Lipid Droplet-Associated Proteins (LDAPs) Are Required for the Dynamic Regulation of Neutral Lipid Compartmentation in Plant Cells

Abstract

Eukaryotic cells compartmentalize neutral lipids into organelles called lipid droplets (LDs), and while much is known about the role of LDs in storing triacylglycerols in seeds, their biogenesis and function in nonseed tissues are poorly understood. Recently, we identified a class of plant-specific, lipid droplet-associated proteins (LDAPs) that are abundant components of LDs in nonseed cell types. Here, we characterized the three LDAPs in Arabidopsis (Arabidopsis thaliana) to gain insight to their targeting, assembly, and influence on LD function and dynamics. While all three LDAPs targeted specifically to the LD surface, truncation analysis of LDAP3 revealed that essentially the entire protein was required for LD localization. The association of LDAP3 with LDs was detergent sensitive, but the protein bound with similar affinity to synthetic liposomes of various phospholipid compositions, suggesting that other factors contributed to targeting specificity. Investigation of LD dynamics in leaves revealed that LD abundance was modulated during the diurnal cycle, and characterization of LDAP misexpression mutants indicated that all three LDAPs were important for this process. LD abundance was increased significantly during abiotic stress, and characterization of mutant lines revealed that LDAP1 and LDAP3 were required for the proper induction of LDs during heat and cold temperature stress, respectively. Furthermore, LDAP1 was required for proper neutral lipid compartmentalization and triacylglycerol degradation during postgerminative growth. Taken together, these studies reveal that LDAPs are required for the maintenance and regulation of LDs in plant cells and perform nonredundant functions in various physiological contexts, including stress response and postgerminative growth.

Figure 1.

Properties of Arabidopsis LDAPs. A, Deduced polypeptide sequence alignment, with positively and negatively charged residues highlighted in red and blue and identical and similar residues indicated with asterisks and colons or periods, respectively. The two Cys residues in LDAP3 (positions 168 and 196) described in the LDAP3 liposome-binding assays (Fig. 2C) are underlined. B, Reverse transcription (RT)-PCR analysis of LDAP gene expression in various tissues and developmental stages, as indicated by labels. ELONGATION FACTOR1-α (EF1α) served as an endogenous control. Additional controls for RT-PCR primer specificity are shown in Supplemental Figure S7B. C, Representative CLSM images of LDAP1-Cherry localization in various vegetative cell types of 15-d-old stably transformed Arabidopsis seedlings. Note the colocalization of LDAP1-Cherry with BODIPY-stained LDs in each cell type, as indicated by labels. Boxes represent the portion of the cell shown at higher magnification, revealing an LDAP1-Cherry torus-shaped fluorescence pattern surrounding the BODIPY-stained TAG core and indicating that LDAP1 is localized to the surface of LDs. Similar subcellular localizations for LDAP2 and LDAP3 in Arabidopsis are shown in Supplemental Figure S2. Also shown for each cell type are the corresponding chlorophyll autofluorescence and differential interference contrast (DIC) images. Bar = 20 μm.

Figure 2.

Subcellular targeting and biophysical interactions of LDAP3 with LDs and synthetic liposomes. A, Truncation analysis of LDAP3 in tobacco cv BY-2 cells. The cv BY-2 cells were transiently transformed with full-length or a modified version of LDAP3-GFP, stained with the neutral lipid dye monodansylpentane (MDH), and imaged using CLSM. The cv BY-2 cells were incubated with LA to induce LD proliferation (Supplemental Fig. S3), unless indicated otherwise. Shown on the left are cartoon representations of the various LDAP3-GFP constructs and their corresponding subcellular localization(s) in cv BY-2 cells (Cyt, cytosol). Shown on the right are representative micrographs for each LDAP3-GFP protein along with the corresponding MDH-stained LDs (false-colored red) in the same cell. Bar = 10 μm. B, Biophysical analysis of LDAP3 interaction with LDs in vivo. LDAP3-GFP, OLEO1-GFP, or GFP-DGAT2 was expressed transiently (as indicated by labels) in cv BY-2 cells incubated with LA. Cells were then fixed and extracted with either digitonin, which perturbs primarily the plasma membrane, or Triton X-100, which perturbs all cellular membranes, and then stained with MDH. Note that LDAP3 was resistant to digitonin extraction, but, unlike OLEO1 and DGAT2, LDAP3 was sensitive to Triton X-100 extraction, whereby the majority of protein was dissociated to the cytosol (left images). Bar = 10 μm. C, LDAP3 synthetic liposome-binding assays. Recombinant LDAP3 was purified (Supplemental Fig. S5), labeled at its single Cys with donor fluorophore, then mixed with a range of concentrations of acceptor fluorophore-labeled liposomes of various phospholipid compositions (Supplemental Table S1). Binding was assessed based on FRET efficiency (i.e. based on the change in fluorescence of the fluor-labeled donor protein when acceptor fluor-containing liposomes were present). While LDAP3 (red curves) exhibited different maximal FRET efficiencies at saturation for liposomes composed of different lipids, the protein displayed similar moderate binding to all liposomes in a manner that was stronger than the negative control protein (GroEL; green curves) but weaker than the positive control protein (BIM; blue curves). The highest concentration of liposomes is the largest amount that could be added to the reactions. Calculated dissociation constant values for protein-liposome-binding assays are presented in Table I. Mito, Mitochondria; PM, plasma membrane.

Figure 3.

LD abundance in Arabidopsis leaves during the diurnal cycle and in LDAP transgenic plants. A, Diurnal regulation of LD abundance in Arabidopsis leaves. Wild-type (WT) plants were grown on one-half-strength Murashige and Skoog (MS) plates for 15 d in a 16-h/8-h day/night cycle (lights on at 7 am and off at 11 pm), then leaves were harvested at the indicated times and LDs were examined by BODIPY staining and CLSM. Representative images are shown on the left, and quantifications of LDs are shown on the right. B, Overexpression of LDAPs in leaves. Two independent, homozygous, single-copy lines were generated for overexpression of each LDAP (i.e. LDAP-Cherry; Supplemental Fig. S7), then leaves were collected and imaged at 11 pm, when LD abundance is low in the wild type (see A). Representative CLSM images of each plant line are shown on the left, and quantifications of LDs are shown in the bar graph on the right. The graphs in the middle show neutral lipid content and composition of plant leaves showing increases in total neutral lipids due primarily to increases in polyunsaturated (i.e. 18:2 and 18:3) fatty acids (FA). C, Suppression of LDAPs in leaves. Two independent T-DNA and/or RNAi lines were generated for each LDAP (Supplemental Fig. S7), then leaves were collected and imaged (using CLSM) at 7 am, when LD abundance is high in the wild type (see A). All of the LDAP-disrupted lines, except ldap3-2, showed decreases in LD abundance (left graph) and no or moderate changes in neutral lipid content (middle graph) or fatty acid composition (right graph). Values of quantified LDs in A to C represent averages and sd from three biological replicates. Values of lipids in B and C represent averages and sd from five biological replicates. Arrowheads represent statistically significant differences above (pointing up) or below (pointing down) the wild-type value as determined by Student’s t test (P < 0.05). FW, Fresh weight. Bars in A, B, and C = 20 μm.

Figure 4.

Proliferation of LDs and LDAP expression in plant leaves during abiotic stress responses. Wild-type (WT) and selected ldap mutant lines were grown on one-half-strength MS plates for 15 d, then a portion of the plates were transferred to either a 4°C chamber for 24 h (A) or a 37°C chamber for 1 h (B). Leaves were collected at 0 and 24 h from control (C) and cold-stressed (CS) plants or at 0 and 1 h for control or heat-stressed (HS) plants, LDs were analyzed by BODIPY staining and CLSM, and transcript levels, including tubulin serving as an endogenous control, were evaluated using RT-PCR. Wild-type plants showed an approximately 10-fold increase in LD abundance in response to cold temperature (bar graph) and significant increases in transcript levels of both LDAP1 and LDAP3 genes (DNA gels). Similar results were observed in the ldap1-1 and ldap2-1 mutants, but the abundance of LDs in ldap3-1 during the cold temperature response was reduced significantly (bar graph). Results from heat stress experiments revealed that LDs proliferated approximately 10-fold in the wild type (bar graph), and LDAP1 transcripts were selectively and strongly induced (DNA gels). LDs were induced similarly in ldap2-1 and ldap3-1 mutants but were reduced significantly in ldap1-1 during the stress response. Values of quantified LDs represent averages and sd from three biological replicates. Arrowheads represent statistically significant differences in comparison with the wild type as determined by Student’s t test (P < 0.05).

Figure 5.

Effects of LDAP overexpression on seed development and during postgerminative growth. Two independent, single-copy, homozygous transgenic lines expressing LDAP-Cherry proteins were generated (Supplemental Fig. S7), seeds/seedlings were visualized by CLSM to evaluate LDAP localization in comparison with BODIPY-stained LDs, and seed oil content and composition were determined. A, Representative CLSM images of mature, dry seeds showing the localization of LDAPs to distinct punctate structures (left images) that do not colocalize with BODIPY-stained LDs (middle images) in merged images (right images). B, Representative CLSM images of seedlings 1 d after the onset of germination, showing the partial colocalization of LDAPs and BODIPY-stained LDs; boxes represent the portions of cells shown at higher magnification, showing the LDAP localization to torus-shaped structures surrounding BODIPY-stained TAG cores. Bars in A and B = 5 μm. C, Total fatty acids (FA) in mature seeds, showing statistically significant changes in two LDAP transgenic lines but no obvious trends due to LDAP overexpression. D, Fatty acid composition analysis of mature seeds, showing small but statistically significant changes but without any obvious trends due to LDAP overexpression. Values in C and D represent averages and sd of five biological replicates. Arrowheads represent statistically significant values above (pointing up) or below (pointing down) wild-type (WT) values as determined by Student’s t test (P < 0.05).

Figure 6.

Effects of LDAP suppression on seed development and postgerminative growth. Two different homozygous suppression mutants, T-DNA knockout and/or RNAi, were generated for each LDAP (Supplemental Fig. S7). A, Representative CLSM images of mature, dry seeds or seedlings 1 d after the onset of germination showing LDs stained with BODIPY. Note the similarity in LD morphology in all dry seeds (top row) and the altered LD phenotype in 1-d-old ldap1-1 seedlings in comparison with the wild type (WT), ldap2-1, or ldap3-1 (bottom row). B, Dry and 1-d-old seedlings from ldap1-2, showing a similar phenotype in comparison with ldap1-1 (see A). C and D, Total fatty acids (FA; C) and fatty acid compositional analysis (D) of mature seeds from the indicated plant lines, showing moderate changes in seed oil content and composition in some of the ldap mutants. E and F, Analysis of seed oil breakdown in wild-type, ldap1-1, and ldap1-2 lines, showing total fatty acids (E) and individual fatty acid (F) amounts in mature seeds and during postgerminative growth (i.e. 1, 2, and 4 d after the initiation of germination). DW, Dry weight; FW, fresh weight. All bar graphs represent averages and sd of five biological replicates, and arrowheads represent statistically significant values above (pointing up) or below (pointing down) wild-type levels as determined by Student’s t test (P < 0.05). G and H, Representative CLSM images of wild-type, ldap1-1, and ldap1-2 lines at 2 d (G) and 4 d (H) after the initiation of germination, showing more similar LD morphology in comparison with the wild type in the two ldap1 mutant lines on day 2 relative to day 1 (compare G with A and B) and normal LD morphology in both mutant lines at day 4 (compare with the wild type in H). Bars in A, B, G, and H = 5 μm.

Figure 7.

LDAPs and oleosin function similarly to compartmentalize lipids, but when ectopically expressed in the same cell, oleosin disrupts the binding of LDAP to LDs. A, Representative CLSM images of tobacco leaves transiently transformed with p19 (serving as a suppressor of gene silencing; Petrie et al., 2010) or p19 and either OLEO1-Cherry or LDAP-Cherry (or a modified version thereof) along with or without Arabidopsis LEC2, as indicated. LDs in all cells were stained with BODIPY. Note the presence of supersized LDs (indicated with arrowheads) in cells transformed with LEC2 and p19 (top row) or LEC2, p19, and LDAP3ΔC46-Cherry (bottom row), which does not target to LDs (Fig. 2A). By contrast, all cells coexpressing LEC2 (and p19) with either oleosin or an LDAP possess normal-sized LDs in comparison with controls without LEC2 (left images). DIC, Differential interference contrast. Bar = 20 μm. B, RT-PCR analysis of LEC2 gene expression, confirming the presence of LEC2 transcripts in all samples cotransformed with LEC2. ACTIN served as an endogenous control. C, Coexpression of oleosin and LDAP3 in tobacco cv BY-2 cells. Representative CLSM images show the localization of OLEO1-Cherry to MDH-stained LDs and the cytosolic (mis)localization of LDAP3-GFP in the same cell (top row; compare with images of oleosin and LDAP3 localized to LDs in individually transformed cv BY-2 cells in Fig. 2, A and B). By contrast, when LDAP3-GFP is coexpressed with the OLEO1-ΔPKM-Cherry mutant, which is retained in the ER (Abell et al., 1997; see also images in the bottom row), the localization of LDAP3-GFP to LDs in the same cell is enhanced (middle row). Bar = 10 μm.