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First published online December 23, 2004; 10.1104/pp.104.050245 Plant Physiology 137:57-69 (2005) © 2005 American Society of Plant Biologists Subcellular Localization of Arabidopsis 3-Hydroxy-3-Methylglutaryl-Coenzyme A Reductase1Departament de Bioquímica i Biologia Molecular, Facultat de Química (P.L., A.B., N.C.), Facultat de Farmàcia (M.A., A.F.), Scientific and Technical Services (S.C., C.L.-I.), and Research Center for Bioelectronics and Nanobioscience, Barcelona Science Park (X.F.-B.), University of Barcelona, E–08028 Barcelona, Spain; Institut de Biologia Molecular de Barcelona, Centre d'Investigació i Desenvolupament-Consejo Superior de Investigaciones Científicas, E–08034 Barcelona, Spain (V.M.G.); and Arizona State University, School of Life Sciences, Tempe, Arizona 85287–4501 (R.N.T.)
Plants produce diverse isoprenoids, which are synthesized in plastids, mitochondria, endoplasmic reticulum (ER), and the nonorganellar cytoplasm. 3-Hydroxy-3-methylglutaryl-coenzyme A reductase (HMGR) catalyzes the synthesis of mevalonate, a rate-limiting step in the cytoplasmic pathway. Several branches of the pathway lead to the synthesis of structurally and functionally varied, yet essential, isoprenoids. Several HMGR isoforms have been identified in all plants examined. Studies based on gene expression and on fractionation of enzyme activity suggested that subcellular compartmentalization of HMGR is an important intracellular channeling mechanism for the production of the specific classes of isoprenoids. Plant HMGR has been shown previously to insert in vitro into the membrane of microsomal vesicles, but the final in vivo subcellular localization(s) remains controversial. To address the latter in Arabidopsis (Arabidopsis thaliana) cells, we conducted a multipronged microscopy and cell fractionation approach that included imaging of chimeric HMGR green fluorescent protein localizations in transiently transformed cell leaves, immunofluorescence confocal microscopy in wild-type and stably transformed seedlings, immunogold electron microscopy examinations of endogenous HMGR in seedling cotyledons, and sucrose density gradient analyses of HMGR-containing organelles. Taken together, the results reveal that endogenous Arabidopsis HMGR is localized at steady state within ER as expected, but surprisingly also predominantly within spherical, vesicular structures that range from 0.2- to 0.6-µm diameter, located in the cytoplasm and within the central vacuole in differentiated cotyledon cells. The N-terminal region, including the transmembrane domain of HMGR, was found to be necessary and sufficient for directing HMGR to ER and the spherical structures. It is believed, although not directly demonstrated, that these vesicle-like structures are derived from segments of HMGR-ER. Nevertheless, they represent a previously undescribed subcellular compartment likely capable of synthesizing mevalonate, which provides new evidence for multiorganelle compartmentalization of the isoprenoid biosynthetic pathways in plants.
The reaction catalyzed by the enzyme 3-hydroxy-3-methylglutaryl-coenzyme A reductase (HMGR) is the first committed step in the mevalonate pathway of isoprenoid biosynthesis (Bach, 1995  ). Plants synthesize an enormous and diverse array of isoprenoids (in excess of 29,000 different compounds), which function in many aspects of growth, development, reproduction, and disease resistance (Chappell, 1995). Consistent with this complexity, all plant species studied so far have several HMGR isoforms that likely are critical in directing the flux of pathway intermediates into specific isoprenoid compounds (Enjuto et al., 1994; Chappell, 1995; Weissenborn et al., 1995). Although synthesis of many of these compounds occurs in the nonorganellar cytoplasm (sterols, sesquiterpenes), several classes of isoprenoids are synthesized in plastids (carotenoids, plastoquinone, and phylloquinone), mitochondria (ubiquinone), or specialized vacuoles (rubber particles; McGarvey and Croteau, 1995). The mevalonate pathway had been considered for many years as the unique source for the synthesis of isoprenoids. However, an alternative mevalonate-independent pathway for isoprenoid biosynthesis, also known as the methylerythritol phosphate pathway, recently has been identified in bacteria, plants, algae, and protozoa. In plants, this novel pathway is confined to the plastidic compartment (Rohdich et al., 2001; Rodríguez-Concepción and Boronat, 2002).
The in vivo subcellular localization of plant HMGRs remains controversial (Bach, 1995
However, an exclusive ER localization raises the question of how the different isoforms of HMGR, which presumably would all be present in the same endomembrane system with their catalytic domain exposed to the cytoplasm, could channel intermediates into specific families of isoprenoids and discriminate against flux into other isoprenoids. Such functional specializations have been suggested to be facilitated by discrete isoforms within distinct ER regions (Campos and Boronat, 1995 ; Chappell, 1995; McCaskill and Croteau, 1998). Studies on this topic often have been hampered by the complexity of membrane-trafficking routes in plant cells (Okita and Rogers, 1996; Hawes et al., 1999; Nebenführ, 2002; Neumann et al., 2003). In support of this hypothesis, however, the multifunctional nature of the plant ER has been attributed in several cases to a multiplicity of specialized ER regions other than the three classical subcompartments, rough ER, smooth ER, and the nuclear envelope (Okita and Rogers, 1996; Staehelin, 1997; Mullen et al., 1999; Choi et al., 2000; Naested et al., 2000; Braun, 2001; Sami-Subbu et al., 2001; Hamada et al., 2003). In this paper, we used a multipronged approach to determine the subcellular site(s) of Arabidopsis HMGR1S, the housekeeping form of the enzyme. Intracellular sorting experiments were performed with transiently expressed green fluorescent protein (GFP) chimeras possessing the N-terminal region of HMGR1S. The main thrust, however, was to determine the site(s) of the endogenous enzyme in wild-type (nontransformed) seedlings, using high-fidelity antibodies raised against the catalytic domain of HMGR1 as a specific probe for in vitro immunoblot analyses of subcellular fractions, confocal immunofluorescence microscopy, and immunogold electron microscopy. The results show that, contrary to the current prevailing hypothesis, the main reservoir of Arabidopsis HMGR in cotyledons is not within the reticular ER, but within novel cytoplasmic and intravacuolar vesicular structures that seem to be derived from subdomains of the ER.
The Amino-Terminal Domain of HMGR1S Is Sufficient for Directing GFP to the ER and to Spherical Structures
With the objective of investigating the subcellular sorting/targeting of HMGR1S and determining the role in these processes played by its amino-terminal domain, a chimeric protein was constructed such that the catalytic domain encoded by HMG1 was replaced with GFP. Thus, the polypeptide fragment of HMGR1S fused to GFP was amino acid residues 1 to 178 that included the cytoplasm-facing N-terminal region, the membrane domain consisting of both transmembrane sequences (H1 and H2) and the luminal stretch, and the linker region (Fig. 1). The resulting chimeric protein 1S:GFP was transiently expressed under control of the cauliflower mosaic virus 35S promoter in epidermal cells of 5-week-old Arabidopsis leaves after microbombardment. Confocal laser microscopy was used to determine the localization pattern of the expressed proteins (Fig. 2). Figure 2A is a positive control image showing the typical nuclear and nonorganellar cytoplasmic localization of transiently expressed, nonchimeric GFP in a transformed leaf epidermal cell. Figure 2B illustrates the results of another positive control, namely, an ER-targeted construct, pEGFPer (Danon et al., 2004
Endogenous Arabidopsis HMGR Is Localized to Spherical Structures in Cotyledon Cells
Overexpressed, membrane-targeted GFP fusion proteins have been reported to form organelle aggregates, which can lead to misinterpretations of sorting pathways for trafficked proteins (Lisenbee et al., 2003b
Rabbit polyclonal antibodies raised against the catalytic domain of HMGR1 (anti-CD1; Manzano et al., 2004
Immunofluorescence experiments (Fig. 3) were done with whole mounts of 6-d-old seedling cotyledons rather than with older plantlets or true leaves because difficulties were encountered with the penetration of the antibodies in the older plant parts, possibly due to the thicker and/or more cross-linked cell walls. Negative controls included omission of primary antibody (data not shown), and positive controls included applications of antiserum raised against the tonoplast membrane protein,  -tonoplast intrinsic protein (-TIP; Höfte and Chrispeels, 1992; Fig. 3A) or antiserum raised against cytoplasmic farnesyl diphosphate synthase isoform 1 (FPS1; Cunillera et al., 1997; Fig. 3B). Figure 3, C and D, shows that most of the HMGR signal (anti-CD1 antibody) is associated with spherical structures distributed throughout the cell, singly or in small groups, with apparent diameters ranging from 0.4 to 1.3 µm. The presence of HMGR in plastids has been a subject of debate (Brooker and Russell, 1975; Wong et al., 1982). For this reason, the autofluorescence of chlorophyll in Figure 3 is shown to provide evidence for the lack of detectable levels of HMGR in plastids.
To determine whether the amino-terminal domain of HMGR is essential for acquisition of the enzyme into the spherical structures, a transgenic line overexpressing the catalytic domain (CD1) was generated. Figure 3E shows that the truncated protein exhibited an expected cytoplasmic localization, and the inset shows that CD1 also existed in the nucleus of transformed cells. This result provides indirect evidence that the amino-terminal domain of HMGR is necessary for directing the protein to spherical structures. Figure 3F, however, shows that spherical structures acquire HMGR1S in cells of plants stably transformed with this protein (González, 2002 ), a result that, when coupled with that presented in Figure 3E, provides more direct evidence for the necessity of the HMGR N-terminal region for this targeting. The confocal immunofluorescence data described above indicate that the spherical structures are the main repository of Arabidopsis HMGR. Figure 3G shows the image of an individual confocal section of nontransformed cotyledon cells when the intensity of the green HMGR signal has been increased significantly over the red channel. This manipulation reveals that the endogenous HMGR immunofluorescence signal emanates also from a reticulated system observed in the spaces between the plastids. This latter image is consistent with a concomitant ER localization, which is similar to the reticular ER localizations observed in our GFP-fusion experiments (Fig. 2). The transgenic line transformed with the sequence coding for isoform 1S had significantly higher HMGR-specific activity than wild-type plants (Fig. 4A), whereas the corresponding increase in protein was relatively modest (Fig. 4B). The CD1-transformed plants showed the highest HMGR-specific activity (Fig. 4A). In these plants, an intense immunoblot signal for CD1 (50 kD) was, intriguingly, accompanied by a clearly increased signal for HMGR1S. Southern blots of the transgenic lines indicated that both contained a single extra DNA copy of the corresponding forms (Fig. 4D).
The cytoplasmic, punctate pattern revealed by application of anti-CD1 antibodies (Fig. 3, C, D, and G) could be consistent with the known peroxisomal localization of mammalian HMGR (Keller et al., 1985 ; Appelkvist and Kalen, 1989). Therefore, it was of interest to determine whether these HMGR-containing structures were plant peroxisomes. Lisenbee et al. (2003a) recently showed that the mouse anti-tobacco catalase monoclonal antibodies and the rabbit anti-cottonseed catalase polyclonal antibodies used in our study recognized peroxisomal catalase in Arabidopsis suspension-cultured cells, but not in plant tissue cells. Figure 5 shows the results of dual-labeling immunofluorescence experiments conducted with these antibodies in 6-d-old wild-type Arabidopsis seedling cotyledons. Punctate structures attributable to immunorecognition of the mouse anti-tobacco catalase antibodies and the rabbit anti-cottonseed catalase antibodies are shown among the red plastids (Fig. 5A) and without chlorophyll detection (Fig. 5B). The corresponding binary mask image, where only coincident pixels are shown (Fig. 5C), reveals colocalization of both antibodies in peroxisomes. These results verified the monoclonal antibody recognition of cotyledon peroxisomal catalase and provided a positive control for colocalization.
Figure 5, D and E, illustrates dual-immunolabeling results with similar cells incubated in the same mouse monoclonal anti-catalase antibodies and rabbit anti-CD1 antiserum. Signals from both antibodies are observed with (Fig. 5D) and without (Fig. 5E) visible plastids. The resulting binary mask image (Fig. 5F) clearly shows that the signals are not colocalized, indicating that, in Arabidopsis seedlings, the HMGR signal is not located within catalase-containing peroxisomes. In confocal images, the mean diameter of HMGR-containing structures is approximately 0.6 µm, whereas peroxisomes are about 1 µm in diameter, in agreement with preexisting data for peroxisome size in Arabidopsis seedlings (Mathur et al., 2002 ). The data presented above indicate that most of Arabidopsis HMGR resides in as yet unidentified abundant spherical structures somewhat smaller in diameter than cotyledon peroxisomes.
Extracts of 9-d-old wild-type Arabidopsis seedlings were cleared of unbroken whole cells and cellular debris with a 200g (5 min) centrifugation. The supernatant (S200) was then centrifuged at 16,000g (20 min), and the resulting pellet (P16) and supernatant (S16) were assayed for their HMGR activity. The results indicated that 58.8% ± 5.7% of total activity was found in P16, and 40.7% ± 5.7% was found in S16. Similar partitioning between the pellet and supernatant was observed with plants grown under long-, short-, and continuous-daylight regimes and with 6-d-old and 5-week-old plants (data not shown). After the 16,000g centrifugation, the sediment contains intact plastids and peroxisomes, while S16 contains the nonorganellar cytoplasmic components and membranes derived from all disrupted organelles, including the ER and Golgi (Leech, 1977
With the objective of further characterizing the structures possessing HMGR, Suc density-gradient centrifugations were employed. P16 and S16 prepared from leaves were loaded onto 35% to 55% (w/v) and 20% to 40% (w/v) Suc gradients, respectively. Gradient fractions were analyzed on immunoblots with anti-CD1 antiserum (Fig. 6). The HMGR signal was observed in all the fractions derived from P16 (Fig. 6A, top blot), and in the less dense region of the 20% to 40% gradient derived from S16 (Fig. 6B, top blot). From densitometric analyses of the immunoblots, the partitioning of HMGR between P16 and S16 was approximately 70:30, respectively, in good agreement with activity data (see above text). A main polypeptide band of approximately 63 kD was identified in P16 fractions, consistent with the expected size of HMGR1S, the main isoform in adult leaves (Lumbreras et al., 1995 Subcellular fraction distributions in these gradient experiments were determined with antiserum against cytoplasmic FPS1 and peroxisomal catalase, and from chlorophyll absorbance for chloroplasts. As expected, FPS1 signal was detected only in S16 fractions (Fig. 6B, middle blot; 40-kD polypeptide). Rabbit antiserum against cottonseed catalase recognized the Arabidopsis homolog (57-kD polypeptide) on immunoblots. As was anticipated, catalase released from ruptured peroxisomes was abundant at the top of the S16 20% to 40% gradients (Fig. 6B, bottom blot). Somewhat surprisingly, however, catalase was found in all fractions of the P16 35% to 55% gradient (Fig. 6A, bottom blot). These results suggest that our w/v Suc density gradients separate peroxisomes of varied density.
Subcellular localizations and ultrastructure of the HMGR-containing spherical bodies were elucidated through extensive immunogold electron microscopy studies of sections prepared from cotyledons of 6-d-old seedlings, which also were used for confocal immunofluorescence studies (Fig. 3). Figure 7, A and B, illustrates representative images of cell cytoplasm observed following dual immunogold labeling with the ER resident protein BiP (Ig heavy-chain binding protein) marked with 15-nm gold particles and HMGR marked with 10-nm gold particles. Double-labeling experiments for HMGR and BiP were done with two different anti-BiP antibodies: polyclonal rabbit anti-tobacco BiP (data shown) and monoclonal mouse anti-spinach BiP (data not shown), obtaining essentially the same labeling results in both cases. The linear arrangement of both sized gold particles is consistent with the expected colocalization of these proteins in ER. Figure 7C illustrates a representative negative gold-particle control image obtained after application of preadsorbed anti-CD1 antiserum (anti-CD1 antibodies preincubated with excess recombinant HMGR1 catalytic domain) and protein A-conjugated gold particles (10 nm). Gold particles are not observed over chloroplasts or anywhere in the nonorganellar cytoplasm surrounding a portion of the central vacuole containing a spherical structure bounded by a single membrane characteristic of vesicles. Results of dual labeling with anti-
The HMGR-containing vesicles also were observed in the nonorganellar cytoplasm, appearing as single structures (Fig. 7F) or in small groups (Fig. 7G). Although gold particles attributable to HMGR and BiP often were observed in neighboring areas in the same sections, they were not colocalized within vesicles in the nonorganellar cytoplasm or vacuoles (Fig. 7, H–L). BiP gold particles were observed over structures surrounded by cytoplasm (Fig. 7, H and I) and over structures adjacent to the tonoplast (Fig. 7, J and K) that resemble previously described BiP bodies (Levanony et al., 1992 ). Similarly, HMGR gold particles were identified in small membrane-bounded vesicles near the tonoplast (Fig. 7, J and L). In most cases, the HMGR-specific gold particles were associated with the membrane of the vesicles (Fig. 7, D–G, I, and L).
Several enzymes of the isoprenoid biosynthetic pathway in plants have been identified in more than one subcellular compartment. Such is the case for FPS in the soluble cytoplasm (Feron et al., 1990 ; Hugueney et al., 1996), mitochondria (Cunillera et al., 1997), and plastids (Sanmiya et al., 1999), and for geranylgeranyl diphosphate synthase in plastids, ER, and mitochondria (Okada et al., 2000). For one of the main regulatory enzymes of this pathway, HMGR, there is a paucity of in vivo data describing its subcellular localization. In vitro, it has been shown that Arabidopsis HMGR1S is inserted into ER microsomal membranes (Enjuto et al., 1994; Campos and Boronat, 1995; Lumbreras et al., 1995). The amino-terminal region of HMGR1S containing the membrane domain (residues 1–178) appears to be necessary (Fig. 3, immunofluorescence data) and sufficient (Fig. 2, GFP data) for targeting to and accumulation in ER and spherical structures. Consistently, the targeting domain of both yeast HMGRs has also been assigned to the amino-terminal transmembrane region of the protein (Hampton et al., 1996), and it was shown that removal of the membrane domain led to increased protein and enzyme activity (Donald et al., 1997). Transgenic plants used in this study carrying an extra CD1 copy showed greatly increased HMGR activity, indicating that when the catalytic domain was removed from its membrane environment, deregulation ensued. This suggests a key role of the endomembrane system in the control of HMGR activity.
Our results show that the main reservoir of Arabidopsis HMGR in cotyledons is not within the reticular ER, but within novel cytoplasmic and intravacuolar vesicular structures that seem to be derived from subdomains of the ER. Given the versatility of the multifunctional plant ER, it seems likely that specialized ER regions exist based on their functional attributes. Papers variously describing such so-called ER domains are plentiful (Okita and Rogers, 1996
The transport of HMGR-enriched vesicles to the vacuole could represent, among several other possibilities, the need for storage of HMGR in cotyledons or the entry of ER vesicles into the lytic vacuole by autophagy as part of a continuous or regulated HMGR flux. Vacuoles occupy up to 90% of the volume of a plant cell and are part of the endomembrane system that also includes the secretory pathway. This system comprises several compartments: ER, Golgi apparatus, trans-Golgi network, prevacuolar compartments, the vacuole, and endosomes (Surpin and Raikhel, 2004
As yet, no coherent rationale has been developed to explain the presumed regulatory roles played by the different HMGR isoforms. However, trafficking of specific forms of HMGR through the ER to other organelles in the endomembrane system has been suggested (Campos and Boronat, 1995
In some subcellular fractionation studies, plant HMGR activity has been detected not only in ER-derived membranes, but also in high-density fractions containing mitochondria, chloroplasts, and peroxisomes (Brooker and Russell, 1975 The dual localization of HMGR in the ER (where it is synthesized and inserted into its membrane) and in cytoplasmic and vacuolar vesicles suggests that, in plants, mevalonate biosynthesis can occur in different cell compartments. This, in turn, could be the molecular basis for a subcellular partitioning of isoprenoid biosynthesis.
Plant Cell Fractionation Seeds from Arabidopsis (Arabidopsis thaliana ecotype Columbia) plants were surface sterilized and then sown in petri dishes containing solid (0.8% w/v agar) germination medium (Murashige and Skoog medium supplemented with 10 g L–1 Suc, 0.5 g L–1 MES, pH 5.7). Dishes were incubated at 22°C ± 2°C for the days indicated for long- (16-h light/8-h dark), short- (8/16), or continuous-day (24/0) illumination regimes with 130 µmol m–2 s–1 daylight fluorescent illumination. Transgenic plants transformed with individual HMGR isoforms were grown on plates with germination medium containing 50 µg mL–1 kanamycin. Thereafter, 9-d-old seedlings were transplanted into soil and allowed to grow for up to 5 weeks under the same light/dark conditions. For subcellular fractionation of plant organs, leaves (400 mg) from 5-week-old plants were ground in a mortar on ice with 3 mL of extraction medium containing 0.1 M Tricine, pH 7.5, 10 mM KCl, 1 mM MgCl2, 1 mM EDTA, 1% (w/v) Ficoll, 0.1% (w/v) bovine serum albumin (BSA; Fraction V; Sigma, St. Louis), 0.5 mM phenylmethylsulfonyl fluoride, 5 mM dithiothreitol, and 20% (w/v) Suc. The ground material was cleared of large debris and nuclei in a short centrifugation at 200g (5 min, 4°C). The resulting supernatant was spun in a bench-top centrifuge at 16,000g (20 min, 4°C) to yield a pellet (P16) and supernatant (S16). Protein was determined with the DC Protein Assay from Bio-Rad (Hercules, CA). The protein content was nearly equal in P16 and S16. S16 was spun again as above and 1 mL of the resulting clarified S16, containing approximately 1.5 mg of protein, was loaded directly on top of a 40-mL continuous 20% to 40% (w/v) Suc gradient freshly prepared in Thinwall Ultra-Clear Beckman tubes (Beckman, Fullerton, CA). P16 was washed once with extraction buffer and the resulting cleaned P16, adjusted to contain approximately 1.5 mg of protein, was taken up in 1 mL of extraction buffer containing 35% (w/v) Suc and loaded on top of a 40-mL continuous 35% to 55% (w/v) Suc gradient prepared as described above. The gradients were centrifuged in a Beckman swinging bucket SW 28 rotor at 25,000 rpm (82,700g at rav) for 6 h (4°C). Approximately 20 2-mL fractions were collected immediately from each gradient by puncturing the bottom of tubes with a red-hot hypodermic needle. Fractions were kept frozen at –20°C.
HMGR activity was assayed as described in Masferrer et al. (2002)
For extraction of plant genomic DNA, approximately 1 g of fresh Arabidopsis leaves was ground in a mortar in the presence of liquid nitrogen. To the fine powder were added 5 mL of DNA extraction buffer (0.1 M EDTA, 0.25 M NaCl, 100 µg/mL proteinase K, 0.1 M Tris-HCl, pH 8.0), and the grinding was resumed until the complete disappearance of undissolved material. The resulting solution was transferred to a conical tube where N-lauroylsarcosine was added to a final concentration of 1% (w/v). After a 2-h incubation at 55°C, the tube was centrifuged (10 min, 6,000 rpm), and the resulting supernatant centrifuged again several times in the same conditions until the solution was clear. The DNA was then precipitated with 0.6 volumes of isopropyl alcohol, washed twice with 70% ethanol, slightly dried, and finally taken up in Tris EDTA buffer (1 mM EDTA, 10 mM Tris-HCl, pH 7.5). Genomic DNA was further purified by adding 1/10 volume of a NaCl/hexadecyltrimethylammonium bromide solution (5 M NaCl, 10% w/v hexadecyltrimethylammonium bromide). After a 20-min incubation at 65°C, the aqueous phase was extracted several times with 1 volume of CHCl3, and the purified DNA finally precipitated with 0.6 volumes of isopropyl alcohol, washed with 70% ethanol, and taken up in Tris EDTA buffer, pH 7.5. A digoxigenin-labeled DNA probe was amplified by PCR from a region of the HMGR1 catalytic domain cDNA lacking EcoRI sites. Primers SP6 (antisense) and an internal primer located on position 722 of the HMGR1L cDNA (sense) were used with the cloned cDNA as a template to generate an 878-bp probe that stretched until position 1,600 of the coding region. Alkali-labile DIG-dUTP (Boehringer Mannheim, Basel) was included in the PCR reaction mixture with a DIG-dUTP:dTTP ratio of 1:3. For Southern-blot analysis, genomic DNA was digested with EcoRI (Gibco BRL, Cleveland), under the conditions specified by the supplier. The digested DNA was electrophoresed in 1% agarose gels, transferred to a positively charged nylon membrane (Boehringer Mannheim) by capillary transfer, and hybridized with the DIG-labeled DNA probe overnight at 42°C. Hybridization, stringency washes, and detection were performed following the instructions in the DIG DNA-labeling kit (Boehringer Mannheim), with a final wash of 0.1x SSC, 0.1% SDS at 68°C.
For whole-mount immunofluorescence experiments, 6-d-old Arabidopsis seedlings grown under a short-daylight regime were fixed for 3 h at room temperature under vacuum in 4% (v/v) formaldehyde (prepared from paraformaldehyde) in PMEG buffer (50 mM KPIPES, pH 7.5, 2 mM MgSO4, 5 mM EGTA) and 10% (v/v) dimethyl sulfoxide, washed three times over 30 min in PMEG, and incubated for 30 min at 37°C in a cell wall-digesting solution containing 0.5% (w/v) pectolyase from Aspergillus japonicus (Sigma; 4.5 units/mg solid), 2% (v/v) Triton X-100, and 1% (w/v) BSA (Fraction V; Merck, Rahway, NJ) in PMEG. Seedlings were washed three more times in PMEG and incubated overnight at 4°C with primary antibody (see above) diluted in 0.1 M phosphate-buffered saline (PBS) containing 3% (w/v) BSA (PBS/BSA). Rabbit antiserum against bean
Plant expression vectors were constructed according to standard molecular biology procedures (Ausubel et al., 1987
For immunogold labeling of nonembedded cryosections, discs (1-mm diameter) excised from cotyledons of 6-d-old Arabidopsis seedlings grown under a short-daylight regime were chemically fixed overnight at 4°C in a mixture of 2% formaldehyde (prepared from paraformaldehyde; Ted Pella) and 0.1% (v/v) glutaraldehyde (EMS) in 0.1 M sodium phosphate buffer (PB), pH 7.5. After washing with PB containing 50 mM Gly, discs were embedded in 10% (w/v) gelatin (Merck; molecular biology grade) and infused with 2.3 M Suc in PB (Raposo et al., 1997
We thank D. Ludevid for providing the antisera against BiP and -TIP; R. Blanvillain and P. Gallois for their gift of the pEGFPer construct; J.D.I. Harper for his helpful comments about immunofluorescence techniques in Arabidopsis; and the Scientific and Technical Services of the University of Barcelona and the Servei de Camps Experimentals (Facultat de Biologia, University of Barcelona) for technical assistance, in particular to Raquel García, Sonia Ruiz, and Josep Matas for their help with the confocal immunofluorescence microscopy, with the preparation of samples for immunogold electron microscopy, and with the maintenance of Arabidopsis plants, respectively. Received July 20, 2004; returned for revision November 5, 2004; accepted November 7, 2004.
1 This work was supported by the Ministerio de Ciencia y Tecnología (MCyT; grant nos. BIO2000–0334 to A.F., BIO2002–00128 to X.F.-B., BMC2003–03450 to N.C., and BMC2003–06833 to A.B.; all grants included Fondo Europeo de Desarrollo Regional funds), and by the National Science Foundation (grant no. MCB–0091826 to R.N.T.). X.F.-B. and N.C. hold Ramón y Cajal tenure-track positions from the MCyT. P.L. acknowledges receipt of a fellowship from the Comissió Interdepartamental de Recerca i Innovació Tecnològica (Generalitat de Catalunya).  Article, publication date, and citation information can be found at www.plantphysiol.org/cgi/doi/10.1104/pp.104.050245. * Corresponding author; e-mail busquets{at}qf.ub.es; fax 34–93–4037181.
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-gliadin and BiP in the developing endosperm (Levanony et al., 1992
