [27]; DAPI (4′,6-diamidino-2-phenylindole)-staining [28]; and chlorophyll estimation [29].

Protein electrophoresis and immunodetection

Protein contents were determined according to the Bradford method [30] using BSA as a standard. Proteins were separated by SDS/PAGE analysis as described previously [31] using the Mini Protean 3 System (Bio-Rad). Proteins were transferred from acrylamide gels on to nitrocellulose membranes (Optitran BA-S 85, 0.45 μm; Whatman) using the Mini Protean 3 Wet-blot system (Bio-Rad). Membranes were washed and blocked as described previously [8]. Immunodetection was performed using the following antibodies: rabbit anti-PMA2 (plasma membrane ATPase 2) antibody [32], rabbit anti-actin antibody (Sigma); rabbit anti-TGA2 (basic/leucine zipper transcription factor 2) antibody (provided by Dr C. Gatz, Department of General and Developmental Plant Physiology, Georg-August-Universität Göttingen, Göttingen, Germany); rabbit anti-BiP (ER-luminal binding protein) antibody [33]; mouse anti-clathrin antibody (Sigma, [34]) and rabbit anti-[V-ATPase (vacuolar H+-ATPase)] antibody [35]. Primary antibodies were detected using appropriate secondary antibodies (Sigma) conjugated to alkaline phosphatase. Alkaline phosphatase was detected using a colour reagent (Sigma) containing Nitro Blue Tetrazolium and 5-bromo-4-chloroindol-3-yl phosphate as described previously [36].

Lipid extraction and biochemical analyses

PIs were extracted using an acidic-extraction protocol [37]. Lipids were separated by TLC on silica gel plates (Merck) using developing solvents for optimal resolution: for PIs, chloroform/methanol/ammonium hydroxide/water [57:50:4:11 (by vol.)] [38]; for PtdCho (phosphatidylcholine) and PtdEtn (phosphatidylethanolamine), acetone/toluene/water [91:30:7 (by vol.)] [39]; and for isolating PtdIns, chloroform/methylacetate/propan-2-ol/methanol/0.25% aqueous potassium chloride [25:25:25:10:9 (by vol.)] [40]. Lipids were isolated and analysed for the presence of associated fatty acids as described previously [41]. Variations in fatty-acid patterns obtained with material sampled on different days did not exceed that denoted by standard deviations. As a result of limiting material in samples representing isolated minor lipids, fatty acids of low abundance may be absent from fatty-acid patterns.

cDNA constructs

cDNA fragments for EYFP (enhanced yellow fluorescent protein) (Clontech) and RedStar [42] tags were amplified from plasmids carrying the authentic clones provided by Dr Martin Fulda (Department of Plant Biochemistry, Georg-August-Universität Göttingen, Göttingen, Germany) using the following oligonucleotide primer combinations: EYFP, 5′-GATCGCGGCCGCCCATGGTGAGCAAGGGCGAG-3′ and 5′-GATCGATATCTTACTTGTACAGCTCGTCCATG-3′; and RedStar, 5′-GATCGTCGACATGAGTGCTTCTTCTGAAGATGTC-3′ and 5′-GATCGCGGCCGCCAAGAACAAGTGGTGTCTACCTT-3′. The coding sequence for the human PLCδ1 PH domain (where PH domain is pleckstrin homology domain and PLC is phospholipase C) was amplified from plasmid DNA provided by Dr Tamas Balla [National Institute of Child Health and Human Development (NICHD), National Institutes of Health (NIH), Bethesda, MD, U.S.A.] [43] and modified to encode a seven-amino-acid linker (Gly–Gly–Ala–Gly–Ala–Ala–Gly) between the PH domain and the RedStar tag as described previously [44], using the primers 5′-GATCGCGGCCGCCGGTGGAGCTGGAGCTGCAGGAATGAGGATCTACAGGCGC-3′ and 5′-GATCGATATCTTAGATCTTGTGCAGCCCCAGCA-3′. The coding sequence for the Arabidopsis clathrin light chain was amplified from Arabidopsis leaf cDNA using the primers 5′-GATCGCGGCCGCCATGGGCTCTGCCTTTGAAGACGATTCCTTC-3′ and 5′-GATCGCGGCCGCTTAAGCAGCAGTAACTGCCTCAGT-3′. cDNA sequences encoding fluorescent tags were subcloned into pGemTeasy and subcloned into pUC18Entr [45] as a NotI- and EcoRV-digested fragment (EYFP) or as a SalI- and NotI-digested fragment (RedStar), creating the plasmids pUC18Entr-EYFP and p18Entr-RedStar respectively. Prior to ligation, the ccdb gene was eliminated from pUC18Entr by EcoRI digestion and religation. The cDNA sequence encoding clathrin was subcloned into pUC18Entr-EYFP as a NotI-digested fragment, and the DNA sequence encoding PLCδ1 PH domain was inserted into pUC18Entr-RedStar as a NotI- and EcoRV-digested fragment. Inserts in the resulting plasmids were transferred into the plasmid pCAMBIA3300 [45] using Gateway Technology (Invitrogen) according to the manufacturer's instructions.

Detection of specific transcripts by RNA–DNA hybridization

RNA was extracted from the rosette leaves of hydroponically grown plants at different time points after exposure to salt treatment as described previously [46]. cDNA probes were amplified from whole Arabidopsis cDNA using the primers 5′-GTCTTCTTCTTCTACATAAAATTG-3′ and 5′-GGAAATTATTAGCGTTGTCATTA-3′ [for HVA (high-voltage activated)] or 5′-AATGTGTACGTCTTTTGCATAAG-3′ and 5′-GTAACATCTTCTCTTATTTATATAA-3′ [for RD20 (responsive to desiccation 20)] and radiolabelled by random priming. Hybridizations were performed as described previously [13].

Particle bombardment and transient expression in onion cells

Transient gene expession in onion epidermal cells was achieved by ballistic bombardment of plant material with DNA-coated gold particles [47]. Plasmid DNA (5 μg) was precipitated on to gold particles (1.0 mm, Bio-Rad). Fresh onions (Allium cepa L) were cut and the pieces were placed on to wet filter paper. Whole pieces were bombarded using a Biolistic PDS 1000/He Biolistic Particle Delivery System (Bio-Rad) with 1350 lbf/in2 (1 lbf/in2=6.9 kPa) rupture discs under a vacuum of 10 kPa. After bombardment, samples were incubated at high humidity for 24 h at 22 °C. Onion skin epidermal-cell layers were peeled and transferred into glass slides for microscopic examination. Stress treatments were applied directly on to microscope slides by adding 0.2 M NaCl in water and drawing the solution under the cover slip.

Confocal imaging

Images were recorded using a Zeiss LSM 510 confocal microscope. EYFP was excited at 514 nm and imaged using an HFT 405/514/633 nm MBS (main beam splitter) and a 530–600 nm band-pass filter; RedStar was excited at 561 nm and imaged using an HFT 405/488/561 nm MBS and a 583–604 nm band-pass filter. In co-expression experiments, EYFP and RedStar were synchronously excited at 488 nm and 561 nm respectively and imaged using an HFT 405/488/561 nm MBS and a 518–550 nm band-pass filter and a 583–636 nm bandpass filter respectively. Images were obtained by confocal microscopy at ×1000 magnification using the Zeiss LSM510 image-aqcuisition system and software (Version 4.0; Zeiss). Fluorescence and transmitted-light images were contrast-enhanced by adjusting the brightness and γ-settings using image-processing software (Adobe Photoshop; Adobe Systems).

RESULTS

In order to delineate a physiological function for PtdIns(4,5)P2 formed upon hyperosmotic stress in Arabidopsis leaves, a direct approach of subcellular fractionation combined with lipid analysis was chosen to systematically test various subcellular compartments for increased PtdIns(4,5)P2 levels after stress application.

Characterization of organellar fractions enriched from Arabidopsis rosette leaves

In order to delineate the subcellular sites of PtdIns(4,5)P2, the first step was to enrich different subcellular fractions from Arabidopsis rosette leaves. The enriched subcellular fractions included the plasma membrane, an F-actin cytoskeletal fraction, ER, nuclei, plastids and CCVs. When the lipid patterns associated with the enriched fractions were analysed (Figure 1), neutral lipids and the plastidial galactolipids monogalactosyldiacylglycerol and digalactosyldiacylglycerol were reduced in the fractions enriched for ER, plasma membrane, F-actin or CCVs compared with total microsomal preparations. In contrast, the nuclear preparation still contained plastidial galactolipids. The F-actin-enriched fraction contained only small amounts of membrane lipids, whereas substantial amounts of PtdEtn were detected in ER-enriched fractions. The CCV-enriched fractions were pooled from five independent preparations because of limited material and may contain additional ribosomal contaminations [48]. As a prerequisite for all further investigations, fractions were analysed for cross-contamination of marker proteins using immunodetection (see Supplementary Figure S1 at http://www.BiochemJ.org/bj/415/bj4150387add.htm). For immunodetection, antibodies against PMA2 [32], actin, the nuclear transcription factor TGA2 [49], BiP [33], clathrin [34] and the microsomal V-ATPase [35] were used. When enriched fractions were individually tested using all antibodies, the resulting detection patterns indicated substantial enrichment of all fractions with only minor cross-contamination between the compartments tested. Because of limiting amounts of material, the CCV-enriched fraction was not tested for actin. The V-ATPase was detected in all fractions, as expected from its broad distribution, and this possibly indicates contamination of all fractions with endomembranes other than those specifically tested for. Enriched fractions were additionally subjected to a variety of marker tests as described previously [2629], indicating substantial enrichment from the crude extracts (see Supplementary Figure S2 at http://www.BiochemJ.org/bj/415/bj4150387add.htm). Note that the fractions used are not pure and represent only an enrichment from total microsomal pellets. Enriched subcellular fractions used in subsequent experiments were prepared by the same method as the fractions shown in Figure 1 and were additionally subjected to PI analysis.

Enrichment of subcellular fractions from Arabidopsis rosette leaves

Figure 1
Enrichment of subcellular fractions from Arabidopsis rosette leaves

Major membrane fractions were enriched from leaf material of 6 week old Arabidopsis plants, and the lipid composition of the individual enriched fractions [total microsomal preparation (MP), plasma membrane (PM), F-actin (F-A), nuclei (N), ER and CCV] was tested by TLC. Lipid extracts were subjected to chromatography and lipids were visualized by charring. Lipids were identified according to co-migration with authentic standards. DGDG, digalactosyldiacylglycerol; diPtdGro, cardiolipin; MGDG, monogalactosyldiacylglycerol; n.i. not identified; NL, neutral lipids; PtdGro, phosphatidylglycerol; PtdOH, phosphatidic acid; SG, sterol glycosides.

Figure 1
Enrichment of subcellular fractions from Arabidopsis rosette leaves

Major membrane fractions were enriched from leaf material of 6 week old Arabidopsis plants, and the lipid composition of the individual enriched fractions [total microsomal preparation (MP), plasma membrane (PM), F-actin (F-A), nuclei (N), ER and CCV] was tested by TLC. Lipid extracts were subjected to chromatography and lipids were visualized by charring. Lipids were identified according to co-migration with authentic standards. DGDG, digalactosyldiacylglycerol; diPtdGro, cardiolipin; MGDG, monogalactosyldiacylglycerol; n.i. not identified; NL, neutral lipids; PtdGro, phosphatidylglycerol; PtdOH, phosphatidic acid; SG, sterol glycosides.

Fatty acids associated with phospholipids do not differ between most organellar fractions in non-stimulated plants

In order to test whether direct detection of unlabelled lipids by the method outlined previously [41] is sufficiently sensitive to detect even minor lipid constituents in plant lipid extracts, phospholipids were isolated from the various preparations and analysed as described previously [41] (Figure 2). The results indicated that non-labelled PIs can be detected in complex plant lipid extracts. Note that the exogenous PtdIns(4,5)P2 was added to the extract in sample II at a concentration not detectable by charring (Figure 2A). The exogenous PtdIns(4,5)P2 contained arachidonic acid, a fatty acid not present endogenously in Arabidopsis, and this fatty acid was detected by subsequent GC analysis of PtdIns(4,5)P2 isolated from sample II (Figure 2B).

Unlabelled PtdIns(4,5)P2 can be detected in plant extracts

Figure 2
Unlabelled PtdIns(4,5)P2 can be detected in plant extracts

The migration of PtdIns(4,5)P2 was tested in relation to more abundant phospholipids. (A) TLC separation of leaf extracts using a developing solvent containing chloroform/methanol/water/acetic acid [10:10:3:1 (by vol.)]. Lipid extracts from non-stressed rosette leaves were used without (I) or with (II) the addition of a spiking amount of 0.5 μg of 1-stearoyl, 2-arachidonoyl PtdIns(4,5)P2 (Avanti). M, authentic PtdIns(4,5)P2 standard (Avanti). (B) PtdIns(4,5)P2 was isolated from TLC plates as described previously [41], and the fatty acids associated with the isolated lipids were analysed by GC. PtdIns(4,5)P2 from plant extract (solid line) and PtdIns(4,5)P2 from plant extract spiked with exogenous PtdIns(4,5)P2 (dotted line) are shown. Peak identities are indicated: 16:0, palmitic acid; 18:0, stearic acid; 18:1, oleic acid; 18:2, linoleic acid; 18:3, linolenic acid; 20:4, arachidonic acid. FID, flame ionization detector; i.s., pentadecanoic acid internal standard. PtdOH, phosphatidic acid; PtdSer, phosphatidylserine.

Figure 2
Unlabelled PtdIns(4,5)P2 can be detected in plant extracts

The migration of PtdIns(4,5)P2 was tested in relation to more abundant phospholipids. (A) TLC separation of leaf extracts using a developing solvent containing chloroform/methanol/water/acetic acid [10:10:3:1 (by vol.)]. Lipid extracts from non-stressed rosette leaves were used without (I) or with (II) the addition of a spiking amount of 0.5 μg of 1-stearoyl, 2-arachidonoyl PtdIns(4,5)P2 (Avanti). M, authentic PtdIns(4,5)P2 standard (Avanti). (B) PtdIns(4,5)P2 was isolated from TLC plates as described previously [41], and the fatty acids associated with the isolated lipids were analysed by GC. PtdIns(4,5)P2 from plant extract (solid line) and PtdIns(4,5)P2 from plant extract spiked with exogenous PtdIns(4,5)P2 (dotted line) are shown. Peak identities are indicated: 16:0, palmitic acid; 18:0, stearic acid; 18:1, oleic acid; 18:2, linoleic acid; 18:3, linolenic acid; 20:4, arachidonic acid. FID, flame ionization detector; i.s., pentadecanoic acid internal standard. PtdOH, phosphatidic acid; PtdSer, phosphatidylserine.

We have reported previously the association of mainly saturated and monounsaturated fatty acids with bulk PIs isolated from non-stimulated Arabidopsis rosette leaves, in contrast with a more unsaturated fatty-acid composition in structural phospholipids, such as PtdCho or PtdEtn [9]. Structural phospholipids and PIs from the enriched subcellular fractions were analysed for their associated fatty acids (Figure 3) to identify subcellular compartments containing unsaturated PtdIns along with saturated or monounsaturated PIs, indicative of candidate locations for stress-induced PI function. The comparison of the fatty-acid patterns of PIs, PtdIns4P and PtdIns(4,5)P2 with those of the structural phospholipids PtdEtn, PtdCho and PtdIns (Figure 3) confirmed an increased association of saturated or monounsaturated fatty acids with PIs in all fractions except in nuclei. In contrast, structural phospholipids contained higher proportions of unsaturated fatty acids. Although the fatty-acid patterns for a given lipid were similar overall between enriched subcellular fractions, the fatty-acid patterns observed for phospholipids isolated from nuclear extracts differed from the other extracts in containing an unusual combination of palmitic and oleic acids in structural phospholipids and PtdIns4P and an increased proportion of linoleic acid associated with PtdIns(4,5)P2. Although the distinct fatty-acid pattern for nuclear lipids supports successful enrichment of nuclei (cf. Figure 1), the physiological relevance of the fatty-acid composition of nuclear lipids in plants is not clear. Though some recent reports from the mammalian field indicate distinct roles for nuclear PIs [50,51], in the course of the present study no further experiments were performed to elucidate the particulars of nuclear phospholipids. With the noted exception, the results indicate that PIs associated with major subcellular fractions do not exhibit characteristic fatty-acid patterns that could serve as a means for easy functional distinction and demonstrate that precursors of PtdIns(4,5)P2 production can be found in all of the compartments tested.

Fatty-acid composition of phospholipids in enriched subcellular fractions

Figure 3
Fatty-acid composition of phospholipids in enriched subcellular fractions

Lipids were extracted from enriched subcellular fractions, isolated, and their associated fatty acids determined. The fractions analysed were plasma membrane (PM)-, F-actin (F-A)-, nuclei (N)-, ER- and CCV-enriched fractions. The fatty acids associated with different phospholipids isolated from the individual fractions are indicated, with the identity of the fatty acids indicated along the x-axis. Results are stated as the mol% of total fatty acids and are means±S.D. (n=3–7). 16:0, palmitic acid; 18:0, stearic acid; 18:1, oleic acid; 18:2, linoleic acid; 18:3, linolenic acid.

Figure 3
Fatty-acid composition of phospholipids in enriched subcellular fractions

Lipids were extracted from enriched subcellular fractions, isolated, and their associated fatty acids determined. The fractions analysed were plasma membrane (PM)-, F-actin (F-A)-, nuclei (N)-, ER- and CCV-enriched fractions. The fatty acids associated with different phospholipids isolated from the individual fractions are indicated, with the identity of the fatty acids indicated along the x-axis. Results are stated as the mol% of total fatty acids and are means±S.D. (n=3–7). 16:0, palmitic acid; 18:0, stearic acid; 18:1, oleic acid; 18:2, linoleic acid; 18:3, linolenic acid.

Transient increases of PtdIns(4,5)P2 in the plasma membrane are followed by increased PtdIns(4,5)P2 levels in endomembranes

Because the analysis of different subcellular fractions did not suggest a particular compartment as the site for stress-induced PtdIns(4,5)P2 formation, subcellular fractions enriched from plants subjected to hyperosmotic stress were systematically tested for increases in PtdIns(4,5)P2. As the plasma membrane represents the immediate barrier between the symplast and its apoplastic surroundings, the first fraction tested was that enriched for plasma membranes (Figure 4). A transient increase in PtdIns(4,5)P2 was evident after 15 min of stress application, which was followed by a decrease in plasma membrane PtdIns(4,5)P2 at later time points. Plasma membrane enrichment by aqueous two-phase partitioning yields a corresponding phase containing the residual endomembranes for each sample. PtdIns(4,5)P2 levels in these residual endomembrane fractions indicate a gradual increase in PtdIns(4,5)P2 in endomembranes at time points beyond 15 min of stimulation (Figure 4). Although plasma membrane PtdIns4P showed a similar dynamic pattern to plasma membrane PtdIns(4,5)P2, PtdIns4P decreased first in endomembranes and showed increased levels only after 60 min of stimulation (Figure 4); PtdIns levels increased gradually with stimulation in both the plasma membrane and in endomembranes (Figure 4).The results indicate that hyperosmotic stress-induced PtdIns(4,5)P2 is generated first in the plasma membrane and can be found in endomembranes at later time points.

Stress-induced dynamic changes in PIs in plasma membranes

Figure 4
Stress-induced dynamic changes in PIs in plasma membranes

Plants grown in hydroponic culture were subjected to hyperosmotic stress and leaf material was collected after stimulation for different time periods. Lipids were extracted from enriched plasma membrane (PM) and endomembrane (EM) fractions, PIs were isolated and their associated fatty acids determined. The total amount of fatty acids associated with individual lipids is shown, with the bar segments indicating the contribution of individual fatty acids: white segment, palmitic acid (16:0); diamond segment, stearic acid (18:0); grey segment, oleic acid [18:1Δ9 (18:1)]; striped segment; linoleic acid [18:2Δ9,12 (18:2)]; black segment; linolenic acid [18:3Δ9,12,15 (18:3)]. Results are stated as as pmol·g−1 of fresh weight and are means±S.D. (n=3).

Figure 4
Stress-induced dynamic changes in PIs in plasma membranes

Plants grown in hydroponic culture were subjected to hyperosmotic stress and leaf material was collected after stimulation for different time periods. Lipids were extracted from enriched plasma membrane (PM) and endomembrane (EM) fractions, PIs were isolated and their associated fatty acids determined. The total amount of fatty acids associated with individual lipids is shown, with the bar segments indicating the contribution of individual fatty acids: white segment, palmitic acid (16:0); diamond segment, stearic acid (18:0); grey segment, oleic acid [18:1Δ9 (18:1)]; striped segment; linoleic acid [18:2Δ9,12 (18:2)]; black segment; linolenic acid [18:3Δ9,12,15 (18:3)]. Results are stated as as pmol·g−1 of fresh weight and are means±S.D. (n=3).

Stress-induced PtdIns(4,5)P2 increases are excluded from ER, nuclei or plastidial membranes

To define the endomembrane location at which PtdIns(4,5)P2 is increased at later time points, subcellular fractions enriched for the major endomembrane systems of leaf tissue were prepared at different time points of stimulation (namely for ER, nuclei or plastids) and the fractions were analysed for the levels of associated PIs, including PtdIns(4,5)P2 (Figure 5). In the ER, nuclei or plastids, detectable levels of PtdIns(4,5)P2 were present (Figure 5); however, no increase in the levels determined was observed to be correlated with the application of stress. PI levels associated with plastids were substantially lower than those detected in the ER or nuclei, indicating little cross-contamination of samples. Overall, the dynamics of the lipids investigated indicate that the three major endomembrane systems tested did not contribute to stress-induced increases in cellular PtdIns(4,5)P2 levels.

Stress-induced dynamic changes in PIs in different endomembranes

Figure 5
Stress-induced dynamic changes in PIs in different endomembranes

Plants grown in hydroponic culture were subjected to hyperosmotic stress and leaf material was collected after stimulation for different time periods. Lipids were extracted from fractions enriched for ER, nuclei, or plastids, as indicated, and their associated fatty acids were determined. Bars represent the total amount of fatty acids associated with individual lipids, with the bar segments indicating the contribution of individual fatty acids: white segment, palmitic acid (16:0); diamond segment, stearic acid (18:0); grey segment, oleic acid [18:1Δ9 (18:1)]; striped segment, linoleic acid [18:2Δ9,12 (18:2)]; black segment, linolenic acid [18:3Δ9,12,15 (18:3)]. Results are stated as as pmol·g−1 of fresh weight and are means±S.D (n=3–5).

Figure 5
Stress-induced dynamic changes in PIs in different endomembranes

Plants grown in hydroponic culture were subjected to hyperosmotic stress and leaf material was collected after stimulation for different time periods. Lipids were extracted from fractions enriched for ER, nuclei, or plastids, as indicated, and their associated fatty acids were determined. Bars represent the total amount of fatty acids associated with individual lipids, with the bar segments indicating the contribution of individual fatty acids: white segment, palmitic acid (16:0); diamond segment, stearic acid (18:0); grey segment, oleic acid [18:1Δ9 (18:1)]; striped segment, linoleic acid [18:2Δ9,12 (18:2)]; black segment, linolenic acid [18:3Δ9,12,15 (18:3)]. Results are stated as as pmol·g−1 of fresh weight and are means±S.D (n=3–5).

Decreasing PI levels in endomembrane fractions are not related to irreversible tissue damage

The failure to detect stress-induced increases in PtdIns(4,5)P2 in the major endomembranes despite extensive experimental efforts raised the question of whether or not the plants were irreversibly damaged by the severe hyperosmotic stress applied. In order to rule out that the decreasing lipid levels in endomembrane fractions observed were a consequence of an irreversible loss of structural integrity, plants were subjected to hyperosmotic stress and tested for increases in stress-inducible RNA transcript levels over a period of 24 h by Northern blot experiments (Figure 6). The stress-inducible transcripts for HVA22 [52] and RD20 [53] showed substantial increases after 4–8 h and 1–24 h respectively, indicating that the induction of stress-inducible transcripts occurred as expected for non-damaged plants. Irreversible damage can thus be ruled out as a cause of the lipid patterns observed. Note that lipid patterns are reported for a period of only 1 h, whereas changes in transcript levels and plant survival were observed for up to 24 h.

Unimpaired transcript induction in plants exposed to salt stress

Figure 6
Unimpaired transcript induction in plants exposed to salt stress

Plants grown in hydroponic culture were subjected to hyperosmotic stress, leaf material was collected after stimulation for different time periods and transcript levels for stress-inducible genes were determined by Northern blot analysis. The ACT8 (actin) transcript was tested as a control. Results are from a representative experiment; the experiment was repeated twice with similar results.

Figure 6
Unimpaired transcript induction in plants exposed to salt stress

Plants grown in hydroponic culture were subjected to hyperosmotic stress, leaf material was collected after stimulation for different time periods and transcript levels for stress-inducible genes were determined by Northern blot analysis. The ACT8 (actin) transcript was tested as a control. Results are from a representative experiment; the experiment was repeated twice with similar results.

Hypertonic stress induces increased formation of CCVs in Arabidopsis leaves

Because the investigation of major endomembrane compartments (Figure 5) had not revealed the location of the stress-induced increase in PtdIns(4,5)P2, minor endomembrane compartments were also considered. A minor endomembrane fraction with relevance to the physiological adaptation to hyperosmotic stress are CCVs. CCVs were isolated from Arabidopsis leaves prior to stimulation and after 60 min of exposure to hyperosmotic stress (Figure 7). CCV-associated proteins from several preparations representing 30 g of fresh leaf weight were pooled and subjected to SDS/PAGE (Figure 6A). In-gel quantification of protein-staining intensities indicates a stress-induced >2-fold increase in CCV-associated proteins within 60 min of stimulation (Figure 7A). Phospholipids were extracted from the same material and quantified (Figure 7B). The raised levels of the structural phospholipids PtdEtn and PtdCho also indicate an increase in CCV mass within 60 min of stimulation, supporting a role for CCV formation in plant-stress adaptation to hypertonic conditions.

Increased formation of CCVs with hyperosmotic stress

Figure 7
Increased formation of CCVs with hyperosmotic stress

Plants grown in hydroponic culture were subjected to hyperosmotic stress, leaf material was collected after stimulation for different time periods and CCVs were enriched. (A) Protein extracts from 30 g of leaf tissue were separated by SDS/PAGE, and proteins stained with Coomassie Brilliant Blue (left-hand side panel). The protein patterns shown are from a representative experiment. Molecular masses are shown on the left-hand side (in kDa). The overall staining intensity of CCV-associated proteins from non-stimulated plants and from plants stimulated for 60 min was determined as indicated (right-hand side panel). The experiment was repeated twice with similar results; results are means±S.D. (n=3). (B) Lipids were extracted from CCV-enriched fractions and their associated fatty acids were determined. Bars represent the total amount of fatty acids associated with individual lipids, with the bar segments indicating the contribution of individual fatty acids: white segment, palmitic acid (16:0); diamond segment, stearic acid (18:0); grey segment, oleic acid [18:1Δ9 (18:1)]; striped segment; linoleic acid [18:2Δ9,12 (18:2)]; black segment; linolenic acid [18:3Δ9,12,15(18:3)]. Results are stated as as pmol·g−1 of fresh weight and are means±S.D. (n=3).

Figure 7
Increased formation of CCVs with hyperosmotic stress

Plants grown in hydroponic culture were subjected to hyperosmotic stress, leaf material was collected after stimulation for different time periods and CCVs were enriched. (A) Protein extracts from 30 g of leaf tissue were separated by SDS/PAGE, and proteins stained with Coomassie Brilliant Blue (left-hand side panel). The protein patterns shown are from a representative experiment. Molecular masses are shown on the left-hand side (in kDa). The overall staining intensity of CCV-associated proteins from non-stimulated plants and from plants stimulated for 60 min was determined as indicated (right-hand side panel). The experiment was repeated twice with similar results; results are means±S.D. (n=3). (B) Lipids were extracted from CCV-enriched fractions and their associated fatty acids were determined. Bars represent the total amount of fatty acids associated with individual lipids, with the bar segments indicating the contribution of individual fatty acids: white segment, palmitic acid (16:0); diamond segment, stearic acid (18:0); grey segment, oleic acid [18:1Δ9 (18:1)]; striped segment; linoleic acid [18:2Δ9,12 (18:2)]; black segment; linolenic acid [18:3Δ9,12,15(18:3)]. Results are stated as as pmol·g−1 of fresh weight and are means±S.D. (n=3).

Association of stress-induced PtdIns(4,5)P2 with enriched CCVs

CCVs enriched from plants exposed to hypertonic stress for different time periods were analysed for PI content (Figure 8). Within 30 min of hyperosmotic stress, the levels of CCV-associated PtdIns increased several-fold, and, after 60 min, dramatic increases were also observed for PtdIns4P and PtdIns(4,5)P2 levels. Although the stress-induced CCV-associated PtdIns and PtdIns4P contained a large proportion of the unsaturated fatty acids linoleic and linolenic acids, the increased PtdIns(4,5)P2 was increased in stearic acid and oleic acid in addition to containing an increased proportion of linolenic acid (Figure 8A). The results identify CCVs as one of the endomembrane compartments harbouring the stress-induced increase in PIs and PtdIns(4,5)P2. The mass ratio of PtdIns(4,5)P2/PtdCho was calculated (Figure 8B) and indicates that increased PtdIns(4,5)P2 levels in CCVs were not merely a reflection of increased CCV mass (cf. Figure 7), but rather were a true increase in the proportion of CCV-associated PtdIns(4,5)P2.

Stress-induced increases in PtdIns(4,5)P2 associated with CCVs

Figure 8
Stress-induced increases in PtdIns(4,5)P2 associated with CCVs

Plants grown in hydroponic culture were subjected to hyperosmotic stress and leaf material was collected after stimulation for different time periods. (A) Lipids were extracted from fractions enriched for CCVs and their associated fatty acids were determined. Bars represent the total amount of fatty acids associated with individual lipids, and the bar segments indicate the contribution of individual fatty acids: white segment, palmitic acid (16:0); diamond segment, stearic acid (18:0); grey segment, oleic acid [18:1Δ9 (18:1)]; striped segment; linoleic acid [18:2Δ9,12 (18:2)]; black segment; linolenic acid [18:3Δ9,12,15(18:3)]. Results are stated as as pmol·g−1 of fresh weight and are means±S.D. (n=4). (B) The mass ratios of PtdIns(4,5)P2/PtdCho for each time point were calculated based on the results in Figure 7(B) and Figure 8(A). An increased ratio indicates an increased association of PtdIns(4,5)P2 with CCVs.

Figure 8
Stress-induced increases in PtdIns(4,5)P2 associated with CCVs

Plants grown in hydroponic culture were subjected to hyperosmotic stress and leaf material was collected after stimulation for different time periods. (A) Lipids were extracted from fractions enriched for CCVs and their associated fatty acids were determined. Bars represent the total amount of fatty acids associated with individual lipids, and the bar segments indicate the contribution of individual fatty acids: white segment, palmitic acid (16:0); diamond segment, stearic acid (18:0); grey segment, oleic acid [18:1Δ9 (18:1)]; striped segment; linoleic acid [18:2Δ9,12 (18:2)]; black segment; linolenic acid [18:3Δ9,12,15(18:3)]. Results are stated as as pmol·g−1 of fresh weight and are means±S.D. (n=4). (B) The mass ratios of PtdIns(4,5)P2/PtdCho for each time point were calculated based on the results in Figure 7(B) and Figure 8(A). An increased ratio indicates an increased association of PtdIns(4,5)P2 with CCVs.

Stress-induced co-localization of PtdIns(4,5)P2 with clathrin at the plasma membrane

In order to verify the biochemical findings of CCV-associated PtdIns(4,5)P2 by an independent method, fluorescence-tagged markers for clathrin and for PtdIns(4,5)P2 were transiently co-expressed in onion epidermal cells, the cells were subjected to hyperosmotic stress, and the fluorescence distribution over time was monitored by confocal microscopy (Figure 9). The clathrin reporter construct encoded a fusion protein of clathrin with a C-terminally attached EYFP tag; the PtdIns(4,5)P2 reporter construct encoded the PH domain of human PLCδ1, which specifically binds to PtdIns(4,5)P2, fused to a C-terminal RedStar tag [42]. Prior to stimulation, the clathrin reporter was distributed evenly throughout the cytosol, whereas the PtdIns(4,5)P2 reporter was located at the plasma membrane (Figure 9). When cells were subjected to hyperosmotic stress, within 2 min the clathrin reporter relocalized to the plasma membrane with a distribution now identical with that of the PtdIns(4,5)P2 reporter. As hypertonic plasmolysis set in at later time points and endocytotic vesicles are too small to be reliably identified in cells undergoing dramatic structural reorganization, no time points later than 2 min were evaluated with confidence. Structural rearrangements of the plasma membrane were seen as early as 60 s after stress application, evident by the rough appearance and multiple membrane invaginations (Figure 10). The observed pattern of fluorescence distribution indicates rapid stress-induced relocalization of clathrin to sites containing PtdIns(4,5)P2, thus corroborating the biochemical results.

Stress-induced co-localization of PtdIns(4,5)P2 with clathrin in onion epidermal cells

Figure 9
Stress-induced co-localization of PtdIns(4,5)P2 with clathrin in onion epidermal cells

Onion epidermal cells were transiently co-transformed with EYFP-tagged clathrin and with a RedStar-tagged reporter for PtdIns(4,5)P2 (RedStar-PLCδ1-PH). The fluorescence distribution of bright field, RedStar (red) and EYFP (green) was synchronously and continuously monitored by confocal microscopy in cells subjected to hyperosmotic stress. The yellow colour in the bottom panels (merge) indicates co-localization. Time points over the first 2 min of stimulation are shown. (A) Bright field; (B) RedStar–PLCδ1-PH; (C) EYFP–clathrin; (D) merged image. Images shown are from a representative experiment; the transformation experiment was repeated twice with similar results. C, cytosol; PM, plasma membrane; T, tonoplast; V, vacuole. Scale bar, 20 μm.

Figure 9
Stress-induced co-localization of PtdIns(4,5)P2 with clathrin in onion epidermal cells

Onion epidermal cells were transiently co-transformed with EYFP-tagged clathrin and with a RedStar-tagged reporter for PtdIns(4,5)P2 (RedStar-PLCδ1-PH). The fluorescence distribution of bright field, RedStar (red) and EYFP (green) was synchronously and continuously monitored by confocal microscopy in cells subjected to hyperosmotic stress. The yellow colour in the bottom panels (merge) indicates co-localization. Time points over the first 2 min of stimulation are shown. (A) Bright field; (B) RedStar–PLCδ1-PH; (C) EYFP–clathrin; (D) merged image. Images shown are from a representative experiment; the transformation experiment was repeated twice with similar results. C, cytosol; PM, plasma membrane; T, tonoplast; V, vacuole. Scale bar, 20 μm.

Stress-induced changes in plasma-membrane appearance

Figure 10
Stress-induced changes in plasma-membrane appearance

The fluorescence distribution of bright field, EYFP (green) and RedStar (red) was synchronously and continuously monitored by confocal microscopy in onion epidermal cells transiently co-transformed with EYFP-tagged clathrin and with a RedStar-tagged reporter for PtdIns(4,5)P2 (RedStar-PLCδ1-PH) which had been subjected to hyperosmotic stress. Plasma-membrane areas indicated in Figure 9 (boxes in bright field at 0 and 60 s time points) were scanned at a higher magnification to resolve the distribution of PtdIns(4,5)P2 and clathrin in relation to rearrangements of the plasma membrane during exposure to stress. The time points chosen reflect the non-stimulated situation (0 s) and the situation 60 s after stimulation. Images shown are from a representative experiment; the transformation experiment was repeated twice with similar results. Scale bar, 5 μm.

Figure 10
Stress-induced changes in plasma-membrane appearance

The fluorescence distribution of bright field, EYFP (green) and RedStar (red) was synchronously and continuously monitored by confocal microscopy in onion epidermal cells transiently co-transformed with EYFP-tagged clathrin and with a RedStar-tagged reporter for PtdIns(4,5)P2 (RedStar-PLCδ1-PH) which had been subjected to hyperosmotic stress. Plasma-membrane areas indicated in Figure 9 (boxes in bright field at 0 and 60 s time points) were scanned at a higher magnification to resolve the distribution of PtdIns(4,5)P2 and clathrin in relation to rearrangements of the plasma membrane during exposure to stress. The time points chosen reflect the non-stimulated situation (0 s) and the situation 60 s after stimulation. Images shown are from a representative experiment; the transformation experiment was repeated twice with similar results. Scale bar, 5 μm.

DISCUSSION

Direct detection of unlabelled PIs was employed to determine the subcellular location of transient increases in PtdIns(4,5)P2 during hyperosmotic stress in Arabidopsis rosette leaves. The direct detection of PIs provides substantial advantages over approaches using radiolabelling because it is possible to quantify total lipid mass changes in comparison with changes in relative radiolabel incorporation, which is additionally subject to a number of factors that are difficult to control, such as uptake and the metabolic state of the labelled tissues. It was the rationale of the present study that the subcellular sites of changes in PI as a result of stress may suggest physiological functions of the increased lipid levels.

Classical techniques for subcellular fractionation were used to enrich for plasma membranes and various endomembrane fractions, and it is clear that the quality of the analyses shown depends on the quality of the enrichment performed. The lipid patterns characteristic for individual subcellular fractions as well as the immunodetection of marker proteins and enzyme tests indicate substantial enrichment of the respective fractions (Figure 1 and Supplementary Figures S1 and S2). It must be noted that the fractions obtained by the methods used will no doubt contain elements of subcellular compartments other than those enriched for. Importantly, however, the immunodetection of marker proteins indicates little cross-contamination between the enriched fractions tested (Supplementary Figure S1), which represent some of the major pools reported to contain PIs in eukaryotic cells. Although contamination by other compartments cannot be ruled out, the effects of mitochondrial membranes on PI patterns, for instance, will be small.

When phospholipids were extracted from enriched subcellular fractions and analysed for the quantity and nature of their associated fatty acids, structural phospholipids and PtdIns contained high proportions of unsaturated fatty acids, in particular linoleic and linolenic acids (Figure 3), as was reported previously for extracts from whole cells [9]. In contrast the PIs PtdIns4P and PtdIns(4,5)P2 isolated from these enriched subcellular fractions contained close to no unsaturated fatty acids, as was also reported previously for whole-cell extracts [9]. These results indicate that there may be no organelle-specific ‘fatty-acid signature’ of phospholipids or PIs. An exception is presented by nuclear extracts, which showed marked differences in the fatty-acid compositions of all phospholipids in comparison with extracts from other enriched fractions (Figure 3). Although lipid patterns (Figure 1) were ambiguous with respect to the purity of the nuclear extracts, the distinct fatty-acid pattern of the nuclear lipids (Figure 3) indicates a substantial degree of enrichment. The distinct fatty-acid patterns of nuclear lipids, especially those of PIs, may prove relevant in the future, possibly in support of recent reports of independent functions for nuclear PIs [50,51]. Nuclear lipid patterns were, however, not a focus of the present study, and because no stress-induced increases in PtdIns(4,5)P2 were associated with nuclei (Figure 5), these results shall not be further discussed at this point. Overall, the fatty-acid patterns for various phospholipids and PIs from enriched subcellular fractions of non-stimulated Arabidopsis leaves (Figure 3) confirmed that various subcellular compartments contain the elements required for PtdIns(4,5)P2 production and thus the question of the subcellular sites of stress-induced PtdIns(4,5)P2 formation remains open.

By using the direct detection of unlabelled lipids, a transient increase in PtdIns4P and PtdIns(4,5)P2 could be located subsequently in enriched plasma membranes (Figure 4). Early increases in plasma membrane PIs with hypertonic stress may be related to the fact that the plasma membrane is the primary site of contact of the cell with its apoplastic surroundings. The transient nature of the plasma membrane PtdIns(4,5)P2 increase and the subsequent increase observed in endomembranes (Figure 4) may reflect lipid relocation either underlying or as a consequence of dynamic membrane rearrangements to maintain cellular integrity during conditions of hyperosmotic stress. From the results presented, it remains unclear whether PIs formed at the plasma membrane are internalized into endomembranes or whether the endomembranes hold stress-activated enzymes for PI biosynthesis. Although to our knowledge no stress-activated isoforms of PI kinases have been reported to date, it appears relevant in this context that the Arabidopsis PtdIns4P 5-kinase isoform 1, when heterologously expressed in tobacco BY-2 cells, relocated from the plasma membrane to endomembranes when the cells were exposed to hyperosmotic stress [54].

Although previous reports have demonstrated the association of PIs with ER [55,56], nuclei [50,51] and plastid-envelope membranes [57,58], the systematic analysis of the different endomembrane fractions revealed that the increased PtdIns4P and PtdIns(4,5)P2 levels observed in total endomembranes were not associated with these locations (Figure 5). A stress-induced increase in PtdIns (Figure 5) was found only in fractions enriched for ER, an observation consistent with the ER-associated PtdIns biosynthesis reported previously [5961]. The present results suggest that the PI pools present in the ER, nuclei or plastids are not sensitive to hypertonic stress. Increases of PtdIns4P and PtdIns(4,5)P2 in reponse to stress were positively identified in fractions enriched in CCVs (Figure 8), which are formed at the plasma membrane in a stress-inducible manner (Figure 7). The fatty-acid composition of the CCV-associated PIs was highly unsaturated (Figure 8) compared with that of PIs that were constitutively present in Arabidopsis (Figure 3). This observation appears relevant in the context of a recent report suggesting that PtdIns(4,5)P2 containing long-chain polyunsaturated fatty acids may be required for synaptic vesicle endocytosis in Caenorhabditis elegans [62]. Although in the present study a protocol for vesicle enrichment was chosen that yielded a vesicle fraction defined by an easily detectable marker (clathrin), vesicle association of PtdIns(4,5)P2 may not be limited to CCVs and the lipid may also associate with other types of vesicles. Although the direct biochemical detection of PtdIns4P and PtdIns(4,5)P2 associated with CCVs has not, to our knowledge, been reported before, it has been demonstrated for mammalian cells that clathrin, the major coat protein of CCVs, is recruited to CCVs in a PtdIns(4,5)P2-dependent fashion [63]. The observation that PtdIns(4,5)P2 increases first at the plasma membrane and subsequently in CCVs (Figures 4 and 8) is corroborated by microscopy (Figure 9), indicating a rapid stress-induced relocation of clathrin from the cytosol to the plasma membrane. In a previous study, it was shown that the PLCδ1 PH domain reporter for PtdIns(4,5)P2 is found in the cytosol of non-stimulated cells and will relocate to the plasma membrane upon hyperosmotic stress [64]. The present observation of plasma-membrane association of the reporter may indicate that our ‘non-stimulated’ cells were already in a stimulated state, possibly as a result of handling and partial desiccation during microscopic procedures. As the clathrin distribution was, however, not affected by this inevitable pretreatment, the interpretation of the results presented in the present study is not impaired. The rough appearance of the plasma membrane after co-localization of clathrin and PtdIns(4,5)P2 (Figure 10) may indicate vesicle budding initiating membrane internalization and bulk-flow endocytosis. Though the dynamic distribution of fluorescent reporters for clathrin and PtdIns(4,5)P2 were recorded over a longer period than 2 min, rapid structural changes in membrane organization and plasmolysis make such images difficult to interpret. Occasional patterns of co-localized clathrin and the PtdIns(4,5)P2 reporter in small punctate cytosolic particles may suggest internalized vesicles (results not shown), which have, however, not been imaged with reliable reproducibility. It must be noted that the time frame of lipid changes observed by biochemical analyses on whole plants grown in hydroponic culture cannot be compared directly with the dynamics of clathrin relocalization in onion epidermal cells exposed directly to hypertonic medium.

In summary, the direct biochemical detection of unlabelled phospholipids in the subcellular fractions of salt-stressed Arabidopsis leaves indicates that stress-induced increases in PtdIns4P and PtdIns(4,5)P2 occur first at the plasma membrane and subsequently in CCVs. CCVs increase during hyperosmotic stress, possibly as one mechanism supporting bulk-flow endocytosis of the membrane area during plasmolytic membrane rearrangement. Confocal imaging of fluorescent reporters transiently expressed in onion epidermal cells confirms the stress-induced co-localization of PtdIns(4,5)P2 with clathrin, the major coat protein of CCVs. The identification of the subcellular site of stress-induced PtdIns(4,5)P2 increases in plant cells will serve as a basis for future investigations into the roles of PtdIns(4,5)P2 in the formation of CCVs and in plant endocytosis.

We thank Dr Tamas Balla [National Institute of Child Health and Human Development (NICHD), National Institutes of Health (NIH), Bethesda, MD, U.S.A.] for providing the HsPLCδ1 PH domain construct. We also thank the following for gifts of antibodies: anti-PMA2 antibody [Dr Marc Boutry (Institute of Life Sciences, Universite Catholique de Louvain, Louvain-la-Neuve, Belgium)]; anti-TGA2 antibody [Dr Christiane Gatz (Department of General and Developmental Plant Physiology, Georg-August-Universität Göttingen, Göttingen, Germany)]; anti-BiP antibody [Dr Alessando Vitale (Instituto Biosintesi Vegetali, Consiglio Nazionale delle Ricerche, Milan, Italy)]; and anti-(V-ATPase) antibody [Dr Rafael Ratajczak (Institute of Botany, Technical University of Darmstadt, Darmstadt, Germany)]. We thank Dr Ivo Feussner (Department of Plant Biochemistry, Georg-August-Universität Göttingen, Göttingen, Germany) and Dr Giselbert Hinz (Heidelberg Institute for Plant Science, University of Heidelberg, Heidelberg, Germany) for helpful discussions. We also thank Dr Martin Fulda (Department of Plant Biochemistry, Georg-August-Universität Göttingen, Göttingen, Germany) for plasmids; Dr Andreas Wodarz and Dr Michael Krahn (Department of Stem Cell Biology, Georg-August-Universität Göttingen, Göttingen, Germany) for access to the confocal microscope and technical support respectively, and Susanne Mesters for expert plant culture. Financial support was provided through an Emmy Noether Grant from the German Research Foundation [DFG (Deutsche Forschungsgemeinschaft)] awarded to I. H.

Abbreviations

     
  • BiP

    ER-luminal binding protein

  •  
  • CCV

    clathrin-coated vesicle

  •  
  • DTT

    dithiothreitol

  •  
  • ER

    endoplasmic reticulum

  •  
  • EYFP

    enhanced yellow fluorescent protein

  •  
  • HVA

    high-voltage activated

  •  
  • MBS

    main beam splitter

  •  
  • PH domain

    pleckstrin homology domain

  •  
  • PI

    phosphoinositide

  •  
  • PLC

    phospholipase C

  •  
  • PMA2

    plasma membrane ATPase 2

  •  
  • PtdCho

    phosphatidylcholine

  •  
  • PtdEtn

    phosphatidylethanolamine

  •  
  • RD20

    responsive to desiccation 20

  •  
  • TGA2

    basic/leucine-zipper transcription factor 2

  •  
  • V-ATPase

    vacuolar H+-ATPase

References

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Supplementary data