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. Author manuscript; available in PMC: 2008 Jan 11.
Published in final edited form as: Plant J. 2007 Nov 12;53(2):275–286. doi: 10.1111/j.1365-313X.2007.03339.x

Subcellular co-localization of Arabidopsis RTE1 and ETR1 supports a regulatory role for RTE1 in ETR1 ethylene signaling

Chun-Hai Dong 1,†, Maximo Rivarola 1,†, Josephine S Resnick 1,†, Benjamin D Maggin 1, Caren Chang 1,*
PMCID: PMC2194639  NIHMSID: NIHMS36995  PMID: 17999643

Summary

Ethylene is an important plant growth regulator perceived by membrane-bound ethylene receptors. The ETR1 ethylene receptor is positively regulated by a predicted membrane protein, RTE1, based on genetic studies in Arabidopsis. RTE1 homologs exist in plants, animals and protists, but the molecular function of RTE1 is unknown. Here, we examine RTE1 expression and subcellular protein localization in order to gain a better understanding of RTE1 and its function in relation to ETR1. Arabidopsis plants transformed with the RTE1 promoter fused to the β-glucuronidase (GUS) reporter gene revealed that RTE1 expression partly correlates with previously described sites of ETR1 expression or sites of ethylene response, such as the seedling root, root hairs and apical hook. RTE1 transcript levels are also enhanced by ethylene treatment, and reduced by the inhibition of ethylene signaling. For subcellular localization of RTE1, a functional RTE1 fusion to red fluorescent protein (RFP) was expressed under the control of the native RTE1 promoter. Using fluorescence microscopy, RTE1 was observed primarily at the Golgi apparatus and partially at the endoplasmic reticulum (ER) in stably transformed Arabidopsis protoplasts, roots and root hairs. Next, a functional ETR1 fusion to a 5xMyc epitope tag was expressed under the control of the native ETR1 promoter. Immunohistochemistry of root hairs not only showed ETR1 residing at the ER as previously reported, but revealed substantial localization of ETR1 at the Golgi apparatus. Lastly, we demonstrated the subcellular co-localization of RTE1 and ETR1. These findings support and enhance the genetic model that RTE1 plays a role in regulating ETR1.

Keywords: RTE1, ETR1, ethylene receptor, Golgi, localization, Arabidopsis

Introduction

Ethylene is a gaseous plant hormone that plays an important role in plant growth, development and responses to environmental stresses (Abeles et al., 1992). Responses to ethylene include promotion of fruit ripening, seed germination, root hair formation, flowering, abscission and senescence. At the cellular level, the ethylene-response pathway is initiated by a family of ethylene receptors, which have similarity to the two-component histidine protein kinase family, and which signal through a series of proteins that lead to changes in gene expression (Chen et al., 2005; Li and Guo, 2007).

Arabidopsis has five ethylene receptors (ETR1, ERS1, EIN4, ETR2 and ERS2) (Chang et al., 1993; Hua et al., 1995, 1998; Sakai et al., 1998), which are negative regulators of ethylene responses (Hall and Bleecker, 2003; Hua and Meyerowitz, 1998; Qu et al., 2007). The receptors consist of an N-terminal membrane-bound region containing the ethylene-binding pocket, followed by a GAF-like domain and a histidine protein kinase-like domain. ETR1, EIN4 and ETR2 also carry a C-terminal receiver domain. The receptors fall into two subfamilies based on structural similarities (Chen et al., 2005). ETR1 and ERS1 comprise subfamily I, with three transmembrane domains followed by a highly conserved histidine kinase domain. EIN4, ETR2 and ERS2 comprise subfamily II, with four N-terminal transmembrane domains and a degenerate histidine kinase domain. Although the five receptors have functional redundancy, subfamily I has a stronger effect than subfamily II in ethylene signaling (Hall and Bleecker, 2003; Qu et al., 2007).

According to the current model for ethylene receptor action, the receptors repress responses when ethylene is not bound, and are turned off when ethylene is bound, resulting in the activation of responses (Chen et al., 2005). The receptors have been shown to bind ethylene (O’Malley et al., 2005; Schaller and Bleecker, 1995) with the help of a copper co-factor Cu(I) (Rodriguez et al., 1999), which requires RAN1, a homolog of the Golgi-bound Menkes/Wilson P-type ATPase copper transporter in mammals (Hirayama et al., 1999; Woeste and Kieber, 2000). The mechanism of ethylene receptor signaling is unknown. ETR1 and ERS1 display histidine autokinase activity (Gamble et al., 1998; Moussatche and Klee, 2004), while the subfamily II receptors, plus ERS1, exhibit autophosphorylation on serine residues (Moussatche and Klee, 2004). However, kinase activity does not appear to play a significant role in ethylene receptor signaling (Gamble et al., 2002; Wang et al., 2003).

The REVERSION-TO-ETHYLENE SENSITIVITY1 (RTE1) gene was recently identified as a positive regulator of ETR1 signal transmission (Resnick et al., 2006). RTE1 encodes a novel predicted membrane protein with no sequence similarity to proteins of known function. RTE1 homologs are found in plants, animals and protists, but currently the only ascribed function for RTE1 is in ethylene signaling in plants (Barry and Giovannoni, 2006; Resnick et al., 2006). Mutants of rte1 were isolated based on their ability to suppress the ethylene insensitivity of the gain-of-function mutant etr1–2. rte1 does not suppress the gain-of-function allele etr1–1, nor does it suppress gain-of-function alleles of the four other ethylene receptor genes, suggesting that RTE1 regulation is specific for the ETR1 receptor and is likely to occur at the protein level (Resnick et al., 2006). The rte1 null mutant displays ethylene hypersensitivity that phenocopies the etr1–7 null mutant, and RTE1 is thought to act in the same pathway as ETR1, as the etr1–7 rte1–2 double mutant is indistinguishable from the etr1–7 and rte1–2 single mutants (Resnick et al., 2006). Over-expression of RTE1 confers reduced ethylene sensitivity that is largely dependent on the ETR1 locus (Resnick et al., 2006; Zhou et al., 2007). Similarly, over-expression of the tomato RTE1 homolog, GREEN-RIPE (GR), confers ethylene insensitivity in tomato (Barry and Giovannoni, 2006).

Less is known about ethylene signaling at the cell biological level. Membrane fractionation studies have placed the ETR1 receptor at the endoplasmic reticulum (ER) (Chen et al., 2002). The Raf-like kinase CTR1, which is the next downstream component in the ethylene-response pathway (Clark et al., 1998; Kieber et al., 1993), is recruited to the ER through physical interaction with the ethylene receptors (Gao et al., 2003). RTE1, on the other hand, has been co-localized with a marker at the Golgi apparatus (in onion epidermal cells) (Zhou et al., 2007). Sequence analyses predict that Arabidopsis RTE1 is an integral membrane protein carrying between two and four transmembrane domains (aramemnon plant membrane protein database: http://aramemnon.botanik.uni-koeln.de/index.ep). Both RTE1 and ETR1 lack an obvious signal sequence for the secretory pathway, and there are no clear predictions of subcellular location for RTE1, RTE1 homologs or ETR1.

In order to gain a better understanding of RTE1 and its function in relation to the ETR1 receptor, we analyzed the gene expression pattern of RTE1 and determined the subcellular localization of the RTE1 protein in Arabidopsis. We show here that RTE1 expression is generally correlated with sites of ETR1 expression and ethylene response. We also demonstrate that the RTE1 protein is localized primarily at the Golgi apparatus and partially at the ER. In the course of this study, we found that the ETR1 receptor localizes not only at the ER as previously reported, but also at the Golgi. By examining RTE1 and ETR1 simultaneously, we show that they exhibit subcellular co-localization. These findings provide cell biological data in support of the model that RTE1 plays a role in regulating ETR1 ethylene signaling.

Results

RTE1 expression patterns in Arabidopsis

Gene array data indicate that Arabidopsis RTE1 is expressed at detectable levels in most organs and stages, and is up-regulated by ethylene (Alonso et al., 2003; Resnick et al., 2006; http://bbc.botany.utoronto.ca/efp/cgi-bin/efpWeb.cgi). Based on gene arrays, RTE1 is most highly expressed in developing seeds and young siliques, with high expression also seen in seedlings and the shoot apex. Using a luciferase reporter, Zhou et al. 2007 observed expression of RTE1 in cotyledons, leaves, the rachis and flowers, with lower expression seen in the seedling root and hypocotyl.

To further examine RTE1 gene expression, we fused the RTE1 promoter region (consisting of a 2.5 kb genomic DNA fragment just upstream of the RTE1 translation start codon and including the 5′ UTR of RTE1) with the β-glucuronidase (GUS) reporter gene, and transformed the resulting construct into wild-type Arabidopsis plants by Agrobacterium infiltration. The expression pattern of RTE1 was observed by staining for GUS activity in the transgenic lines. As shown in Figure 1, there is strong expression of RTE1 in 1–4-day-old seedlings in the apical hook, cotyledons, root vascular tissue, root tip and root hairs, with little or no expression in the hypocotyl. The pattern of expression was similar in dark- and light-grown seedlings (Figure 1a–d). In light-grown seedlings, expression could also be seen in the apex and young leaves (Figure 1d,g,i), and disappeared from the cotyledons by 10 days (Figure 1i). In mature plants, RTE1 was expressed in floral buds (Figure 1j), the style of mature flowers (Figure 1k), stems and the rachis (not shown).

Figure 1.

Figure 1

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RTE1promoter–GUSexpression patterns. (a–e,g–k)Representative GUS expression is seen in the following wild-type tissues: (a) cotyledons, apical hook and root of a 1-day-old dark-grown seedling; (b) cotyledons, apical hook and root of a 3-day-old dark-grown seedling; (c) cotyledons and root of a 1-day-old light-grown seedling; (d) cotyledons, root and shoot apex of a 3-day-old light-grown seedling; (e) vascular tissue and the root tip of a 3-day-old dark-grown seedling; (g) cotyledons and shoot apex of a 3-day-old lightgrown seedling; (h) root, including root hairs, of a 3-day-old light-grown seedling; (i) developing leaves and roots of a 10-day-old light-grown seedling; (j) floral buds; (k) style of mature flower. (f) No expression is detected in the hypocotyl of a 3-day-old dark-grown seedling.

(l–w) Representative 4-day-old dark-grown seedlings subjected to various treatments: (l) no treatment; (m) root (close-up) of seedling with no treatment; (n) germinated on medium containing 100 μM ACC; (o) root (close-up) of seedling germinated on medium containing 100 μM ACC; (p) hypocotyl (close-up) of seedling germinated on medium containing 100 μM ACC; (q) root tip (close-up) of seedling germinated on medium containing 100 μM ACC; (r) germinated on medium containing 10 μM AgNO3 (an inhibitor of ethylene response); (s) root of seedling grown on medium containing 10 μM AgNO3, showing weak expression at the root tip; (t) etr1–1 seedling with no treatment; (u) etr1–1 seedling root, showing weak expression at the root tip; (v) etr1–1 seedling hypocotyl (close-up), showing no detectable expression; (w) etr1–1 seedling cotyledons, showing no detectable expression.

Scale bars = 1 mm in (a–d,i–l,n,o,r–u,w) and 100 μm in (e–h,m,p,q,v).

To analyze the effect of ethylene treatment on RTE1 expression, we examined the GUS staining pattern of etiolated seedlings germinated in the presence of the ethylene precursor 1-aminocyclopropane-1-carboxylic acid (ACC). We also examined the effect of AgNO3 (an inhibitor of the ethylene response; Beyer, 1976), as well as the effect on expression when the reporter construct was transformed into the ethylene-insensitive etr1–1 mutant background. Etiolated seedlings responding to ethylene or ACC treatment display the ‘triple response’ phenotype, which consists of shortening and radial swelling of the hypocotyl, inhibition of root growth, proliferation of root hairs and an exaggerated apical hook, whereas seedlings treated with AgNO3 or in the etr1–1 background have longer hypocotyls and roots (Bleecker et al., 1988). Based on a qualitative assessment of GUS staining, RTE1 transcript levels in comparison to untreated seedlings were enhanced by ACC and reduced by AgNO3 and etr1–1, although the pattern itself was unaltered by the treatments (Figure 1l–w). These results suggest that there is a negative feedback mechanism in ethylene signaling via regulation of RTE1 expression, as RTE1 is a negative regulator of ethylene responses.

Localization of RFP–RTE1 at the Golgi apparatus and ER in Arabidopsis protoplasts

Using antiserum raised against an RTE1 peptide, we detected RTE1 in the microsomal fraction of protein extracts from Arabidopsis seedlings, confirming the prediction that RTE1 is localized at the membrane (data not shown). It was not feasible, however, to use this antiserum to determine the subcellular localization of RTE1, because Western blots showed cross-reaction with numerous nonspecific bands. In order to detect the RTE1 protein, we therefore created a reporter fusion to RTE1. To construct the reporter, the open reading frame of red fluorescence protein (RFP) was fused in-frame to the RTE1 sequence at the N- or C-terminus of RTE1 (Figure 2a). Each fusion was placed under the control of the native RTE1 promoter region (identical to that used in the GUS expression analysis) and cloned into the binary transformation vector pMLBart to create RFP–RTE1 and RTE1–RFP, respectively. To test the functionality of RFP–RTE1 and RTE1–RFP in planta, the constructs were stably transformed into the double mutant etr1–2 rte1–3 to test for rescue of the rte1–3 mutation. Five independent transgenic lines of each construct were examined for the ability to restore ethylene insensitivity in etr1–2 rte1–3 (by alleviating rte1–3 suppression of etr1–2). For the RFP–RTE1 construct, all five lines exhibited substantial rescue of the rte1–3 mutation based on the seedling triple response, indicating that RTE1 carrying an N-terminal RFP tag retains substantial function (Figure 2b). For the other construct, none of the five lines rescued rte1–3 to the same extent as the RFP–RTE1 construct (Figure 2b). Therefore, all subsequent analyses of the RTE1 protein were performed using RFP–RTE1, in which RFP is fused at the N-terminus of RTE1.

Figure 2.

Figure 2

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RFP-tagged RTE1 rescues the Arabidopsis rte1–3 null mutation. (a) Diagram of RFP-tagged RTE1 constructs carrying RFP (red) at either the Nterminus or C-terminus of the RTE1 coding sequence (yellow), which was amplified from genomic DNA. Both constructs are driven by the native RTE1 promoter region and include the native RTE1 terminator sequence. The gray portions represent the 5′ and 3′ UTRs of RTE1. A small portion of the 3′ UTR of each flanking gene in the genome (blue) is also present in each construct. Arrows indicate the direction of transcription.

(b) Representative 4-day-old dark-grown seedlings germinated on 50 μM ACC, showing that RFP–RTE1 largely rescues the rte1–3 mutation in the etr1–2 rte1–3 double mutant, while RTE1–RFP does not. Scale bar = 2 mm.

To determine the intracellular localization of RTE1, protoplasts were prepared from wild-type plants that had been stably transformed with the RFP–RTE1 construct, and then various GFP-tagged organelle markers were transiently expressed in the protoplasts to compare their localization patterns with those of RFP–RTE1. The markers were: GFP– HDEL (ER), ST–GFP (Golgi), GmMan1(tTMsC)–GFP (cis- Golgi), GFP–δTIP (vacuole), GFP–CPK9 (plasma membrane), GFP–SKL (peroxisome), COX4ts–GFP (mitochondria) and cp targeting signal–YFP (plastid). Using confocal laser scanning microscopy, we examined at least 20 protoplasts expressing each marker. Substantial co-localization of RFP–RTE1 with the Golgi and cis-Golgi markers was detected, in addition to partial co-localization with the ER marker (Figure 3a–c). There was potentially some slight co-localization with the vacuole marker (Figure 3d). In contrast, we did not detect co-localization of RFP–RTE1 with GFP-tagged markers for the plasma membrane, peroxisome, mitochondria or plastid (Figure 3e–h). These results suggest that RTE1 in protoplasts localizes primarily to the Golgi apparatus and partially to the ER.

Figure 3. Localization of RFP–RTE1 at the Golgi apparatus and ER in Arabidopsis protoplasts.

Figure 3

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Confocal laser scanning microscopy images of representative protoplasts showing fluorescence of various organelle markers (left panel), RFP–RTE1 fluorescence (middle panel), and merged images of the left and middle panels (right panel).

(a–h) RFP–RTE1 localization is compared with that of the following markers: (a) GFP–HDEL (ER); (b) ST–GFP (Golgi); (c) GmMan1(tTMsC)–GFP (cis-Golgi); (d) GFP–δTIP (vacuole); (e) GFP–CPK9 (plasma membrane); (f) GFP–SKL (peroxisome); (g) COX4ts–GFP (mitochondria); (h) cp targeting signal–YFP (plastid).

Scale bars = 10 μm.

Localization of RFP–RTE1 at the Golgi apparatus and ER in Arabidopsis root and root hair cells

In order to confirm the Golgi and ER localization, we examined the localization of RFP–RTE1 in planta. For this, we stably transformed GFP–HDEL (ER) and ST–GFP (Golgi) into wild-type Arabidopsis, and then crossed the resulting transformants to the above RFP–RTE1 transformants. The F1 were allowed to self-pollinate, and several F2 progeny were analyzed for the presence of both RFP–RTE1 and the ER or Golgi marker. We examined fluorescence in seedling root cells, which were easy to access and because GUS staining had been pronounced in the seedling root. Confocal laser scanning microscopy showed that RFP–RTE1 co-localized well with the Golgi marker in both root cells and root hair cells, consistent with the Golgi apparatus localization in protoplasts (Figure 4a,c). There also appeared to be partial co-localization with the ER marker. The RFR–RTE1 signal was more punctate (thus more Golgi-like) than that of the ER marker (Figure 4b,d).

Figure 4. Localization of RFP–RTE1 at the Golgi apparatus and ER in root and root hair cells of Arabidopsis.

Figure 4

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Representative images from confocal laser scanning microscopy showing fluorescent markers (left panel), RFP–RTE1 fluorescence (middle panel) and merged images (right panel) from stably transformed plants expressing RFP–RTE1 and the corresponding organelle marker.

(a,b) Root cells of 4-day-old light-grown seedlings expressing both ST–GFP (Golgi) and RFP–RTE1.

(c) Root hair cell of a 4-day-old light-grown seedling expressing both ST–GFP (Golgi) and RFP–RTE1.

(d,e) Root cells of 4-day-old light-grown seedlings expressing both GFP–HDEL (ER) and RFP– RTE1.

(f) Root hair cell of 4-day-old light-grown seedling expressing both GFP–HDEL (ER) and RFP– RTE1.

Scale bars = 10 μm.

To test whether ethylene affects the subcellular localization of RTE1, we compared the localization pattern of RFP–RTE1 with that of the GFP–HDEL and ST–GFP markers in roots of 2–4-day-old etiolated seedlings germinated in the presence and absence of the ethylene precursor ACC. No change was detected in the localization of RFP–RTE1 resulting from ACC treatment (data not shown).

Localization of ETR1–5xMyc at the Golgi apparatus and ER in Arabidopsis root hair cells

Next, we were interested in investigating whether ETR1 and RTE1 share similar subcellular localization. To visualize the localization of RTE1 and ETR1, we initially prepared protoplasts from the leaves of RFP–RTE1 transgenic plants and transfected the protoplasts with a construct that expressed ETR1 fused to a C-terminal CFP tag. The ETR1–CFP fusion yielded only weak fluorescence, however (data not shown). To increase the sensitivity for detection of ETR1, we switched to a 5xMyc epitope tag fused at the C-terminus of ETR1. The ETR1–5xMyc fusion was expressed under the control of the native ETR1 promoter region (comprising 3.2 kb upstream of the ETR1 translation start site) (Figure 5a). We determined that the ETR1–5xMyc fusion construct possessed wild-type ETR1 activity by transforming it into the etr1 etr2 ein4 triple null mutant, which has a constitutive ethylene-response phenotype (Huo and Meyerowitz, 1998). We examined five independent transgenic lines, and found that the triple null phenotype was rescued to the less severe etr2 ein4 double null mutant phenotype (Figure 5b). In all subsequent analyses with the ETR1 protein, the ETR1– 5xMyc construct was stably transformed into either the etr1–7 null mutant or wild-type plants.

Figure 5. Function and detection of the ETR1 receptor fused with an epitope tag (5xMyc).

Figure 5

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(a) Diagram of the Myc epitope (5xMyc)-tagged ETR1 construct driven by the native ETR1 promoter region. Shown are the promoter region, which includes a small portion of the flanking gene in the genome (light blue), the ETR1 5′ UTR (gray) with native intron, the ETR1 coding sequence (dark blue), the 5xMyc epitope translational fusion (orange) and 3′ OCS terminator. Arrows indicate the direction of transcription.

(b) Rescue of the etr1–7 null mutation in the Arabidopsis triple receptor null mutant (etr1–6 etr2–3 ein4–4) by ETR1–5xMyc. Representative 4-day-old darkgrown seedlings in air (no ethylene treatment) show that ETR1–5xMyc rescues the etr1–6 mutation, alleviating the constitutive triple response and restoring the triple mutant to the etr2–3 ein4–4 double null phenotype. Scale bar = 2 mm.

(c) Western blot showing the intact ETR1–5xMyc monomer isolated from the microsomal membrane fraction of Arabidopsis seedlings run on denaturing PAGE and detected by an anti-c-myc antibody. ETR1–5xMyc transformed into etr1–7 gives a predominant band of approximately 80 kDa (left lane), which is absent in the untransformed wild-type (right lane). A non-specific band of lower molecular weight is detected in both samples. ECA1, an ER-membrane protein (Liang et al., 1997), was used as a loading control.

To ensure that the ETR1–5xMyc fusion protein was intact in the transformed lines, we isolated protein from the transformed plants and visualized the protein on a Western blot using an anti-Myc antibody. The results consistently showed a single band of the correct monomer size on a Western blot of a denaturing PAGE gel (Figure 5c). There was also a single band of the predicted molecular weight for the dimer under non-denaturing conditions (data not shown). Occasionally, a single non-specific background band of smaller size was detected, whether or not the samples carried the ETR1–5xMyc protein (Figure 5c).

Given that the anti-Myc antibody detected an intact fusion protein, we proceeded with immunohistochemistry of root hair cells of plants that had been stably transformed with ETR1–5xMyc. The GFP–HDEL and ST–GFP marker constructs (used above) were transformed into the ETR1–5xMyc lines to generate separate lines expressing ETR1–5xMyc with each marker. Notably, substantial co-localization of ETR1–5xMyc was observed with the Golgi marker (Figure 6a), and partial co-localization was also seen with the ER marker (Figure 6b). No signal was observed in root hair cells of untransformed seedlings that were fixed and treated in parallel with the anti- Myc antibody, indicating that the background band seen in the Western blot was not detected by this method (data not shown).

Figure 6. Localization of ETR1–5xMyc at the Golgi apparatus and ER in Arabidopsis root hair cells.

Figure 6

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Representative root hair cells viewed by confocal laser scanning microscopy.

(a) Root hair cell of a 5-day-old light-grown seedling expressing both ST–GFP (Golgi) and ETR1–5xMyc, visualized by immunohistochemistry using an antic-myc antibody.

(b) Root hair cell of a 5-day-old light-grown seedling expressing both GFP– HDEL (ER) and ETR1–5xMyc, visualized by immunohistochemistry using an anti-c-myc antibody.

Scale bars = 10 μm.

Co-localization of RTE1 and ETR1 in Arabidopsis root hair cells

Finally, we examined whether RTE1 co-localizes with the ETR1 receptor. To obtain Arabidopsis plants harboring both RFP–RTE1 and ETR1–5xMyc, we crossed the individual transformants (above) together, and allowed the resulting F1 to self-pollinate to produce F2 seeds. Whole roots from 4-day-old F2 seedlings were analyzed by immunohistochemistry using a monoclonal anti-Myc antibody. The immunolocalization of ETR1–5xMyc, as well as RFP fluorescence from RFP–RTE1, was viewed in root hair cells by confocal laser scanning microscopy. As shown in Figure 7, co-localization of RTE1 and ETR1 was observed. No antibody signal was detected in root hair cells of approximately one fourth of the segregating F2s indicating the absence of non-specific background signal for this method (data not shown).

Figure 7. Co-localization of RTE1 and ETR1 in Arabidopsis root hair cells.

Figure 7

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Representative root hair cells viewed by confocal laser scanning microscopy. (a–c) Three root hair cells of 5-day-old light-grown seedlings expressing both ETR1–5xMyc and RFP–RTE1. RFP–RTE1 is visualized by fluorescence (as in Figure 4) and ETR1–5xMyc is visualized by immunohistochemistry (as in Figure 6). Scale bars = 10 μm.

Discussion

Previous genetic analyses have indicated that Arabidopsis RTE1 is a positive regulator of ETR1 ethylene receptor function (Resnick et al., 2006; Zhou et al., 2007). In this paper, we advance the understanding of RTE1 and ETR1 function at the cell biological level, providing data that support and enhance the genetic model.

GUS reporter analysis of the RTE1 promoter revealed that RTE1 has discrete and specific expression patterns, some of which can be correlated with sites of ETR1 expression and ethylene response. RTE1 is strongly expressed in the seedling apical hook, root tip and root hairs – all cells that are linked to ethylene-inducible rapid cell division and/or cell elongation (Dolan, 2001; Ortega-Martínez et al., 2007; Raz and Koornneef, 2001). While RTE1 shows little or no expression in the hypocotyl, the hypocotyl is derived from cells that have passed through the apical hook (Raz and Ecker, 1999) where RTE1 expression is high. RTE1 is also expressed in developing leaves, young cotyledons, stems, rachis and style. The RTE1 expression pattern partly overlaps with the pattern of expression of the ETR1 receptor gene, as detected by in situ hybridization in etiolated seedlings (Hua et al., 1998; Raz and Ecker, 1999), although ETR1 expression is higher in the hypocotyl and weaker in the apical hook in 2- and 3-day-old seedlings (Raz and Ecker, 1999). ETR1 is also expressed in stems and leaves, and in the locules of anthers, developing carpels and ovules (Hua et al., 1998). Unlike ETR1 expression, which is not ethyleneinduced (Hua et al., 1998), RTE1 expression is enhanced upon ethylene treatment and reduced when ethylene signaling is blocked, suggesting a mechanism of negative feedback on the response pathway. The ethylene-enhanced expression that we observed is consistent with array data indicating that exposure to ethylene results in a fourfold increase in RTE1 transcript levels (Alonso et al., 2003), as also seen in RNA blots (Resnick et al., 2006). These findings, showing that RTE1 is expressed preferentially at several important sites for ethylene response, and that expression is responsive to ethylene, are consistent with RTE1 having a regulatory role in ethylene signaling.

The RTE1 protein was visualized in living Arabidopsis cells (protoplasts, root cells and root hair cells) using the RTE1 native promoter and a red fluorescent protein tag. We found that RTE1 is localized predominantly at the Golgi apparatus and partially at the ER. We do not rule out the possibility of a small amount of RTE1 localization at the vacuole, based on the examination of protoplasts co-expressing RFP-RTE1 and a vacuole marker. The ER is one of the major components of the endomembrane system, closely connected with the Golgi apparatus and vacuoles (Hawes and Satiat-Jeunemaitre, 2005). There was no obvious localization of RTE1 at the plasma membrane, peroxisome, mitochondrion or plastid organelles. In addition, we did not detect any alteration in the subcellular localization of RTE1 when the seedlings were treated with ethylene, consistent with the findings of Zhou et al. 2007, who showed that a CaMV 35S-driven GFPtagged RTE1 fusion was localized at the Golgi apparatus in onion epidermal cells.

Interestingly, we found that the ETR1 receptor is localized primarily at the Golgi apparatus and partially at the ER in Arabidopsis root hair cells. Chen et al. 2002 previously reported localization of ETR1 at the ER, but did not rule out the possibility of Golgi localization; ER localization was based on the co-fractionation of ETR1 and an ER marker by sucrose density gradient centrifugation, in which the Golgicontaining fractions showed a similar, but slightly broader distribution, than that of the ER fractions (Chen et al., 2002, 2007). The ER and Golgi fractions exhibited the same shift from higher to lower density in the absence of Mg2+, indicating that the ER and Golgi apparatus are not easily resolved by this method (Chen et al., 2002, 2007). There are known structural and functional links between the ER and Golgi apparatus, and, in fact, a continuum between the ER and Golgi apparatus has been proposed by Hawes and Satiat-Jeunemaitre (2005). Conceivably, ETR1 is differentially localized depending on the stage or type of cell, thus yielding different results depending on the cell types examined. For the previously published sucrose density gradient centrifugation, protein was extracted from plants grown in liquid culture, containing predominantly green tissue. Additionally, leaf cells were examined by immunoelectron microscopy, which again did not rule out the possibility of ETR1 localized at the Golgi apparatus (Chen et al., 2002). In the study presented here, the ETR1–5xMyc fusion was localized by immunohistochemistry of intact root hair cells. Although the 5xMyc epitope tag could potentially lead to artifacts, the ETR1–5xMyc construct was able to rescue an etr1 null mutation, and the ETR1–5xMyc fusion protein was seen in Western blots as an intact band of the expected molecular weight. There was no distinct background signal detected by immunofluorescence microscopy of root hair cells, even though a single faint band was occasionally detected in Western blots whether or not the plants carried the ETR1–5xMyc construct.

Localization of ETR1 at the Golgi apparatus presents an interesting modification to our current understanding of ethylene receptor signaling, but is consistent with the overall model of ethylene signaling. Due to the solubility of ethylene in aqueous and lipid environments, ethylene should be readily perceived by receptors residing at either organelle (Abeles et al., 1992). The receptors require a copper co-factor in order to bind ethylene (Rodriguez et al., 1999), and it is believed that this copper is delivered by RAN1 (Hirayama et al., 1999; Woeste and Kieber, 2000). RAN1 is a homolog of the mammalian Menkes/Wilson P-type ATPase copper transporter, which has been localized (in mammals) at the Golgi membrane and delivers copper to the lumen (Petris et al., 1996). If RAN1 is similarly localized at the Golgi apparatus in plants, then copper could be directly supplied to the Golgi-associated ETR1 receptor, providing a cell biological link between RAN1 and ethylene receptor signaling. Another possible connection is the fact that certain ethylene-induced responses require Golgi-specific functions, thereby associating ETR1 and RTE1 with a site of ethylene response. For example, cell-wall synthesis is required for the processes of cell elongation and expansion, which occur in certain responses to ethylene, such as at the apical hook and in root hair elongation. The components for cell-wall synthesis are produced at the Golgi apparatus (Lerouxel et al., 2006), and thus the regulation of these processes by ethylene could involve co-localization of ETR1 and RTE1 with components in the Golgi apparatus, in a manner similar to that proposed by Chen et al. 2005 for ERlocalized ethylene receptors. Not all ethylene receptors may be localized at the ER or Golgi apparatus. ETR2 (Chen et al., 2007) and the melon ethylene receptor CmERS1 (subfamily I) have been localized to the ER using sucrose density gradient fractionation (Ma et al., 2006), but tobacco NTHK1 (subfamily II) appears to localize at the plasma membrane (PM) (Xie et al., 2003), and unpublished work by Klee and Tieman (University of Florida) suggests that tomato NEVERRIPE (subfamily I) may also be localized at the PM.

The subcellular co-localization of RTE1 and ETR1 supports the possibility that RTE1 promotes ETR1 signaling through physical interaction with ETR1. Whether a physical interaction occurs between these proteins is currently under investigation. If RTE1 acts directly on ETR1, then RTE1 might serve as a molecular chaperone or co-factor for ETR1, or affect the membrane trafficking or stability of ETR1. Alternatively, RTE1 could exert an indirect effect, such as altering the conformation of ETR1 via changes to the membrane or other proteins, or changes in the status of copper. If the other ethylene receptors in Arabidopsis prove to be localized primarily to other tissues or membranes relative to RTE1, then co-localization with ETR1 might be an underlying basis for the specificity of RTE1 for ETR1. Differential tissue localization of ethylene receptors has been postulated for the non-global ethylene effects of GR over-expression in tomato (Barry and Giovannoni, 2007). Further insight into the connections between RTE1 and ETR1 should advance our understanding of the basis for RTE1’s regulation of, and specificity for, the ETR1 receptor in ethylene signaling.

Experimental procedures

Plant growth and transformation

Arabidopsis thaliana plants [ecotype Columbia (Col-0) were grown in soil under a 16 h light/8 h dark photoperiod in a controlled environment chamber at 20°C under white fluorescent light 100 μmol m−2 s−1. For seedling growth, seeds were sown on MS plates containing 0.8% agar. After stratification for 3 days at 4°C, the seeds were incubated at 20°C either under continuous light or in the dark for the indicated lengths of time. Transgenic plants were generated by the floral dip infiltration method (Clough and Bent, 1998) mediated by Agrobacterium tumefaciens strain GV3101. To select for transformed plants, we used either hygromycin (250 mg l−1) or Basta (0.1% Finale™ [Bayer Crop Science; http://www.bayercropscience.com] sprayed onto seedlings), depending on the binary vector used.

The triple response assay was performed as described previously (Resnick et al., 2006) using the stated concentrations of ACC or AgNO3 in the medium.

Construction of RTE1 and ETR1 reporter fusions

To construct the RTE1promoter–GUS fusion, a DNA fragment containing the RTE1 promoter region (2485 bp upstream from the RTE1 start codon, which includes the intron located in the RTE1 5′ UTR) was PCR-amplified from Arabidopsis wild-type genomic DNA using the primers 5′-GGATGATGTGATCACCATCG-3′ and 5′-TTTTAGATTCCTAATCACACAAGAC-3′. The PCR product was cloned into the pCR8/GW/TOPO TA cloning plasmid vector (Invitrogen, http://www.invitrogen.com/) and verified by nucleotide sequencing. Using the Gateway recombination system (Invitrogen), the RTE1 promoter region was inserted upstream of the GUS reporter gene in binary vector pBGWFS7 (Karimi et al., 2002).

To generate constructs encoding RFP-tagged RTE1 fusion proteins, the RFP coding sequence was PCR-amplified from pDSRed2- C1 kindly provided by Biao Ding (Ohio State University). For the eventual fusion of RFP at the N-terminus of RTE1, primers 5′-CCTAGGATGGCCTCCTCCGAGAACGTC-3′ and 5′-GCTAGCTCTAGATCCGGTGGATCCCGG- 3′ were used to PCR-amplify the RFP coding sequence, eliminating the stop codon and incorporating an AvrII restriction site at the 5′ end and a BmtI restriction site at the 3′ end. For the eventual fusion of RFP at the C-terminus of RTE1, primers 5′-CCTAGGATGGCCTCCTCCGAGAACGTC-3′ and 5′-GCTAGCTTATCTAGATCCGGTGGATCC- 3′ were used to PCR-amplify the RFP coding sequence, incorporating an AvrII restriction site at the 5′ end and a BmtI restriction site at the 3′ end. The resulting fragments were cloned into pGEM-T Easy (Promega, http://www.promega.com/) and verified by nucleotide sequencing. The clones were designated pGEM-NT-RFP and pGEM-CT-RFP, respectively.

For the N-terminal RFP tag on RTE1, primers 5′-CCTAGGTTGGATGATGTGATCACCATCG- 3′ and 5′-CCTAGGTTTTAGATTCCTAATCACACAAGAC- 3′ were used to PCR-amplify a 2487 bp fragment from genomic DNA encompassing the RTE1 promoter region and 5′ UTR, incorporating flanking AvrII restriction sites. Primers 5′-GCTAGCATGTCACGTGGAAGAGGAGTTCC-3′ and 5′-GCTAGCTCACTGTTGGTACAACTTTGTGG-3′ were used to PCRamplify a 1298 bp fragment from genomic DNA encompassing the coding sequence of RTE1 and the terminator region, incorporating flanking BmtI restriction sites. The amplified fragments were cloned separately into pGEM-T Easy and verified by nucleotide sequencing. The fragments were released and ligated in succession into the respective AvrII and BmtI sites of pGEM-NT-RFP. For the C-terminal RFP tag on RTE1, primers 5′-CCTAGGTTGGATGATGTGATCACCATCG- 3′ and 5′-CCTAGGAGTAATTATGTTCTTAAAACAGTAAC-3′ were used to PCR-amplify a 3351 bp fragment from genomic DNA, encompassing the RTE1 promoter region through to the end of the RTE1 coding sequence, eliminating the stop codon and incorporating flanking AvrII restriction sites. Primers 5′-GCTAGCAGCAGTATGAGAGAAAT-3′ and 5′-GCTAGCTCACTGTTGGTACAACTTTGTGG-3′ were used to PCR-amplify a 431 bp fragment from genomic DNA, encompassing the 3′ UTR and terminator region of RTE1 and incorporating flanking BmtI restriction sites. These amplified fragments were cloned separately into pGEM-T Easy and verified by nucleotide sequencing. The AvrII and BmtI fragments were released and ligated in succession into the respective AvrII and BmtI sites of pGEM-CT-RFP. Finally, the composite fragments for both the C-terminal and N-terminal RFP fusions to RTE1 were released with NotI and ligated into the NotI site of the binary vector pMLBart (Gleave, 1992), which contains the bar gene for the selection of stably transformed plants.

To generate the ETR1–5xMyc construct, a 3.9 kb PstI–BstXI genomic DNA fragment containing the ETR1 promoter region (3167 bp upstream from the ETR1 start codon including the native intron located in the 5′ UTR) plus 733 bp of the ETR1 coding sequence was cloned into plasmid pBJ36 (Gleave, 1992) just upstream of the 3′ UTR OCS terminator sequence. Just downstream of this ETR1 fragment, we inserted a 1593 bp BstXI–BamHI ETR1 cDNA fragment (including the stop codon and 25 bp of the ETR1 3′ UTR). Next, a fragment of approximately 400 bp containing the 3′ end of the ETR1 coding region was PCR-amplified, replacing the stop codon with StuI–BamHI restriction sites. After digesting the fragment with both AflII (a natural internal site in the ETR1 coding sequence) and BamHI, the fragment was used to replace the AflII– BamHI fragment of the above construct (in which the BamHI site was located just after the ETR1 stop codon). An StuI–StuI DNA fragment containing five copies of the Myc epitope (5xMyc) followed by a stop codon (from clone CD3-128; Arabidopsis Biological Resource Center, Ohio State University) was then cloned in-frame into the introduced StuI site. The clone was verified by nucleotide sequencing, and then the entire composite gene including the OCS terminator was released with NotI and ligated into the NotI site of the binary vector pMLBart for stable plant transformation.

Fluorescent protein-tagged markers for organelle localization

The established fluorescent protein markers used in this study were GFP–HDEL (pVKH18En6-mGFPer) for the ER (Saint-Jore et al., 2002), ST–GFP (pVKH18En6-STtmd-GFP) for the Golgi apparatus (Saint- Jore et al., 2002), GmMan1(tTMsC)–GFP (pAN33) for the cis-Golgi (Nebenführ et al., 1999), GFP–CPK9 for the plasma membrane (Padmanaban et al., 2007), GFP–δTIP for the vacuole (Padmanaban et al., 2007), GFP–SKL (pAN81) for the peroxisome (Nelson et al., 2007), COX4ts–GFP (pAN107) for the mitochondria (Nelson et al., 2007), and cp targeting signal–YFP (pAN186) for the plastid (Nelson et al., 2007).

Transfection of plant protoplasts

Transient gene expression in Arabidopsis mesophyll protoplasts was carried out as described previously (Sheen, 2001). In brief, Arabidopsis protoplasts were isolated from the leaves of 25-day-old plants (10 h light/14 h dark, 22°C day/20°C night) that had been stably transformed with RFP–RTE1. Leaf strips were digested in a buffer containing cellulose R-10 and macerozyme R-10 (Yakult Pharmaceutical; http://www.yakult.co.jp). An equal volume of protoplasts was mixed with PEG buffer (40% PEG4000 (Sigma; http://www.sigmaaldrich.com), 20% 1 M mannitol, 10% 1 M CaCl2) after adding 60–120 μg of plasmid marker DNA, and then incubated at room temperature for 30 min. After gentle washing, the protoplasts were kept in the dark at room temperature overnight, and then viewed by confocal laser scanning microscopy as described below.

Histochemistry

GUS staining was carried out as previously described (Dong et al., 2001). Images of GUS-stained plants were obtained using a Nikon SMZ1000 dissecting microscope or a Nikon Eclipse E600 microscope (Nikon; http://www.nikon-instruments.jp) using differential interference contrast (DIC) microscopy.

For immunohistochemistry of ETR1–5xMyc, Arabidopsis seedlings were grown under white light and prepared essentially as described by Friml et al. 2003. In brief, 5-day-old light-grown seedlings were fixed in 4% paraformaldehyde in MTSB (50 mM PIPES, 5 mM EGTA, 5 mM MgSO4, pH 7, adjusted with KOH) for 1 h. Samples were washed with MTSB/0.1% Triton (5–10 min) and with de-ionized water (5–10 min). Cell walls were digested with 1% cellulase and 0.1% maceroenzyme in MTSB for 30 min, and then samples were washed with MTSB/0.1% Triton (5–10 min). Incubation with 10% DMSO/3% NP-40 in MTSB for 1 h followed. After another washing in MTSB/0.1% Triton (5–10 min), seedlings were pre-incubated in 2% BSA/MTSB (1 h at 37°C), and then incubated overnight (4°C) with the primary antibody, which was mouse monoclonal anti-c-Myc antibody (Invitrogen) at 1:200 dilution. After extensive washing with MTSB/0.1% Triton (8–10 min), the seedlings were incubated with a 1:500 dilution of the appropriate secondary antibody in 3% BSA/MTSB (3 h at 37°C). The secondary antibody for co-localization with the GFP-tagged ER and Golgi markers was Alexa Fluor 633 goat anti-mouse IgG (H+L) (Invitrogen). For co-localization with RFP–RTE1, the secondary antibody was Alexa Fluor 488 goat anti-mouse IgG (H+L) (Invitrogen). The samples were washed with MTSB/0.1% Triton (5–10 min, then overnight) and transferred into VectaShield mounting medium (Vector Laboratories; http://www.vectorlabs.com).

Fluorescence microscopy

Imaging of fluorescent proteins in protoplasts or seedling roots was conducted under a laser scanning confocal microscope (Zeiss LSM510, http://www.zeiss.com/). The excitation wavelengths for GFP (or YFP) and RFP were 488 and 543 nm, respectively, and the emission filter wavelengths were 505–530 nm for GFP 505–550 nm for YFP and 560–615 nm for RFP. Protoplasts were directly mounted on a glass slide in buffer solution (0.5 M mannitol, 4 mM MES, pH 5.7, 20 mM KCl), and seedling root fragments were mounted in water for visualization of the fluorescent proteins. For immunohistochemistry imaging of ETR1–5xMyc using Alexa Fluor 633, the excitation and emission wavelengths were 633 and 650 nm, respectively. For immunohistochemistry imaging of ETR1–5xMyc using Alexa Fluor 488, the same confocal microscopy settings were used as for GFP.

Membrane protein isolation, SDS–PAGE and Western blotting

For isolation of Arabidopsis membranes, 8-day-old etiolated seedlings were homogenized at 4°C in extraction buffer (50 mM Tris, pH 8.0, 150 mM NaCl, 10 mM EDTA and 20% v/v glycerol) containing a protease inhibitor cocktail (Sigma, http://www.sigmaaldrich.com/). The homogenate was strained through Miracloth (Calbiochem- Novabiochem; http://www.emdbiosciences.com) and centrifuged at 8000 g for 15 min. The supernatant was centrifuged at 100 000 g for 30 min, and the membrane pellet resuspended in 10 mM Tris, pH 7.5, 150 mM NaCl, 1 mM EDTA and 10% v/v glycerol with protease inhibitors. Immunoblot analysis was performed as described by Gamble et al. 2002. In brief, membrane proteins were treated with 100 mM DTT at 37°C for 1 h and then fractionated by SDS–PAGE on an 8% w/v polyacrylamide gel. After electrophoresis, proteins were electroblotted to a supported nitrocellulose membrane (Bio-Rad, http://www.bio-rad.com/). To detect ETR1–5xMyc, a 1:1000 dilution of the primary rabbit polyclonal anti-myc antibody (Sigma) was used, followed by a 1:5000 dilution of the goat anti-rabbit HRP secondary antibody (Pierce; http://www.piercenet.com). For the ECA1 protein loading control, we used an anti-ECA1 antibody (Liang et al., 1997) kindly provided by Heven Sze. Immunodecorated proteins were visualized by enhanced chemiluminescence detection using the SuperSignal West femto maximum sensitivity kit (Pierce Chemical).

Acknowledgments

We thank Chris Hawes (Oxford Brookes University, UK) for pVKH18En6-STtmd-GFP and pVKH18En6-mGFPer, Andreas Nebenführ (University of Tennessee, Knoxville, USA) for pAN33, pAN81, pAN107 and pAN186, Jeff Harper (University of Nevada, Reno, USA) for GFP–CPK9 and GFP–δTIP, Biao Ding (Ohio State University, Columbus, Ohio, USA) for pDSRed2-C1, Flanders Interuniversity Institute for Biotechnology for pBGWFS7, and Heven Sze for the anti-ECA1 antibody. We thank Amy Beaven for confocal microscopy instruction, H. Sze and Iqbal Hamza for helpful discussions, and H. Sze, June Kwak, Zhongchi Liu and members of the Chang lab for critical comments on the manuscript. Nucleotide sequencing was carried out by the Center for Biosystems Research DNA Sequencing Facility of University of Maryland Biotechnology Institute.

This work was supported by grants from the National Institutes of Health (1R01GM071855) and the United States Department of Energy (DE-FG02-99ER20329) to C.C. C.C. was supported in part by the University of Maryland Agricultural Experiment Station. B.D.M. was supported in part by a Senior Summer Scholarship as well as an Undergraduate Honors Research Grant from the University of Maryland.

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