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First published online April 25, 2008; 10.1105/tpc.107.055632 The Plant Cell 20:1073-1087 (2008) © 2008 American Society of Plant Biologists Functional and Physiological Characterization of Arabidopsis INOSITOL TRANSPORTER1, a Novel Tonoplast-Localized Transporter for myo-Inositol[W]
a Molekulare Pflanzenphysiologie, Friedrich-Alexander-Universität Erlangen-Nürnberg, D-91058 Erlangen, Germany 1 Address correspondence to nsauer{at}biologie.uni-erlangen.de.
Arabidopsis thaliana INOSITOL TRANSPORTER1 (INT1) is a member of a small gene family with only three more genes (INT2 to INT4). INT2 and INT4 were shown to encode plasma membrane–localized transporters for different inositol epimers, and INT3 was characterized as a pseudogene. Here, we present the functional and physiological characterization of the INT1 protein, analyses of the tissue-specific expression of the INT1 gene, and analyses of phenotypic differences observed between wild-type plants and mutant lines carrying the int1.1 and int1.2 alleles. INT1 is a ubiquitously expressed gene, and Arabidopsis lines with T-DNA insertions in INT1 showed increased intracellular myo-inositol concentrations and reduced root growth. In Arabidopsis, tobacco (Nicotiana tabacum), and Saccharomyces cerevisiae, fusions of the green fluorescent protein to the C terminus of INT1 were targeted to the tonoplast membranes. Finally, patch-clamp analyses were performed on vacuoles from wild-type plants and from both int1 mutant lines to study the transport properties of INT1 at the tonoplast. In summary, the presented molecular, physiological, and functional studies demonstrate that INT1 is a tonoplast-localized H+/inositol symporter that mediates the efflux of inositol that is generated during the degradation of inositol-containing compounds in the vacuolar lumen.
In plants, sugars and sugar alcohols play an essential role as transport and storage forms of fixed carbon, they represent the main energy source, they are the initial substrates for numerous biosynthetic reactions, and they may be accumulated in response to several kinds of stress. Their biosynthesis occurs in fully developed and mature source leaves, from where they are allocated via the phloem to the different sink tissues and organs of higher plants.
The relative amounts of sugars and sugar alcohols that are transported in the phloem vary between different plant species. In many plants, sucrose is the exclusive or by far dominant form of assimilated carbon (e.g., in Arabidopsis thaliana or tobacco [Nicotiana tabacum]). Other species, however, transport only low concentrations of sucrose in combination with raffinose family oligosaccharides, such as raffinose, stachyose, or verbascose (Sauer, 2007
The long-distance allocation and partitioning of linear and cyclic polyols depends on the presence of specialized biosynthetic machineries in the source leaves of the respective plant species and on the activity of transport proteins that are able to load mannitol, sorbitol, or inositol into the phloem of these plants. Genes or cDNAs encoding transporters for polyol substrates were first cloned from plants that translocate polyols in their phloem (Chauhan et al., 2000
Arabidopsis has six genes with significant similarity to genes encoding transporters for linear polyols in polyol-translocating plants (PLT genes; Klepek et al., 2005
The monosaccharide-transporter(-like) gene family of Arabidopsis has >50 members, and most of the genes analyzed to date encode plasma membrane transporters (Büttner, 2007
So far, no transport proteins for polyols have been characterized from plant endomembranes. Although Chauhan et al. (2000) Here, we present a detailed characterization of INT1, a member of the Arabidopsis INT family. We report on the characterization of the functional and physiological properties of this protein by heterologous expression of its cDNA in baker's yeast (Saccharomyces cerevisiae) and by analyses of two independent int1 mutant lines, on the subcellular localization of an INT1-GFP (for green fluorescent protein) fusion, and of the tissue specificity of the INT1 promoter. Finally, we present patch-clamp analyses of isolated plant vacuoles that characterize INT1 as a tonoplast-localized transporter that is capable of mediating H+ symport of myo-inositol. The physiological role of INT1 is discussed.
The Arabidopsis INT1 Protein Sequence Cloning of a full-length INT1 cDNA has been described previously (Schneider et al., 2006  ). Analyses of these sequence data showed that the INT1 gene (At2g43330) is interrupted by four introns and encodes an mRNA with an open reading frame (ORF) of 1527 bp. The INT1 protein (509 amino acids) has a predicted molecular mass of 54.2 kD, a pI of 5.05, and 12 predicted transmembrane helices. It is significantly smaller than the two previously characterized Arabidopsis inositol transporters, INT2 and INT4, that have molecular masses of 62.9 kD (pI = 8.70) and 63.4 kD (pI = 8.25), respectively. This size difference is mainly due to an elongated, Cys-rich loop that is found between the predicted transmembrane helices IX and X. The Cys-rich region (69 amino acids) of this loop is present in INT2 and INT4 but missing from INT1.
Within the monosaccharide-transporter(-like) superfamily of Arabidopsis, INT1 clearly clusters together with the three other INT proteins (Büttner, 2007
INT1 Mediates the Accumulation of myo-Inositol in S. cerevisiae The functional properties of the previously described inositol transporters from Arabidopsis, INT4 (Schneider et al., 2006 ) and INT2 (Schneider et al., 2007), had been studied by complementation of a baker's yeast mutant (S. cerevisiae strain D458-1B; Nikawa et al., 1991) deficient in myo-inositol uptake ( itr1) and biosynthesis (ino1). Wild-type yeast strains possess two genes for inositol transporters, ITR1 and ITR2. Whereas ITR2 is constitutively expressed and the encoded protein catalyzes myo-inositol uptake at only low rates, ITR1 expression is induced by low medium concentrations of inositol, and ITR1-driven myo-inositol uptake occurs at high rates (Nikawa et al., 1993; Miyashita et al., 2003). Both proteins, ITR1 and ITR2, are plasma membrane–localized (Nikawa et al., 1991), and like INT2 and INT4 from Arabidopsis, they possess a long loop between their predicted transmembrane helices IX and X. A yeast transporter of the INT1 type has not been identified.
An INT1 cDNA was expressed in the sense and antisense orientations in the D458-1B yeast mutant. When the resulting yeast strains SSY36 (sense orientation) and SSY37 (antisense orientation) were grown on media with different myo-inositol concentrations, no functional complementation of the
To discriminate between these different possibilities, we analyzed the intracellular localization of INT1 in the yeast wild-type strain SEY2102 (Emr et al., 1983 ), which carried a construct for an INT1 protein fused to the N terminus of the GFP. This INT1-GFP–expressing strain was named SSY16. Figure 2B demonstrates that INT1-GFP localizes preferentially to the yeast vacuoles (arrows). Labeling of the plasma membrane was not detected. This suggested that INT1 might be a tonoplast protein. In some cells, additional dot-like structures showed strong GFP labeling. These structures might be aggresomes that are known to occur sometimes upon overexpression of foreign proteins (García-Mata et al., 1999).
We next determined the total concentrations of myo-inositol in various yeast strains at different growth stages (Figure 2C). In particular, we studied the SEY2102 (wild type)–derived strains SSY1 (INO1, ITR1, INT1), SSY9 (INO1, ITR1), and SSY16 (INO1, ITR1, INT1-GFP) and the D458-1B (mutant)–derived strains SSY36 (
This difference became even more obvious when similar analyses were performed on stationary-phase cells. In these analyses, all cells that expressed INT1 or INT1-GFP showed strongly increased myo-inositol concentrations, whereas both controls (wild-type SSY9 and mutant SSY37) did not. This demonstrated that INT1 and INT1-GFP are functional myo-inositol transporters in yeast and that their activity is responsible for the observed accumulation of myo-inositol. The low myo-inositol concentration observed in stationary-grown SSY9 cells may reflect either reduced myo-inositol biosynthesis as a consequence of reduced glucose availability (Granot and Snyder, 1991
To test whether the observed myo-inositol accumulation (Figure 2C) results from sequestration within the yeast vacuoles, uptake analyses were performed with [3H]myo-inositol on isolated yeast vacuoles. As a control for the integrity and functionality of the vacuoles, we determined the ATP-dependent uptake of [14C]Lys. This amino acid is imported into yeast vacuoles by a Lys/H+ antiporter that is energized by the vacuolar H+-ATPase (Sato et al., 1984
INT1 Localizes to the Tonoplast in Arabidopsis and Tobacco
These results demonstrated that INT1 is a protein of Arabidopsis tonoplast membranes. In combination with the accumulation data obtained for INT1-GFP–expressing yeast cells (Figure 2C) that demonstrate that INT1-GFP is a functional myo-inositol transporter, a mistargeting of the INT1-GFP fusion due to the C-terminal tag is quite unlikely.
Tissue- and Organ-Specific Expression of INT1 None of the INT1 promoter/GFP plants showed detectable GFP fluorescence, suggesting that the activity of the INT1 promoter is rather low. This was supported by published microarray analyses (http://www.bar.utoronto.ca/efp/cgi-bin/efpWeb.cgi) and by the results obtained from the INT1 promoter/GUS lines (Figure 4 ). GUS histochemical staining was found in almost all cells and all tissues at all developmental stages. Germinating (Figure 4A) or young seedlings (Figure 4B), rosette leaves (Figure 4C), and flowers (Figure 4E) showed comparable intensities of GUS staining. In cross sections of rosette leaves, this staining was found in the mesophyll but was lacking in both epidermis cell layers (Figure 4D). Flowers showed less intense staining in petals than in most other floral tissues (Figure 4E), and GUS staining seen in anthers was restricted mainly to pollen grains. Surprisingly, no GUS staining was seen in developing seeds at any developmental stage (Figure 4G; see Supplemental Figure 1 online). In principle, these results confirmed data from publicly accessible microarray analyses that also described low and ubiquitous expression of INT1 in all tissues and organs (http://www.genevestigator.ethz.ch/ and http://bbc.botany.utoronto.ca). The only difference observed was the complete absence of GUS staining in seeds, which was not confirmed by the microarray data.
Generation of Anti-INT1 Antisera For an independent confirmation of the GUS analyses presented in Figure 4, we aimed to generate INT1-specific antibodies and to perform immunolocalization analyses. To this end, a fusion construct (pSS11-AK1) was made of the last 31 C-terminal amino acids (starting with Trp-479 of the INT1 sequence) that was fused to the C terminus of the maltose binding protein (MBP) from Escherichia coli. The resulting MBP-INT1 fusion protein was used to generate polyclonal antisera.
A protein gel blot with anti-INT1 antiserum (
 INT1 was used to immunolocalize INT1 protein in sections from different tissues and organs of Arabidopsis plants; however, we were not able to detect INT1-specific fluorescence. We also tested the quality of the INT1 on thin sections of SSY36 and SSY37 cells. These sections were prepared using the identical fixation and embedding protocol that had also been used for the plant material. Again, the INT1 treatment did not yield any immunolabeling in the INT1-expressing SSY36 cells. These results demonstrate that either the antigenic epitope is destroyed during the fixation procedure or the amount of INT1 is too low for immunodetection in tissue sections.
Characterization of Mutant Lines Carrying T-DNA Insertions in the INT1 Gene
For both insertion lines, homozygous plants could be isolated (Figure 6B). PCR analyses performed with cDNA that had been synthesized from total RNA of int1.1 plants showed that the T-DNA insertion in these plants resulted in a complete loss of full-length INT1 mRNA and of RNA sequences downstream from the insertion site (Figure 6C). However, a truncated mRNA corresponding to the region upstream from the insertion site could be identified (Figure 6C). Similarly, a full-length INT1 mRNA was no longer detected in int1.2 plants. In contrast with int1.1 plants, however, a weak band for a truncated downstream mRNA was identified in int1.2 plants (Figure 6C). This may result from a promoter activity in the left border region of the T-DNA. No PCR was performed to identify a truncated mRNA from the upstream region, as the insertion was too close to the 5' end of the gene in the int1.2 allele.
Homozygous int1.1 and int1.2 plants were used for analyses of potential phenotypic differences between mutant and wild-type plants. Germination analyses of seeds on agar medium, comparisons of plant growth on soil, and analyses of soil-grown plants after NaCl stress or cold treatment showed no differences. However, for int1.1 and int1.2 plants, reduced root lengths were detected on agar medium. Figure 7A
shows 14-d-old seedlings that were grown on Murashige and Skoog (MS) medium (Murashige and Skoog, 1962
These data suggest that int1.1 plants suffer from a reduced availability of myo-inositol. Therefore, we determined the myo-inositol concentrations in ethanol extracts from rosette leaves of 75-d-old int1.1 and wild-type plants. Surprisingly, the myo-inositol concentration in these leaves was  2-fold higher than in wild-type leaves (Figure 7C). For comparison, we also determined the myo-inositol concentrations of rosette leaves of int2.1 (Schneider et al., 2007) and int4.2 (Schneider et al., 2006) mutant plants of the same age. These mutants carry T-DNA insertions that cause null mutations in the INT2 and INT4 genes, respectively. The analyzed plants showed the identical myo-inositol concentrations as wild-type plants (Figure 7C). In contrast with the myo-inositol concentrations, the concentrations of glucose, fructose, and sucrose were almost identical in all mutant lines and in wild-type plants (Figure 7C). The difference in total myo-inositol concentrations observed between int1.1 and wild-type plants was also seen when younger plants were analyzed or with plants after cold treatment or NaCl stress (data not shown). Moreover, this difference was observed in all double mutants that carried the int1.1 mutation (e.g., in int1.1 int2.1) but never in double mutants in which the INT1 gene was intact (e.g., in int2.1 int4.2; data not shown). Finally, analyses of the total myo-inositol concentrations in rosette leaves of int1.2 plants showed a similar, although less pronounced, increase (Figure 7D). This smaller increase in the myo-inositol concentration in int1.2 plants might be explained by an observation already mentioned above. In contrast with int1.1, which seems to be a real knockout allele (Figure 6C), int1.2 plants possess a truncated mRNA corresponding to sequences downstream from the T-DNA insertion site (Figure 6C). Therefore, they may still have some residual INT1 transport activity. The observed reduced availability of myo-inositol, the inositol-dependent root growth, and the simultaneous increase in cellular inositol can only be explained by the sequestration of inositol in an intracellular compartment, most likely within the vacuoles.
Patch-Clamp Analyses of Isolated Vacuoles from Arabidopsis Plants
In fact, when the inositol concentration in the bath medium of vacuoles from Arabidopsis wild-type plants was raised from 0 to 50 mM (i.e., to a most likely saturating concentration), immediate positive currents were observed (Figures 8A and 8C). By convention (Bertl et al., 1992 ), currents in this direction are defined as currents out of the cytoplasm (i.e., in this case into the vacuole). This result suggests that INT1 transports myo-inositol and H+ out of the vacuole. When identical analyses were performed on vacuoles from int1.1 mutant plants, this current could not be resolved (Figures 8A and 8C). As expected from the data presented in Figure 6C (which shows truncated int1.2 mRNA) and in Figure 7D (which shows a less pronounced change in the cellular inositol concentration of int1.2 mutant leaves), analyses of vacuoles from the int1.2 mutant yielded detectable currents that were significantly lower than those measured in the wild type (Figures 8A and 8C). These data confirm that int1.1 represents a knockout mutant, whereas int1.2 is a knockdown mutant. The comparative analyses of cellular sugar and inositol concentrations in wild-type and different knockout plants (Figure 7C) revealed changes only for myo-inositol. Cellular glucose and fructose concentrations were not affected, suggesting that these hexoses are not substrates for INT1. This suggested that INT1 might be myo-inositol–specific, just like the related plasma membrane transporters INT2 and INT4. Nevertheless, we compared the capacity of vacuoles from wild-type and int1.1 mutant plants to mediate the efflux of glucose (Figures 8B and 8C). Obviously, glucose transport activities could be determined in vacuoles from both plants. The presented patch-clamp data independently confirm that INT1 is a tonoplast-localized inositol transporter. Most importantly, however, they demonstrate that INT1 mediates the symport of myo-inositol and protons across the tonoplast. Na+ can be excluded as a potentially cotransported ion, as sodium salts were absent from both the bath solution and the buffer in the patch pipette.
Higher plant vacuoles occupy up to 95% of the volume of mature cells, represent the main storage compartment for solutes, are involved in the regulation of cell volume and turgor, mediate essential steps in the degradation of cellular components, and play a key role in cellular ion homeostasis. Moreover, vacuoles are indispensable for the deposition of secondary metabolites (Marinova et al., 2007 ) and for multiple detoxification processes, including heavy metals and xenobiotics (Wink, 1993). For all of these different tasks, vacuoles depend on specific and efficient transport proteins that are embedded into their membrane, the tonoplast (Martinoia et al., 2007). Despite these numerous essential functions of the tonoplast for plant growth and development, few tonoplast proteins have been functionally characterized. To date, only four solute transporters for amino acids or sugars were described in Arabidopsis: one amino acid transporter (CAT2; Su et al., 2004), two different monosaccharide transporters (TMT1; Wormit et al., 2006; VGT1; Aluri and Büttner, 2007), and one transporter for the disaccharide sucrose (SUC4; Endler et al., 2006). In sugar beet (Beta vulgaris), another vacuolar solute transporter has been identified (Chiou and Bush, 1996) that is similar to proteins (At1g75220 and At1g19450) of a so far uncharacterized subgroup of the Arabidopsis monosaccharide-transporter(-like) gene family (Büttner, 2007).
During the last few years, several groups performed analyses of the higher plant vacuolar proteome (Carter et al., 2004
INT1 Is a Tonoplast-Localized, Energy-Dependent Transporter for myo-Inositol
Attempts to express INT1 in Xenopus laevis oocytes for more detailed analyses of substrate specificity, kinetic properties, or energy dependence of INT1 were not successful, and expression of an INT1-GFP fusion construct in Xenopus oocytes demonstrated that the fusion protein did not localize to the plasma membrane (U. Hammes, P. Udvardi, and N. Sauer, unpublished data). Therefore, it was not possible to study the transport properties of INT1 in this expression system. However, the results obtained from Xenopus oocytes with the closely related plasma membrane–localized transporters INT2 and INT4 (Schneider et al., 2007 Due to the lack of detectable INT1 transport activity in these well established expression systems, we aimed to characterize the function of INT1 by comparing mesophyll vacuoles isolated from wild-type plants or from transgenic Arabidopsis lines that carried a T-DNA insertion in the INT1 gene (int1.1 plants).
The results obtained from these analyses identified a myo-inositol transport activity that was present in wild-type plants but absent from int1.1 plants (Figures 8A and 8C). This demonstrates (1) that this activity can be attributed to the INT1 protein and (2) that INT1 is targeted to the tonoplast with (Figure 3) or without a C-terminal GFP fusion. Second, these analyses characterize INT1 as an energy-dependent transporter that generates inositol-dependent currents into the vacuoles of wild-type plants. However, as already pointed out above, for experimental reasons the patch-clamp analyses shown in Figure 8 were performed in the inverse mode (i.e., at high vacuolar and low cytoplasmic pH values). This allows changes in the myo-inositol concentrations during studies of a possible symporter activity of INT1. Previous characterizations of the closely related INT2 and INT4 proteins as H+/inositol symporters of the plasma membrane (Schneider et al., 2006 The complete absence of inositol currents in int1.1 vacuoles (Figures 8A and 8C) demonstrates that INT1 is the only inositol exporter at the tonoplast, a fact that was most likely indispensable for the identification of the clear differences between vacuoles from wild-type and knockout plants (Figure 8C). Surprisingly enough, the comparison of glucose-induced currents in wild-type and int1.1 vacuoles (Figures 8A and 8C) showed reduced currents for this substrate in mutant vacuoles (50% of the wild type; Figure 8C). However, whether this reduction is due to the absence of INT1 that may have some marginal glucose transport activity, or whether INT1 is highly specific for inositol and the observed reduction is an indirect effect (e.g., of the altered inositol availability in the cytoplasm; see analyses in Figure 7), cannot be said. This question can only be answered in the absence of interfering transport activities of other tonoplast transporters. Crosses with knockout lines of already characterized and possibly yet to be identified tonoplast transporter genes will be necessary for detailed analyses of substrate specificities in planta.
Nevertheless, two lines of evidence suggest that INT1 is most likely a specific transporter for inositols: (1) the previously characterized transporters INT2 and INT4 are highly specific for different inositol epimers (Schneider et al., 2006 Our patch-clamp data obtained from reverse mode analyses are in agreement with the predicted physiological function of INT1 as a H+/inositol exporter from the vacuolar lumen into the cytoplasm.
INT1 Drives myo-Inositol Export from Arabidopsis Vacuoles
What Is the Origin of Vacuolar myo-Inositol?
However, to our knowledge, the uptake of cytoplasmic myo-inositol into plant vacuoles or even accumulation to high concentrations within the vacuoles has not been described. Even in roots of ice plant, where myo-inositol is delivered from the phloem, myo-inositol accumulates in the cytoplasm and not in vacuoles (Nelson et al., 1999
However, IP6 biosynthesis occurs mainly in seeds, where it is deposited as an organic storage form for phosphorus and minerals (Brinch-Pedersen et al., 2002
Nevertheless, IP6 or enzymes involved in IP6 biosynthesis were also found in leaves, roots, and tubers of different plant species (Brearley and Hanke, 2000
Even a complete hydrolysis of this IP6 would result in an increase in free myo-inositol concentrations of only 4 µg/g, a value that is too small to account for the observed increase in myo-inositol in int1.1 plants (50 µg/g in Figure 7C). Moreover, it is not entirely clear whether all of the IP6 found in leaves is in vacuoles (Bentsink et al., 2003 The sum of these and other reactions might result in the production of significant amounts of myo-inositol in the vacuolar lumen, and INT1 seems to mediate the release of this recycled myo-inositol into the cytoplasm. A lack of this transporter would then result in the observed phenotype. Similar reactions are likely to occur in all other plant species as well, and the identification of one or several transporters of the INT1 type in all plant species analyzed so far (Figure 1) suggests similar functions for these proteins.
Strains and Growth Conditions Arabidopsis thaliana (Columbia wild type) plants were grown in growth chambers on potting soil or on MS plates under long-day conditions (16 h of light/8 h of dark) at 22°C and 60% relative humidity for most analyses. For ion chromatographic analyses, plants were grown on soil under short-day conditions (8 h of light/16 h of dark). The Saccharomyces cerevisiae strain D458-1B (Nikawa et al., 1991 ) was used for heterologous expression of INT1 cDNA. The Escherichia coli strain DH5 (Hanahan, 1983) was used for basic cloning steps, and the MBP-INT1 fusion protein for antisera was expressed in E. coli strain Rosetta (Novagen). Transformation of Arabidopsis was performed using Agrobacterium tumefaciens strain GV3101 (Holsters et al., 1980).
cDNA Cloning and Constructs for Yeast Expression
INT1 Promotor/GUS and INT1 Promotor/GFP Constructs
Transient Expression of INT1-GFP
Complementation Analyses in Yeast
Preparation of Yeast and Plant Material for Ion Chromatographic Analyses
Ion Chromatography
Analysis of the T-DNA Insertion Lines int1.1 and int1.2 The primers INT1-5-Pci (5'-ACATGTCATTGACGATCCCAAACG-3'), INT1-3-Pci (5'-ACATGTTGGCAGATTGAGATCCCTGCTCGAG-3'), Int1cs218 (5'-CAGGAAACTATTGTAAGTATGGCTTT-3'), Int1g557r (5'-ATGGCACCAACTAAAGCCATAC-3'), Int1cs774 (5'-TCTTTCAGCGGCTGAAGAGG-3'), and Int1cs1220r (5'-GGGTATATCTCTGAGTTCACCG-3') were used for RT-PCRs with total RNA from wild-type and mutant Arabidopsis plants. The primers AtACT2g+846f (5'-ATTCAGATGCCCAGAAGTCTTGTT-3') and AtACT2g+1295r (5'-GAAACATTTTCTGTGAACGATTCCT-3') were used to amplify the ACT2 mRNA.
Analyses of Arabidopsis Root Lengths
Immunohistochemical Techniques and Protein Gel Blot Analyses
Protein extracts of total membrane fractions from baker's yeast were prepared as described (Sauer and Stolz, 2000
Microscopy and Detection of GFP Fluorescence GUS plants were analyzed with a stereomicroscope (Leica MZFLIII; Leica Microsystems) or a microscope (Zeiss Axioskop; Carl Zeiss). Images were processed using the analySIS Doku 3.2 software (Soft Imaging System).
Patch-Clamp Analyses of Isolated Plant Vacuoles
Patch-clamp experiments on mesophyll vacuoles were performed in the whole-vacuole configuration essentially as described by Schulz-Lessdorf and Hedrich (1995)
Phylogenetic Analyses
Accession Numbers
Supplemental Data
We thank Enrico Martinoia (University of Zürich) for his help in the large-scale preparation of plant vacuoles. This work was supported by Grant SA 382/13 from the Deutsche Forschungsgemeinschaft to N.S.
The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantcell.org) is: Norbert Sauer (nsauer{at}biologie.uni-erlangen.de).
[W] Online version contains Web-only data. www.plantcell.org/cgi/doi/10.1105/tpc.107.055632 Received September 10, 2007; Revision received March 25, 2008. accepted March 31, 2008.
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-TIP and 
-TIP (Klebl et al., 2003