Skip to main content
NCBI home page
The Plant Cell logoLink to The Plant Cell
. 2010 Sep 24;22(9):3066–3075. doi: 10.1105/tpc.110.077974

Arabidopsis ABCG Transporters, Which Are Required for Export of Diverse Cuticular Lipids, Dimerize in Different Combinations[W]

Heather E McFarlane a, John JH Shin a, David A Bird b, A Lacey Samuels a,1
PMCID: PMC2965547  PMID: 20870961

Arabidopsis ATP binding cassette (ABC) transporters, ABCG11 and ACBG12, are required for export of different lipids from the epidermis to the cuticle. This study shows that ABCG11 and ABCG12 can form an obligate heterodimer and that ABCG11 can homodimerize. Live-cell imaging in epidermal cells demonstrates that localization of these ABCGs to the plasma membrane is dependent upon dimerization.

Abstract

ATP binding cassette (ABC) transporters play diverse roles, including lipid transport, in all kingdoms. ABCG subfamily transporters that are encoded as half-transporters require dimerization to form a functional ABC transporter. Different dimer combinations that may transport diverse substrates have been predicted from mutant phenotypes. In Arabidopsis thaliana, mutant analyses have shown that ABCG11/WBC11 and ABCG12/CER5 are required for lipid export from the epidermis to the protective cuticle. The objective of this study was to determine whether ABCG11 and ABCG12 interact with themselves or each other using bimolecular fluorescence complementation (BiFC) and protein traffic assays in vivo. With BiFC, ABCG11/ABCG12 heterodimers and ABCG11 homodimers were detected, while ABCG12 homodimers were not. Fluorescently tagged ABCG11 or ABCG12 was localized in the stem epidermal cells of abcg11 abcg12 double mutants. ABCG11 could traffic to the plasma membrane in the absence of ABCG12, suggesting that ABCG11 is capable of forming flexible dimer partnerships. By contrast, ABCG12 was retained in the endoplasmic reticulum in the absence of ABCG11, indicating that ABCG12 is only capable of forming a dimer with ABCG11 in epidermal cells. Emerging themes in ABCG transporter biology are that some ABCG proteins are promiscuous, having multiple partnerships, while other ABCG transporters form obligate heterodimers for specialized functions.

INTRODUCTION

ATP binding cassette (ABC) transporters are universal components of cells from all kingdoms and play diverse roles, including lipid transport. Many ABC transporters are encoded as fully functional units, consisting of two ATP binding cassettes and two transmembrane domains. ABCG family members that are encoded as half-transporters must dimerize to form a full, functional ABC transporter. In Arabidopsis thaliana, the ABCG gene family contains 28 genes annotated as half-transporters (Verrier et al., 2008). ABCG half-transporters have been implicated in the transport of abscisic acid (Kuromori et al., 2010) and export of kanamycin (Mentewab and Stewart, 2005), sporopollenin (Quilichini et al., 2010; Xu et al., 2010), and cuticular lipids (Pighin et al., 2004; Bird et al., 2007; Luo et al., 2007; Panikashvili et al., 2007; Ukitsu et al., 2007). Whether these half-transporters function as homodimers or heterodimers is not known.

A model of ABCG half-transporters forming different heterodimers to perform multiple functions has been proposed in other eukaryotic systems. In Drosophila melanogaster, WHITE, SCARLET, and BROWN genes are involved in eye pigment accumulation and WHITE is also involved in cGTP transport (Sullivan and Sullivan, 1975; Sullivan et al., 1979; Mackenzie et al., 2000; Evans et al., 2008). Mutant phenotypes have indicated that WHITE is required for accumulation of both guanine-derived (red) or Trp-derived (brown) eye pigments, as the eyes of white mutants lack any pigmentation (Sullivan et al., 1980; Ewart et al., 1994). Similarly, brown and scarlet mutants indicated that these half-transporters are required for accumulation of guanine-derived or Trp-derived eye pigments, respectively (Ewart et al., 1994). This implies that WHITE and BROWN dimerize to transport guanine-derived pigments, while WHITE and SCARLET dimerize to transport Trp-derived pigments. Whether these gene products physically interact in vivo and whether they directly transport these substrates has not been tested.

ABCG transporters in mammals appear to form either obligate heterodimers, such as the ABCG5/ABCG8, or homodimers, such as ABCG2 (Tarr et al., 2009). While both ABCG5 and ABCG8 can interact promiscuously with other ABCG transporters tested in vitro (Graf et al., 2003), mouse ABCG5 and ABCG8 must dimerize to exit the endoplasmic reticulum (ER), undergo posttranslational modification in the Golgi, and traffic to their usual plasma membrane location (Graf et al., 2002, 2003). If either ABCG5 or ABCG8 is expressed individually, or if heterodimerization is disrupted by a point mutation in either transporter, both transporters are retained in the ER and degraded, indicating that formation of the ABCG5-ABCG8 heterodimer is a prerequisite for plasma membrane localization (Graf et al., 2002, 2004). These studies suggest that only a subset of the ABCG transporter dimers that may be detected by immunoprecipitation are capable of exiting the protein maturation machinery in the ER and that ER exit can be used as an assessment of the biological relevance of functional dimers.

Two ABCG half-transporters, ABCG11 and ABCG12, have been implicated in the export of lipids from the epidermis to the cuticle, which seals and protects the aerial tissues of the plant body (Bird, 2008). The cuticle is a tough cross-linked cutin polyester scaffold composed of hydroxy-C16 and C18 fatty acids and glycerol (Pollard et al., 2008), surrounded by and covered with a hydrophobic wax mixture dominated by very-long-chain fatty acid derivatives (Jetter et al., 2006; Samuels et al., 2008). ABCG half-transporters required for accumulation of both cutin and wax at the cell surface have been identified by mutant analysis in Arabidopsis. abcg11/wbc11/desperado/cof1 mutants display reduced cutin and wax levels (Bird et al., 2007; Luo et al., 2007; Panikashvili et al., 2007; Ukitsu et al., 2007). Detailed analyses of these chemical phenotypes revealed that all wax and cutin constituents were decreased in these mutants. Assuming that ABCG11 can directly transport cuticular lipids, these data imply that ABCG11 has a broad substrate specificity for a variety of structurally diverse cuticular lipids (Bird et al., 2007; Luo et al., 2007; Panikashvili et al., 2007; Ukitsu et al., 2007). The closely related ABCG12/CER5 transporter is required for wax (Pighin et al., 2004), but not cutin, export (Bird et al., 2007). abcg12 mutants have a reduction only in wax components, suggesting that ABCG12 has a narrower substrate specificity than ABCG11 (Pighin et al., 2004). Fluorescently tagged ABCG11 or ABCG12 is localized to the plasma membrane and rescues cuticular lipid deficiencies of the knockout mutants in stably transformed lines (Pighin et al., 2004; Bird et al., 2007). Consistent with a role in cuticular lipid export, both genes are highly expressed in the stem epidermis, where wax and cutin synthesis and secretion are extremely high (Pighin et al., 2004; Suh et al., 2005; Bird et al., 2007; Luo et al., 2007; Panikashvili et al., 2007; Ukitsu et al., 2007). However, ABCG11 is also expressed in tissues in which cuticular lipids are not synthesized (e.g., emerging lateral roots), suggesting that it may play roles beyond cuticular lipid export (Bird et al., 2007; Luo et al., 2007; Panikashvili et al., 2007; Ukitsu et al., 2007). Indeed, abcg11 mutants display pleiotropic phenotypes, including dwarfism, loss of apical dominance, and sterility, while abcg12 mutants are phenotypically normal except for their glossy stems (Pighin et al., 2004; Bird et al., 2007; Luo et al., 2007; Panikashvili et al., 2007; Ukitsu et al., 2007). Epidermal cells of both abcg11 and abcg12 mutants accumulate lipidic inclusions, presumably due to the aggregation of wax molecules that are synthesized, but not exported, from these mutants (Pighin et al., 2004; Bird et al., 2007; Panikashvili et al., 2007). abcg11 abcg12 double mutants have the same levels of residual wax as either of the single mutants, suggesting that ABCG11 and ABCG12 act in the same pathway or complex in cuticular wax export (Bird et al., 2007).

Based on the pleiotropic phenotypes of abcg11 mutants, we hypothesized that different dimerization combinations of ABCG transporters could account for the multiple functions of the ABCG11 half-transporter (Bird et al., 2007). In this study, ABCG11/ABCG12 heterodimers and ABCG11 homodimers were demonstrated using bimolecular fluorescence complementation (BiFC), while ABCG12 homodimers were not. Heterodimerization between ABCG11 and ABCG12 was further supported by the behavior of the ABC transporters during their biosynthesis and secretion in epidermal cells, which actively synthesize and secrete cuticular lipids. The different dimer combinations demonstrated by BiFC can account for the behavior of these half-transporters during their trafficking. ABCG12 was retained in lipidic inclusions in the absence of ABCG11, and these inclusions were contiguous with the ER. This study emphasizes the flexibility of the half-transporter system of ABCG transporters, in which specific dimer combinations perform specialized functions and different combinations of ABCG transporters can perform different functions.

RESULTS

ABCG11 and ABCG12 Form a Heterodimer, and ABCG11 Can Homodimerize

Given that ABCG11 and ABCG12 are both half-transporters (Verrier et al., 2008) and that the double mutant phenotype indicated that they likely act in the same pathway or complex in wax export (Bird et al., 2007), their physical interaction was tested in vivo using BiFC (Figure 1). This plant-based system was used because it has proven difficult, both in our hands and as documented by others (Geisler and Murphy, 2006; Yang and Murphy, 2009), to express plant ABC transporters in Saccharomyces cerevisiae, thus limiting yeast protein–protein interaction studies. In the BiFC system, enhanced yellow fluorescent protein (EYFP) is split into two halves: N-terminal (nYFP) and C-terminal (cYFP), which are fused to the proteins of interest. Cotransformation of these constructs into a transient expression system in Arabidopsis leaf mesophyll protoplasts allows reconstitution of YFP if the two proteins are in close physical proximity, thereby allowing detection of YFP signal (Citovsky et al., 2006). A significant proportion of protoplasts cotransformed with cYFP-ABCG11 and nYFP-ABCG12, or conversely, with cYFP-ABCG12 and nYFP-ABCG11, displayed bright yellow fluorescence in the plasma membrane, indicating that these two proteins are capable of forming a heterodimer in vivo (P < 0.05) (Figures 1C, 1D, and 1I; see Supplemental Figure 1 online). This also confirms that ABCG BiFC constructs with either the C- or N-terminal pieces of YFP are capable of generating yellow fluorescence, regardless of the combination used.

Figure 1.

Figure 1.

Open in a new tab

ABCG11 and ABCG12 Heterodimerize and ABCG11 Homodimerizes in the BiFC System.

Only protoplasts with intact plasma membranes, shown with bright-field light microscopy (A, C, E, G), were tested for the presence of yellow fluorescence, indicating protein–protein interaction due to assembly of split YFP, shown with confocal microscopy ([B], [D], [F], and [H]). Cotransformation of cYFP-ABCG11 and nYFP-ABCG11 into protoplasts with intact plasma membranes (A) generates yellow fluorescence (false-colored green) at the plasma membrane, surrounding chloroplast autofluorescence (false-colored magenta) in confocal (B). Cotransformation of cYFP-ABCG11 and nYFP-ABCG12 also generates yellow fluorescence at the plasma membrane ([C] and [D]). However, cotransformation of cYFP-ABCG12 and nYFP-ABCG12 ([E] and [F]) or cYFP-ABCG11 and NRT3.1-nYFP ([G] and [H]) generated only faint yellow fluorescence in a low proportion of protoplasts. Results were quantified as the percentage of fluorescent protoplasts, relative to the number of fluorescent protoplasts in the positive control (full-length YFP) (I). n = 8 independent experiments (total of >1300 protoplasts scored), bars represent se, * denotes statistically significant differences between samples (P < 0.05), and bars = 10 μm. All protoplasts (except for the overexpressing YFP control) were imaged at the same exposure time and detector gain to allow for comparisons of relative fluorescence.

When cYFP-ABCG11 and nYFP-ABCG11 were cotransformed into protoplasts, yellow fluorescence was observed in the plasma membrane of a significant proportion of these cells, indicating that ABCG11 can also homodimerize (P < 0.05) (Figures 1A, 1B, and 1I). By contrast, when cYFP-ABCG12 and nYFP-ABCG12 were cotransformed, only a small proportion of protoplasts fluoresced faintly when imaged under the same conditions (Figures 1E, 1F, and 1I; see Supplemental Figure 1 online). To determine whether this signal was due to a genuine, low-level homodimerization of ABCG12 or whether it was an artifact of colocalization of two half-YFP constructs to the plasma membrane, each of the ABCG BiFC constructs were cotransformed with BiFC constructs for components of a plasma membrane–localized root nitrate transporter complex (Arabidopsis NRT2.1 or NRT3.1) (Yong et al., 2010), which are not predicted to interact with ABCG transporters. Protoplasts cotransformed with NRT2.1-cYFP and either nYFP-ABCG11 or nYFP-ABCG12 or with NRT3.1-nYFP and either cYFP-ABCG11 or cYFP-ABCG12 displayed the same faint fluorescence as cYFP-ABCG12 with nYFP-ABCG12 (Figures 1E to 1I; see Supplemental Figure 1 online). These proportions of fluorescent protoplasts were not significantly different from the proportion observed when the two ABCG12 constructs were cotransformed (P > 0.35). Thus, these low proportions of faintly fluorescing protoplasts are likely background signal due to chance associations between proteins that are plasma membrane localized but do not legitimately interact.

ABCG11 Trafficking to the Plasma Membrane Is Independent of ABCG12

Studies of ABCG half-transporters in mammalian cells indicate that both partners of a heterodimer must be present for the dimer to traffic to the plasma membrane (Graf et al., 2003, 2004). To test whether ABCG11 or ABCG12 can traffic normally only in the presence of their partners and to put these interactions into the context of cuticular lipid secretion in the epidermis, stably transformed plant lines expressing fluorescently tagged ABCGs (Pighin et al., 2004; Bird et al., 2007) were crossed into the single (abcg11) and double (abcg11 abcg12) knockout mutants. YFP-ABCG11 successfully trafficked to the plasma membrane in the presence of its wild-type ABCG12 partner (i.e., in the single abcg11 mutant background; Figure 2A). Plasma membrane localization of YFP-ABCG11 was visualized as two distinct signals from neighboring cells lining the adjacent cell wall, which was labeled with propidium iodide (Figures 2B and 2C). In the absence of ABCG12 (in abcg11 abcg12 double knockout mutant background), YFP-ABCG11 was also localized to the plasma membrane (Figure 2E), where it colocalized with the plasma membrane marker FM4-64 (Figures 2F and 2G). YFP-ABCG11 localization in the double mutant background was confirmed using transmission electron microscopy (TEM) immunogold labeling of cryo-fixed, freeze-substituted stem epidermal cells (Figure 2D), probed with an anti-green fluorescent protein (GFP) antibody. The plasma membrane of epidermal cells was labeled by anti-GFP in transgenic plant lines carrying YFP-ABCG11 (Figure 2H), but no signal above background was detected in wild-type control plants or in samples probed without a primary antibody (see Supplemental Figure 2 online). There was no evidence of polar localization of YFP-ABCG11 in stem epidermal cells using this high-resolution TEM technique, in contrast with previous reports using confocal microscopy, where signal intensities can be distorted by optical conditions or dissection (Panikashvili et al., 2007). These results demonstrate that ABCG11 is able to traffic to the plasma membrane even in the absence of ABCG12. This indicates that ABCG11 may heterodimerize with other ABCG half-transporters that are present in the stem epidermis, such as ABCG18 and ABCG19 (Suh et al., 2005). Alternatively, ABCG11 may form a homodimer in vivo, an interpretation that is consistent with the BiFC data.

Figure 2.

Figure 2.

Open in a new tab

Trafficking of ABCG11 to the Plasma Membrane Is Independent of ABCG12.

In abcg11 mutants ([A] to [C]), YFP-ABCG11 is localized to the plasma membrane (A), as shown by counterstaining with propidium iodide (B) and merge (C). TEM of the wild-type stem illustrates epidermal cell morphology (D). In abcg11 abcg12 double mutants ([E] to [H]), YFP-ABCG11 is also localized to the plasma membrane (E), where it colocalizes with FM4-64 (F) in the merge (G). TEM immunogold localization with anti-GFP confirms YFP-ABCG11 localization to the plasma membrane (H). The white box in (G) highlights an area that is representative of the field of view in (H). Circles highlight gold particles, and arrowheads denote the plasma membrane. Bars = 10 μm in confocal images ([A] to [C] and [E] to [G]), 5 μm in (D), and 200 nm in (H).

ABCG12 Trafficking to the Plasma Membrane Is Dependent on ABCG11

If ABCG12 forms an obligate heterodimer with ABCG11, then ABCG11 is predicted to be required for normal traffic of ABCG12 to the plasma membrane. Therefore, GFP-ABCG12 was localized in abcg12 and abcg11 abcg12 mutants to investigate its trafficking in the presence and absence of its ABCG11 partner, respectively. In the single abcg12 knockout mutants, GFP-ABCG12 signal was detected at the plasma membrane (Figure 3A), adjacent to the cell wall, visualized with propidium iodide (Figures 3B and 3C). Immunogold TEM labeling of the single abcg12 mutant expressing the GFP-ABCG12 construct revealed that anti-GFP signal was predominantly at the plasma membrane, with little intracellular and cell wall background label (Figure 3D; see Supplemental Figure 2 online). However, in the abcg11 abcg12 double mutant background, GFP-ABCG12 signal was detected in a reticulate network resembling the ER and in large aggregations in the middle of the cells (Figure 3E). This signal had a low level of colocalization with FM4-64 (Figures 3F and 3G), which is not consistent with plasma membrane localization. In TEM, anti-GFP labeling was concentrated in the large aggregations of sheet-like inclusions that have been detected within the cytoplasm of abcg11 and abcg12 mutants (Figure 3H; Pighin et al., 2004; Bird et al., 2007; Panikashvili et al., 2007). GFP-ABCG12 retention in the ER in the absence of ABCG11 is consistent with the prediction that ABCG12 dimerizes only with ABCG11 in the stem epidermal cells and that, as in mammals, ABCG half-transporter dimer formation is required for trafficking to the plasma membrane (Graf et al., 2003, 2004).

Figure 3.

Figure 3.

Open in a new tab

Trafficking of ABCG12 to the Plasma Membrane Is Dependent on ABCG11.

In abcg12 mutants ([A] to [D]), GFP-ABCG12 is localized to the plasma membrane (A), as shown by counterstaining with the cell wall dye propidium iodide (B), merge (C), and TEM immunogold localization with anti-GFP (D). In abcg11 abcg12 double mutants ([E] to [H]), GFP-ABCG12 is retained within the ER and in lipidic inclusions of these mutants (E) and fails to colocalize with the plasma membrane dye FM4-64 (F) in the merge (G). TEM immunogold localization with anti-GFP confirms GFP-ABCG12 localization to inclusions (H). White boxes in (C) and (G) highlight an area that is representative of the field of view in (D) and (H), respectively, circles highlight gold particles, and arrowheads denote the plasma membrane. Bars = 10 μm in confocal images ([A] to [C] and [E] to [G]) and 200 nm in TEM ([D] and [H]).

Accumulation of GFP-ABCG12 in the abcg11 abcg12 mutant inclusions could be due to a nonspecific ER stress response and/or a general trafficking defect in abcg11 mutants. Immunogold TEM labeling for another plasma membrane–localized protein, the PIP2 aquaporin (Bots et al., 2005), detected PIP2 in the plasma membrane of wild-type and abcg11 mutant cells (Figures 4A and 4B). However, no labeling above background was detected in the inclusions (Figure 4C; see Supplemental Figure 3 online). This confirms that the failure of GFP-ABCG12 to traffic to the plasma membrane in abcg11 mutants is specific to ABCG12 and that the inclusions are not a general sink for plasma membrane–localized proteins. Similarly, RT-PCR analysis of genes typically upregulated by the unfolded protein response (Martinez and Chrispeels, 2003; Kamauchi et al., 2005) confirmed that the presence of inclusions does not induce significant unfolded protein response (see Supplemental Figure 4 online). These data agree with a recent microarray analysis of gene expression changes in abcg11 mutants (Panikashvili et al., 2010). Therefore, GFP-ABCG12 retention in inclusions is not due to a general protein or membrane trafficking defect in abcg11 mutants.

Figure 4.

Figure 4.

Open in a new tab

Trafficking of PIP2 to the Plasma Membrane Is Independent of ABCG11.

In wild-type stem epidermal cells, the PIP2 aquaporin is localized to the plasma membrane (A). PIP2 is also localized to the plasma membrane in abcg11 mutants (B) and is not retained within the abcg11 mutant inclusions (C). Circles highlight gold particles, arrowheads denote the plasma membrane, and bars = 200 nm.

Membrane Inclusions in abcg11 Mutants Are Contiguous with the ER

Because GFP-ABCG12 was retained in the ER and in the inclusions of abcg11 abcg12 double mutants, the relationship between these inclusions and the ER was investigated further. TEM was used to examine the morphology of abcg11 stem epidermal cells where inclusions protruded into the vacuole of epidermal cells (Figure 5A). ER membranes are often closely associated with the inclusions, either layered between inclusions, or at the tips of the inclusions (Figures 5B and 5C). By contrast, there was no relationship between the Golgi apparatus and the inclusions (Figure 5B).

Figure 5.

Figure 5.

Open in a new tab

Membrane Inclusions in abcg11 Mutants Are Closely Associated with the ER.

Stem epidermal cells of abcg11 mutants accumulate inclusions that protrude into the large central vacuole (A). These inclusions are closely associated and interspersed with the electron-lucent ER ([B] and [C]); however, the morphology of the Golgi apparatus seems unaffected (B). Arrows in (B) and (C) highlight close associations between ER and inclusions. Bars = 5 μm in (A), 500 nm in (B), and 200 nm in (C).

To confirm the association of the ER with the inclusions, ER was positively identified with established markers in abcg11 mutants. In wild-type epidermal cells, the ER marker GFP-HDEL (Batoko et al., 2000) labeled the ER network, including the cortical ER (Figure 6A). In abcg11 mutant stem epidermal cells, GFP-HDEL labeled large aggregations in the center of these cells, in addition to the normal ER network (Figure 6E). This pattern is strikingly similar to the lipidic inclusions labeled with Nile Red detected in abcg12 mutants (Pighin et al., 2004). In abcg11 mutants, Nile Red stained a similar pattern to GFP-HDEL (Figures 6B, 6C, 6F, and 6G), suggesting that these ER accumulations are lipid rich and may be composed of cuticular lipids that are synthesized, but not secreted, in abcg11 mutants.

Figure 6.

Figure 6.

Open in a new tab

Membrane Inclusions in abcg11 Mutants Are Contiguous with the ER.

In wild-type stem epidermal cells ([A] to [D]), GFP-HDEL labels the ER network (A), and Nile Red (a lipidic dye) faintly labels cellular membranes (B), which partially colocalize with GFP-HDEL in the merge (C). TEM immunogold labeling with anticalreticulin confirms the presence of this epitope in the wild-type electron-lucent ER (D). In abcg11 mutants ([E] to [H]), GFP-HDEL labels the reticulate ER network as well as large aggregations in the center of the cells (E). Nile Red signal indicates that these inclusions are lipidic (F), and GFP-HDEL and Nile Red overlap in inclusions in the merge (G). TEM immunogold labeling with anticalreticulin shows an accumulation of this ER epitope in inclusions (H). White boxes in (C) and (G) highlight an area that is representative of the field of view in (D) and (H), respectively, and circles highlight gold particles. Bars = 10 μm in confocal images ([A] to [C] and [E] to [G]) and 200 nm in TEM ([D] and [H]).

Since the ER was often layered between inclusions, the live-cell imaging with GFP-HDEL might reflect trapping of ER membranes between inclusions, rather than expansion of the ER membranes to form inclusions. To differentiate between these possibilities, TEM immunogold labeling was employed with anticalreticulin, an established marker for the ER (Coughlan et al., 1997). In wild-type cells, anticalreticulin labeled the electron-lucent ER lumen (Figure 6D; see Supplemental Figure 5 online). In abcg11 mutants, anticalreticulin labeled both ER lumen and inclusions (Figure 6H; see Supplemental Figure 5 online). This confirms that the inclusions themselves contain bona fide ER markers. Together, these data suggest that in abcg11 mutants, lipids are retained in inclusions that are associated with ER-derived membranes.

DISCUSSION

Specific dimer pairings of ABCG transporters have been hypothesized to allow multiple functions for a single gene product in both Arabidopsis cuticular lipid export by ABCG11 and ABCG12 (Bird et al., 2007, Bird, 2008) and in Drosophila eye pigment accumulation by WHITE, BROWN, and SCARLET (Sullivan and Sullivan, 1975; Sullivan et al., 1979; Mackenzie et al., 2000; Evans et al., 2008). However, these dimer combinations have been hypothesized based on sequence analysis and mutant phenotypes and have not been directly tested, until now. Here, physical interactions between Arabidopsis ABCG11 and ABCG12 are experimentally demonstrated by BiFC. BiFC data indicate which pairings are physically possible, assuming these gene products are expressed in the same cell at the same time. To place ABCG transporter dimerization in the context of the epidermal cell during lipid secretion, biosynthetic traffic of ABCG transporters from the ER to the plasma membrane was examined in these cells. The behavior of these half-transporters during their biosynthesis and trafficking are consistent with the different dimer combinations demonstrated by BiFC. The consistency between these two data sets, together with functional data deduced from mutant phenotypes (Pighin et al., 2004; Bird et al., 2007; Luo et al., 2007; Panikashvili et al., 2007; Ukitsu et al., 2007), lead to the models proposed below for specific dimerization pairings of ABCG transporters in diverse functions.

ABCG11 Undergoes Flexible Dimerization and Performs Multiple Functions

abcg11 mutants display pleiotropic phenotypes, including reduced surface wax and cutin monomers, stunted growth, organ fusions, reduced fertility, and reduced apical dominance, indicating that ABCG11 is involved in processes in addition to wax export (Bird et al., 2007; Luo et al., 2007; Panikashvili et al., 2007; Ukitsu et al., 2007). Furthermore, the expression pattern of ABCG11 extends beyond that of ABCG12 to tissues in which neither cutin nor wax is being synthesized (e.g., emerging lateral roots) (Suh et al., 2005; Toufighi et al., 2005; Bird et al., 2007; Luo et al., 2007; Panikashvili et al., 2007). Based on mutant phenotypes and expression pattern, we propose that ABCG11 acts as a generalist ABCG transporter, pairing with different half-transporters in different tissues to transport structurally diverse substrates. Consistent with this, ABCG11 was able to exit the ER and traffic to the plasma membrane independently of ABCG12, indicating that it can dimerize with other ABCG half-transporters in epidermal cells. Additionally, homodimerization of ABCG11 was demonstrated in BiFC assays. These results are consistent with the model in which ABCG11 homodimerizes to export cutin precursors from the stem epidermis (Bird et al., 2007). Determining whether ABCG11 functions in cutin export as a homodimer will require further experiments. While analysis of several cutin biosynthesis mutants has revealed genes required for cutin monomer biosynthesis and assembly, the nature of cutin precursors that are exported for assembly in the cuticle is not known (Pollard et al., 2008). This gap in our understanding, as well as the problem that wax constituents are solid at biologically relevant temperatures, makes assessment of the possible substrates of ABCG11 challenging. Both BiFC and trafficking data demonstrate that ABCG11 is capable of promiscuous pairing with multiple ABCG half-transporters. It is possible that ABCG11 dimerizes with multiple partners to form a variety of full transporters to perform diverse functions, as has been hypothesized in the Drosophila WHITE/BROWN/SCARLET model. Indeed, ABCG11 has been implicated in functions as diverse as maintenance of apical dominance, flower development, embryogenesis, and root suberin export (Panikashvili et al., 2010). This gives rise to a model in which the pleiotropic effects of abcg11 mutation are due to the loss of this variety of full transporters.

ABCG12 Specifically Forms Heterodimers with ABCG11 for Wax Export

In contrast with the broad expression pattern of ABCG11 and the pleiotropic phenotypes of abcg11 mutants, mutants in abcg12 display only wax-related phenotypes, and expression of ABCG12 is enriched in the stem epidermis where wax is actively being synthesized and secreted (Pighin et al., 2004; Suh et al., 2005; Toufighi et al., 2005). There is a precedent for strict ABCG heterodimerization in Medicago truncatula arbuscular mycorrhizal symbiosis (Zhang et al., 2010). During arbuscule formation, two ABCG half-transporter genes, STUNTED ARBUSCULE (STR) and STR2, are highly expressed. STR and STR2 heterodimerize and are incapable of forming homodimers. Their identical mutant phenotypes and expression patterns suggest that, like ABCG12, these two ABCG half-transporters are specialists (Zhang et al., 2010). Consistent with the hypothesis that ABCG12 is a specialist, GFP-ABCG12 was only able to exit the ER and move to the plasma membrane of epidermal cells in the presence of ABCG11, indicating that ABCG12 forms an obligate heterodimer with ABCG11 in stem epidermal cells. This is supported by the BiFC interaction data, which indicate that ABCG12 can heterodimerize with ABCG11 but is incapable of forming a homodimer. Therefore, the wax phenotypes of the abcg11 and abcg12 single mutants and the abcg11 abcg12 double mutants are a result of defects in this heterodimer.

Waxes That Are Not Exported in ABCG Mutants Accumulate in ER-Derived Inclusions

In abcg11 or abcg12 mutants, membranous lipidic inclusions accumulate in the epidermal cells (Pighin et al., 2004; Bird et al., 2007; Panikashvili et al., 2007). Here, we demonstrate that the inclusions in abcg11 mutants contain markers of the ER but not markers of the plasma membrane. It is possible that ER expansion is a stress response to sequester insoluble cuticular lipids. Similarly, in yeast, the ER increases in size to accommodate defective proteins, independent of the unfolded protein response (Schuck et al., 2009). Alternatively, the inclusions could reflect the nature of lipid traffic during cuticular lipid secretion. All of the enzymes that have been characterized in wax synthesis have been localized to the ER membranes (Zheng et al., 2005; Greer et al., 2007; Bach et al., 2008; Li et al., 2008). If intermediate secretory compartments, such as the Golgi apparatus, are required for wax secretion to the cell surface, then wax accumulation in these compartments would be expected to occur when transport is blocked. However, Golgi and plasma membrane morphology appeared unaffected in TEM of abcg11 knockout mutants. While the subcellular compartments involved in wax secretion are not known, accumulation of waxes only in the ER is consistent with wax moving directly from the ER to the plasma membrane via nonvesicular lipid traffic, by analogy to yeast lipid transport systems (Levine and Loewen, 2006).

In summary, Arabidopsis ABCG11 is capable of flexible dimerization to form either a heterodimer or a homodimer at the plasma membrane in vivo. By contrast, ABCG12 forms an obligate heterodimer with ABCG11, and heterodimerization is required for normal trafficking to the plasma membrane. Based on these data and on previous ABCG studies, there are two emerging paradigms: flexible pairing of one ABCG gene product with multiple partners to perform diverse functions and formation of obligate ABCG heterodimers for specialized functions. These paradigms have been formerly deduced but not experimentally tested in the canonical WHITE/BROWN/SCARLET complex in Drosophila. In mammalian cells, cases of biologically relevant flexible ABCG transporter partnering have not been described, although several lines of evidence hint that this is possible (Graf et al., 2002; Cserepes et al., 2004). This study demonstrates that mechanisms of ABCG transporter dimerization are conserved across biological kingdoms, despite the divergent functional roles played by these transporters.

METHODS

Plant Material

Seedlings were sown on AT media and grown in an environmental growth chamber at 21°C, 70 to 80% humidity, and 24 h light (80 to 100 μE m−2 s−1) for roughly 10 d before transfer to Sunshine Mix 5 soil. The cer5-2, wbc11-3, cer5-2+CER5pro:GFP-CER5, and wbc11-3+35Spro:YFP-WBC11 lines (all in the Columbia-0 [Col-0] background) have been previously described (Pighin et al., 2004; Bird et al., 2007), and Col-0 seeds carrying the 35Spro:GFP-HDEL ER marker were a generous gift from Hugo Zheng (Batoko et al., 2000).

BiFC Transgene Construction and Protoplast Transformation

ABCG12 cDNA was amplified from the GFP:CER5 vector (Pighin et al., 2004), and XmaI and EcoRI sites were added using the primers ABCG12.P3 and ABCG12.P4 (see Supplemental Table 1 online). ABCG11 cDNA was amplified from YFP:WBC11 (Bird et al., 2007), and XmaI and EcoRI sites were added using the primers ABCG11.P3 and ABCG11.P4 (see Supplemental Figure 6 online). The resulting products were ligated into the EcoRI and XmaI sites of pSAT4-cEYFP-C1-B and pSAT4-nEYFP-C1 (Citovsky et al., 2006) and verified by sequencing. Control BiFC vectors containing components of the Arabidopsis thaliana nitrate transport machinery were a generous gift from Z. Kotur and A. Glass (Yong et al., 2010). Plasmid DNA was purified from Escherichia coli cultures using an endotoxin-free plasmid maxiprep kit (Qiagen). Protoplasts from Col-0 leaves were prepared and transformed as described (Tiwari et al., 2006) with 10 μg of each vector used in each protoplast transformation. The number of fluorescent protoplasts was scored out of the total number of live protoplasts (i.e., those with an intact plasma membrane when viewed with bright-field microscopy) and presented as a percentage relative to the number of fluorescent protoplasts in the positive control (pSAT6+35Spro:EYFP) as an estimation of transformation efficiency. Data from converse transformations (e.g., cYFP-ABCG11+nYFP-ABCG12 and cYFP-ABCG12+nYFP-ABCG11) were grouped. To meet the assumptions of analysis of variance (Whitlock and Schluter, 2009), data from eight independent experiments were natural log (ln) transformed and corrected for variations in transformation efficiency among experiments by subtracting the mean transformation efficiency of that experiment from each value. Means were compared using analysis of variance and a Tukey post-hoc test using SPSS (IBM).

Confocal Microscopy

Transformed protoplasts were mounted in WI solution, or dissected stem segments from the top 3 cm from the shoot apical meristem (where cuticle synthesis and secretion is the highest; Suh et al., 2005) were mounted in distilled water and immediately imaged. Stem segments were stained with 1 μg/mL propidium iodide (Sigma-Aldrich) or 10 μM FM4-64 (Molecular Probes) for 10 min. Images were collected using a Zeiss 510 Meta scan head on a Zeiss Axiovert 200 M with a Zeiss AxioCam HRm CCD camera or using a Quorum Wave FX spinning-disk scan head on a Leica DMI6000 microscope with a Hamamatsu ImagEM CCD camera. On the Zeiss microscope, GFP was detected using a 488-nm laser with a 505- to 530-nm filter, YFP was detected using a 514-nm laser with a 535- to 580-nm filter, and propidium iodide and FM4-64 were detected using a 514-nm laser with a 600- to 650-nm filter. On the Quorum system, GFP and YFP were detected using a 491-nm laser with a 528- to 566-nm filter. To allow comparison of the relative brightness between different BiFC treatments, all protoplasts (except the YFP overexpressing positive control) were imaged under identical conditions, including detector gain and exposure time. Images were processed using Volocity (Improvision) or ImageJ, and Adobe Illustrator.

High-Pressure Freezing, TEM, and Immunogold Labeling

Stem tissue from the top 1 to 3 cm from the shoot apical meristem was frozen in 0.2 M sucrose in B sample holders (Ted Pella) using a Leica HPM-100 high-pressure freezer. Freeze substitution, resin infiltration, sectioning, poststaining, and imaging were performed as described (McFarlane et al., 2008). Immunolabeling of high-pressure frozen material was performed as described (McFarlane et al., 2008). Primary antibodies were 1/50 polyclonal anti-GFP (A6455 from Molecular Probes); 1/20 polyclonal anti-calreticulin, generated against calreticulin from castor bean (Ricinus communis cv Hale), a generous gift from Sean Coughlan (Coughlan et al., 1997); and 1/50 polyclonal anti-PIP2, generated against PIP2 from tobacco (Nicotiana tabacum cv Petit Havana SR1), a generous gift from Ralf Kaldenhoff, (Bots et al., 2005). Secondary antibody was 1/100 10 nm gold-conjugated goat-anti-rabbit (Ted Pella). Images were processed using ImageJ and Adobe Illustrator.

Gene Expression Analysis

Total RNA was extracted using TRIzol reagent (Invitrogen) from stem segments 1 to 3 cm from the shoot apical meristem or from 10-d-old seedlings grown in liquid AT media plus 1% sucrose, with and without 5 mM DTT (as positive and negative controls for the unfolded protein response, respectively). cDNA was synthesized from 15 μg of RNA using an oligo dT18 primer and SuperScript III reverse transcriptase (Invitrogen). RT-PCR was performed for 20 cycles using 1 μL of cDNA with intron-flanking, gene-specific primers for HSP90.7, BiP1/BiP2 (not specific to either gene alone, since these are 97% identical), CALNEXIN1, CALRETICULIN2, and PDI-LIKE9 (see Supplemental Table 1 online). cDNA levels were normalized using primers for the UBC10 ubiquitin conjugating enzyme gene for 20 cycles. PCR products were visualized using SYBR-Safe (Invitrogen).

Accession Numbers

Sequence data from this article can be found in the Arabidopsis Genome Initiative or GenBank/EMBL databases under the following accession numbers: At1g17840 (ABCG11), At1g51500 (ABCG12), At5g53300 (UBC10), At4g24190 (HSP90.7), At5g28540 (BiP1), At5g42020 (BiP2), At5g61790 (Calnexin1), At1g09210 (Calreticulin2), and At2g32920 (PDI-like9).

Supplemental Data

The following materials are available in the online version of this article.

Supplementary Material

Supplemental Data
supp_22_9_3066__index.html (653B, html)

Acknowledgments

We thank the ABRC for providing seed stocks and BiFC vectors, Sean Coughlan for anticalreticulin, Ralf Kaldenhoff for anti-PIP2, Zorica Kotur for the nitrate transporter BiFC constructs, Hugo Zheng for the GFP-HDEL construct, the University of British Columbia Bioimaging Facility for technical assistance, and Ljerka Kunst, Mathias Schuetz, and Teagen Quilichini for helpful discussions and comments on the manuscript. This work was funded by Canadian Natural Sciences and Engineering Research Council Discovery Grants to D.A.B. and A.L.S. and by a Canada Graduate Scholarship-D3 to H.E.M.

References

  1. Bach L., et al. (2008). The very-long-chain hydroxy fatty acyl-CoA dehydratase PASTICCINO2 is essential and limiting for plant development. Proc. Natl. Acad. Sci. USA 105: 14727–14731 [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Batoko H., Zheng H.Q., Hawes C., Moore I. (2000). RAB1 GTPase is required for transport between the endoplasmic reticulum and Golgi apparatus and for normal Golgi movement in plants. Plant Cell 12: 2201–2218 [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Bird D., Beisson F., Brigham A., Shin J., Greer S., Jetter R., Kunst L., Wu X., Yephremov A., Samuels L. (2007). Characterization of Arabidopsis ABCG11/WBC11, an ATP binding cassette (ABC) transporter that is required for cuticular lipid secretion. Plant J. 52: 485–498 [DOI] [PubMed] [Google Scholar]
  4. Bird D.A. (2008). The role of ABC transporters in cuticular lipid secretion. Plant Sci. 174: 563–569 [Google Scholar]
  5. Bots M., Feron R., Uehlein N., Weterings K., Kaldenhoff R., Mariani T. (2005). PIP1 and PIP2 aquaporins are differentially expressed during tobacco anther and stigma development. J. Exp. Bot. 56: 113–121 [DOI] [PubMed] [Google Scholar]
  6. Citovsky V., Lee L.Y., Vyas S., Glick E., Chen M.H., Vainstein A., Gafni Y., Gelvin S.B., Tzfira T. (2006). Subcellular localization of interacting proteins by bimolecular fluorescence complementation in planta. J. Mol. Biol. 362: 1120–1131 [DOI] [PubMed] [Google Scholar]
  7. Coughlan S.J., Hastings C., Winfrey R., Jr (1997). Cloning and characterization of the calreticulin gene from Ricinus communis L. Plant Mol. Biol. 34: 897–911 [DOI] [PubMed] [Google Scholar]
  8. Cserepes J., Szentpétery Z., Seres L., Ozvegy-Laczka C., Langmann T., Schmitz G., Glavinas H., Klein I., Homolya L., Váradi A., Sarkadi B., Elkind N.B. (2004). Functional expression and characterization of the human ABCG1 and ABCG4 proteins: indications for heterodimerization. Biochem. Biophys. Res. Commun. 320: 860–867 [DOI] [PubMed] [Google Scholar]
  9. Evans J.M., Day J.P., Cabrero P., Dow J.A.T., Davies S.A. (2008). A new role for a classical gene: White transports cyclic GMP. J. Exp. Biol. 211: 890–899 [DOI] [PubMed] [Google Scholar]
  10. Ewart G.D., Cannell D., Cox G.B., Howells A.J. (1994). Mutational analysis of the traffic ATPase (ABC) transporters involved in uptake of eye pigment precursors in Drosophila melanogaster. Implications for structure-function relationships. J. Biol. Chem. 269: 10370–10377 [PubMed] [Google Scholar]
  11. Geisler M., Murphy A.S. (2006). The ABC of auxin transport: The role of p-glycoproteins in plant development. FEBS Lett. 580: 1094–1102 [DOI] [PubMed] [Google Scholar]
  12. Graf G.A., Cohen J.C., Hobbs H.H. (2004). Missense mutations in ABCG5 and ABCG8 disrupt heterodimerization and trafficking. J. Biol. Chem. 279: 24881–24888 [DOI] [PubMed] [Google Scholar]
  13. Graf G.A., Li W.P., Gerard R.D., Gelissen I., White A., Cohen J.C., Hobbs H.H. (2002). Coexpression of ATP-binding cassette proteins ABCG5 and ABCG8 permits their transport to the apical surface. J. Clin. Invest. 110: 659–669 [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Graf G.A., Yu L., Li W.P., Gerard R., Tuma P.L., Cohen J.C., Hobbs H.H. (2003). ABCG5 and ABCG8 are obligate heterodimers for protein trafficking and biliary cholesterol excretion. J. Biol. Chem. 278: 48275–48282 [DOI] [PubMed] [Google Scholar]
  15. Greer S., Wen M., Bird D., Wu X., Samuels L., Kunst L., Jetter R. (2007). The cytochrome P450 enzyme CYP96A15 is the midchain alkane hydroxylase responsible for formation of secondary alcohols and ketones in stem cuticular wax of Arabidopsis. Plant Physiol. 145: 653–667 [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Jetter R., Kunst L., Samuels L. (2006). Composition of plant cuticular waxes. In Biology of the Plant Cuticle, Riederer M., Muller C., (Oxford, UK: Blackwell Publishers; ), pp. 145–181 [Google Scholar]
  17. Kamauchi S., Nakatani H., Nakano C., Urade R. (2005). Gene expression in response to endoplasmic reticulum stress in Arabidopsis thaliana. FEBS J. 272: 3461–3476 [DOI] [PubMed] [Google Scholar]
  18. Kuromori T., Miyaji T., Yabuuchi H., Shimizu H., Sugimoto E., Kamiya A., Moriyama Y., Shinozaki K. (2010). ABC transporter AtABCG25 is involved in abscisic acid transport and responses. Proc. Natl. Acad. Sci. USA 107: 2361–2366 [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Levine T., Loewen C. (2006). Inter-organelle membrane contact sites: Through a glass, darkly. Curr. Opin. Cell Biol. 18: 371–378 [DOI] [PubMed] [Google Scholar]
  20. Li F., Wu X., Lam P., Bird D., Zheng H., Samuels L., Jetter R., Kunst L. (2008). Identification of the wax ester synthase/acyl-coenzyme A: diacylglycerol acyltransferase WSD1 required for stem wax ester biosynthesis in Arabidopsis. Plant Physiol. 148: 97–107 [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Luo B., Xue X.Y., Hu W.L., Wang L.J., Chen X.Y. (2007). An ABC transporter gene of Arabidopsis thaliana, AtWBC11, is involved in cuticle development and prevention of organ fusion. Plant Cell Physiol. 48: 1790–1802 [DOI] [PubMed] [Google Scholar]
  22. Mackenzie S.M., Howells A.J., Cox G.B., Ewart G.D. (2000). Sub-cellular localisation of the white/scarlet ABC transporter to pigment granule membranes within the compound eye of Drosophila melanogaster. Genetica 108: 239–252 [DOI] [PubMed] [Google Scholar]
  23. Martínez I.M., Chrispeels M.J. (2003). Genomic analysis of the unfolded protein response in Arabidopsis shows its connection to important cellular processes. Plant Cell 15: 561–576 [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. McFarlane H.E., Young R.E., Wasteneys G.O., Samuels A.L. (2008). Cortical microtubules mark the mucilage secretion domain of the plasma membrane in Arabidopsis seed coat cells. Planta 227: 1363–1375 [DOI] [PubMed] [Google Scholar]
  25. Mentewab A., Stewart C.N., Jr (2005). Overexpression of an Arabidopsis thaliana ABC transporter confers kanamycin resistance to transgenic plants. Nat. Biotechnol. 23: 1177–1180 [DOI] [PubMed] [Google Scholar]
  26. Panikashvili D., Savaldi-Goldstein S., Mandel T., Yifhar T., Franke R.B., Höfer R., Schreiber L., Chory J., Aharoni A. (2007). The Arabidopsis DESPERADO/AtWBC11 transporter is required for cutin and wax secretion. Plant Physiol. 145: 1345–1360 [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Panikashvili D., Shi J.X., Bocobza S., Franke R.B., Schreiber L., Aharoni A. (2010). The Arabidopsis DSO/ABCG11 transporter affects cutin metabolism in reproductive organs and suberin in roots. Mol. Plant 3: 563–575 [DOI] [PubMed] [Google Scholar]
  28. Pighin J.A., Zheng H., Balakshin L.J., Goodman I.P., Western T.L., Jetter R., Kunst L., Samuels A.L. (2004). Plant cuticular lipid export requires an ABC transporter. Science 306: 702–704 [DOI] [PubMed] [Google Scholar]
  29. Pollard M., Beisson F., Li Y., Ohlrogge J.B. (2008). Building lipid barriers: Biosynthesis of cutin and suberin. Trends Plant Sci. 13: 236–246 [DOI] [PubMed] [Google Scholar]
  30. Quilichini T.D., Friedmann M., Samuels A.L., Douglas C.J. (2010). ATP-binding cassette transporter G26 (ABCG26) is required for male fertility and pollen exine formation in Arabidopsis. Plant Physiol., in press [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Samuels L., Kunst L., Jetter R. (2008). Sealing plant surfaces: Cuticular wax formation by epidermal cells. Annu. Rev. Plant Biol. 59: 683–707 [DOI] [PubMed] [Google Scholar]
  32. Schuck S., Prinz W.A., Thorn K.S., Voss C., Walter P. (2009). Membrane expansion alleviates endoplasmic reticulum stress independently of the unfolded protein response. J. Cell Biol. 187: 525–536 [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Suh M.C., Samuels A.L., Jetter R., Kunst L., Pollard M., Ohlrogge J., Beisson F. (2005). Cuticular lipid composition, surface structure, and gene expression in Arabidopsis stem epidermis. Plant Physiol. 139: 1649–1665 [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Sullivan D.T., Bell L.A., Paton D.R., Sullivan M.C. (1979). Purine transport by malpighian tubules of pteridine-deficient eye color mutants of Drosophila melanogaster. Biochem. Genet. 17: 565–573 [DOI] [PubMed] [Google Scholar]
  35. Sullivan D.T., Bell L.A., Paton D.R., Sullivan M.C. (1980). Genetic and functional analysis of tryptophan transport in Malpighian tubules of Drosophila. Biochem. Genet. 18: 1109–1130 [DOI] [PubMed] [Google Scholar]
  36. Sullivan D.T., Sullivan M.C. (1975). Transport defects as the physiological basis for eye color mutants of Drosophila melanogaster. Biochem. Genet. 13: 603–613 [DOI] [PubMed] [Google Scholar]
  37. Tarr P.T., Tarling E.J., Bojanic D.D., Edwards P.A., Baldán Á. (2009). Emerging new paradigms for ABCG transporters. Biochim. Biophys. Acta 1791: 584–593 [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Tiwari S., Wang S., Hagen G., Guilfoyle T.J. (2006). Transfection assays with protoplasts containing integrated reporter genes. Methods Mol. Biol. 323: 237–244 [DOI] [PubMed] [Google Scholar]
  39. Toufighi K., Brady S.M., Austin R., Ly E., Provart N.J. (2005). The Botany Array Resource: e-Northerns, expression angling, and promoter analyses. Plant J. 43: 153–163 [DOI] [PubMed] [Google Scholar]
  40. Ukitsu H., et al. (2007). Cytological and biochemical analysis of COF1, an Arabidopsis mutant of an ABC transporter gene. Plant Cell Physiol. 48: 1524–1533 [DOI] [PubMed] [Google Scholar]
  41. Verrier P.J., et al. (2008). Plant ABC proteins: A unified nomenclature and updated inventory. Trends Plant Sci. 13: 151–159 [DOI] [PubMed] [Google Scholar]
  42. Whitlock M., Schluter D. (2009). The Analysis of Biological Data. (Greenwood Village, CO: Roberts and Co. Publishers; ). [Google Scholar]
  43. Xu J., Yang C., Yuan Z., Zhang D., Gondwe M.Y., Ding Z., Liang W., Zhang D., Wilson Z.A. (2010). The ABORTED MICROSPORES regulatory network is required for postmeiotic male reproductive development in Arabidopsis thaliana. Plant Cell 22: 91–107 [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Yang H., Murphy A.S. (2009). Functional expression and characterization of Arabidopsis ABCB, AUX 1 and PIN auxin transporters in Schizosaccharomyces pombe. Plant J. 59: 179–191 [DOI] [PubMed] [Google Scholar]
  45. Yong Z., Kotur Z., Glass A.D. (2010). Characterization of an intact two-component high-affinity nitrate transporter from Arabidopsis roots. Plant J. 63: 739–748 [DOI] [PubMed] [Google Scholar]
  46. Zhang Q., Blaylock L.A., Harrison M.J. (2010). Two Medicago truncatula half-ABC transporters are essential for arbuscule development in arbuscular mycorrhizal symbiosis. Plant Cell 22: 1483–1497 [DOI] [PMC free article] [PubMed] [Google Scholar]
  47. Zheng H., Rowland O., Kunst L. (2005). Disruptions of the Arabidopsis Enoyl-CoA reductase gene reveal an essential role for very-long-chain fatty acid synthesis in cell expansion during plant morphogenesis. Plant Cell 17: 1467–1481 [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supplemental Data
supp_22_9_3066__index.html (653B, html)
supp_tpc.110.077974_Samuels_Supplemental_Data_Final.pdf (2.7MB, pdf)

Articles from The Plant Cell are provided here courtesy of Oxford University Press

RESOURCES