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First published online October 17, 2008; 10.1104/pp.108.130575 Plant Physiology 148:1868-1882 (2008) © 2008 American Society of Plant Biologists OPEN ACCESS ARTICLE
Interaction of the WD40 Domain of a Myoinositol Polyphosphate 5-Phosphatase with SnRK1 Links Inositol, Sugar, and Stress Signaling1,[W],[OA]Department of Biochemistry, Virginia Tech, Blacksburg, Virginia 24061 (E.A.A., G.E.G., R.N.B.); and Department of Biology, Salisbury University, Salisbury, Maryland 21801 (A.E., F.L.E.)
In plants, myoinositol signaling pathways have been associated with several stress, developmental, and physiological processes, but the regulation of these pathways is largely unknown. In our efforts to better understand myoinositol signaling pathways in plants, we have found that the WD40 repeat region of a myoinositol polyphosphate 5-phosphatase (5PTase13; At1g05630) interacts with the sucrose nonfermenting-1-related kinase (SnRK1.1) in the yeast two-hybrid system and in vitro. Plant SnRK1 proteins (also known as AKIN10/11) have been described as central integrators of sugar, metabolic, stress, and developmental signals. Using mutants defective in 5PTase13, we show that 5PTase13 can act as a regulator of SnRK1 activity and that regulation differs with different nutrient availability. Specifically, we show that under low-nutrient or -sugar conditions, 5PTase13 acts as a positive regulator of SnRK1 activity. In contrast, under severe starvation conditions, 5PTase13 acts as a negative regulator of SnRK1 activity. To delineate the regulatory interaction that occurs between 5PTase13 and SnRK1.1, we used a cell-free degradation assay and found that 5PTase13 is required to reduce the amount of SnRK1.1 targeted for proteasomal destruction under low-nutrient conditions. This regulation most likely involves a 5PTase13-SnRK1.1 interaction within the nucleus, as a 5PTase13:green fluorescent protein was localized to the nucleus. We also show that a loss of function in 5PTase13 leads to nutrient level-dependent reduction of root growth, along with abscisic acid (ABA) and sugar insensitivity. 5ptase13 mutants accumulate less inositol 1,4,5-trisphosphate in response to sugar stress and have alterations in ABA-regulated gene expression, both of which are consistent with the known role of inositol 1,4,5-trisphosphate in ABA-mediated signaling. We propose that by forming a protein complex with SnRK1.1 protein, 5PTase13 plays a regulatory role linking inositol, sugar, and stress signaling.
Myoinositol (inositol) signaling pathways are important for many different developmental and physiological processes in eukaryotes (Boss et al., 2006  ; Berridge, 2007). In plants, inositol signaling is used in the response to abscisic acid (ABA; Sanchez and Chua, 2001; Xiong et al., 2001; Burnette et al., 2003; Gunesekera et al., 2007; Lee et al., 2007), salt stress (DeWald et al., 2001; Takahashi et al., 2001), gravity (Perera et al., 2001, 2006), and pathogens (Ortega and Perez, 2001; Andersson et al., 2006). The inositol signaling pathway makes use of an inositol 1,4,5-trisphosphate (InsP3) second messenger that triggers intracellular Ca2+ release from various sources within the cell (Berridge, 1993; Tang et al., 2007), although the precise mechanism in plants is not yet known (Krinke et al., 2007). To terminate signaling events driven by InsP3, cells utilize the myoinositol polyphosphate 5-phosphatases (5PTases; EC 3.1.3.56) to remove the 5-phosphate, thus initiating second messenger breakdown (Astle et al., 2007). Eukaryotes have a family of diverse 5PTases, with the Arabidopsis (Arabidopsis thaliana) genome containing 15 genes predicted to encode conserved 5PTases (Berdy et al., 2001). Of the 15 Arabidopsis 5PTase genes, four encode 5PTase enzymes with a large, N-terminal extension containing five to six WD40 repeat regions (Zhong and Ye, 2004). WD40 repeats are found in a number of eukaryotic regulatory proteins (Li and Roberts, 2001), where they are speculated to serve as a stable propeller-like platform to which proteins can bind either stably or reversibly. In some cases, sequences outside of the WD40 repeats may confer a new functional specificity to the protein even as conserved protein interactions are maintained through the WD40 repeats, potentially adapting basic cellular mechanisms to organism-specific processes. WD40-containing 5PTases are unique to plants and certain fungi, as no other genomes contain candidate genes for these proteins (Zhong and Ye, 2004).
By examining plants containing either a gain or loss of function in specific 5PTases, investigators have established that 5PTases are critical in plant development and in ABA signaling. In particular, mutations in the CVP2 (5PTase6; At1g05470) and 5PTase13 (At1g05630) genes have been linked to altered cotyledon vascular patterning and/or blue light responses and phototropin 1 signaling (Carland and Nelson, 2004
5PTases have been reported to play a role in Glc sensing/metabolism in animal cells (Wada et al., 2001
We report here that the WD40 repeat region of the 5PTase13 gene interacts specifically with a Suc nonfermenting-1-related kinase (SnRK1.1, AKIN10; At3g01090), which functions as a sensor of energy and stress in plants (Baena-Gonzalez et al., 2007
5PTase13 Is a Member of a Unique Group of WD40-Containing Proteins
Besides containing a conserved inositol polyphosphate 5-phosphatase catalytic domain, 5PTase13 also contains five WD40 repeat regions in the N terminus that may allow for unique protein interactions. Three other genes in the Arabidopsis genome, 5PTase12 (At2g43900), 5PTase14 (At2g31830), and FRA3 (At1g65580), encode similar proteins (Berdy et al., 2001
The similarity of the 5PTases (Zhong and Ye, 2004 ) suggests that these proteins may function in a redundant manner. Microarray data from GENEVESTIGATOR (Zimmermann et al., 2004) indicates that FRA3 is broadly and abundantly expressed compared with 5PTase12, 5PTase13, and 5PTase14 (Supplemental Fig. S1A). We conclude that the WD40-containing 5PTases are a small group of proteins with the potential to form protein complexes and that functional redundancy may be present.
To investigate the ability of the WD40 regions of 5PTase13 to participate in protein complexes, we used the yeast two-hybrid system. The 533 N-terminal amino acids from 5PTase13 containing the WD40 repeats were used as bait in a yeast two-hybrid screen of an Arabidopsis 3-d-old etiolated seedling cDNA library. We screened over 1 million yeast transformants and obtained a positive clone containing the C-terminal domain of the SnRK1.1 gene (At3g01090; also known as AKIN10). We retransformed this positive clone into yeast and verified the interaction (Fig. 2 ). Negative controls, including the empty DNA binding domain and activation domain vectors, established that SnRK1.1 binds to the WD40 repeat region of the 5PTase13 protein in yeast.
To examine the interaction between 5PTase13 and SnRK1.1 in vitro, we fused the sequence encoding the Xpress epitope tag to the C terminus of the WD40 repeat region of 5PTase13 (13WDX) and the V5 epitope tag sequence to the C terminus of SnRK1.1 (SnRKV5) and expressed both proteins in Escherichia coli. The SnRKV5 construct directs the expression of a 61.7-kD protein detected by an anti-V5 monoclonal antibody, and in most cases we detected two SnRKV5 bands, perhaps as a result of phosphorylation or proteolytic cleavage (Fig. 3 , lane 1). The 13WDX construct directs the expression of a 65.9-kD protein detected by an anti-Xpress monoclonal antibody (Fig. 3, lane 2). These interactions are specific, as SnRKV5 is not detected by the anti-Xpress antibody and 13WDX is not detected by the anti-V5 antibody (data not shown). To determine whether the SnRK1.1 recombinant protein can "pull down" 13WDX, immunoprecipitations using anti-V5:protein A-Sepharose beads were performed, and the resulting complex was then analyzed by western blotting with an anti-Xpress antibody (Fig. 3, IP lanes). As shown in Figure 3, the WD40 repeat region from 5PTase13 (13WDX) is only detected in this pull-down assay when it has been incubated in the presence of SnRKV5. We conclude that the WD40 repeat region from 5PTase13 and SnRK1.1 can form a protein complex in vitro.
SnRK1.1, along with its closely related gene family member SnRK1.2, encodes a Suc nonfermenting-1-related kinase implicated as a central integrator of energy signaling and metabolic regulation in yeast, plants, and animals. The interaction of 5PTase13 and SnRK1.1 is novel and is perhaps unique to plants in that yeast and animal 5PTases do not contain WD40 regions. This interaction may indicate that InsP3 signal termination via 5PTase13 function affects the SnRK1 "energy sensor" in plants. Data obtained from the GENEVESTIGATOR database indicate that SnRK1.1, SnRK1.2, and 5PTase13 are detected in most plant tissues examined, although 5PTase13 expression levels are very low compared with SnRK1 gene expression (Supplemental Fig. S1B).
To further explore the link between InsP3 signaling and the energy sensor, SnRK1.1, we isolated two independent T-DNA insertion mutants in the 5PTase13 gene. Two potential mutants were identified in the SALK T-DNA database and named 5ptase13-1 (SAIL_350_FO1) and 5ptase13-2 (SALK_081991) and were compared with their corresponding wild-type accessions (Fig. 4A
). The presence of the T-DNA insertion was verified by diagnostic PCR in each mutant using genomic DNA and primers specific for the T-DNA left border (LB) and gene-specific primers that flank the T-DNA insertion (Fig. 4, A and B). The resulting LB gene-specific fragments were sequenced, indicating that in 5ptase13-1 mutants, a second T-DNA insertion is found in tandem in the fourth exon (Fig. 4, A and B, LB-R band). This is in contrast to the previously reported analysis by Lin et al. (2005)
Using primers specific for the 3' end of 5PTase13, we detected a PCR product in both wild-type lines used (CS60000 and CS908) but not in 5ptase13-1 and 5ptase13-2 mutants (5PTase13 in Fig. 4C). Using primers that amplify the 5' end, we detected a 1.68-kb product in both wild-type lines and in the 5ptase13-1 mutant, but not in 5ptase13-2 (Fig. 4C). We conclude that the 5ptase13-2 mutant is totally lacking 5PTase13 expression and that both 5ptase13-1 and 5ptase13-2 mutant lines do not express transcripts capable of encoding a full-length 5PTase13 protein. To examine how the loss of 5PTase13 affects the expression of its binding partner SnRK1.1, we examined the expression of SnRK1.1 and its closely related isoform, SnRK1.2, in 7-d-old dark-grown 5ptase13 and wild-type seedlings. The results reveal that there are no large changes in SnRK1.1 and SnRK1.2 in 5ptase13 mutants (Fig. 4D). As shown in Figure 4D, we found that FRA3 expression remains unchanged in 5ptase13 mutants, and 5PTase14 expression is barely detectable but also unchanged. In contrast, the expression of 5PTase12 is increased in both 5ptase13 mutant lines, revealing a possible means of compensation for the loss of 5PTase13 function (Fig. 4D).
Under standard laboratory conditions, 5ptase13 mutants did not exhibit any abnormalities in plant growth or development. Since Lin et al. (2005)
To determine whether 5PTase13 affects SnRK1 function, we measured the activity of SnRK1 in 5ptase13 mutants and wild-type seedlings grown under various nutrient conditions. It is well documented that SnRK1 regulates multiple transcription cascades in response to sugar or energy deprivation (Baena-Gonzalez et al., 2007 As expected from its role as a low-energy sensor, SnRK1 activity is higher in seedlings grown on low nutrients compared with extracts prepared from seedlings grown on optimal nutrients or 6% Glc (Fig. 5 ). Figure 6A shows that the activity of SnRK1 significantly increases in 5ptase13-1 and 5ptase13-2 mutants when seedlings are grown with no nutrients (8.1- and 11.3-fold increases, respectively). In contrast, SnRK1 activity decreases in 5ptase13-1 and 5ptase13-2 seedlings compared with wild-type seedlings when grown with low nutrients (Fig. 6B). Furthermore, the decline in SnRK1 activity is even more dramatic when 6% Glc is added (7.1- and 3.3-fold reduction, respectively; Fig. 6C). We conclude that a loss of 5PTase13 function affects SnRK1 activity, and the impact differs depending on nutrient availability. Our data support a role for 5PTase13 as a negative regulator of SnRK1 activity in the absence of nutrients and as a positive regulator when either low nutrients or 6% Glc is present.
5PTase13 Affects SnRK1.1 Stability
To explore whether 5PTase13 regulates SnRK1.1 stability, we used a similar approach as Lee et al. (2008)
To test the hypothesis that 5PTase13 destabilizes SnRK1.1 under the no-nutrient conditions, we examined a 30-min time point for analysis as a midpoint to total SnRKV5 degradation. The data indicate no difference in SnRKV5 stability in wild-type and 5ptase13 extracts under the no-nutrient conditions (Fig. 7C). In contrast, when extracts are prepared from seedlings grown on low nutrients, we find less SnRKV5 accumulation in 5ptase13 extracts compared with wild-type extracts (Fig. 7D). We conclude that under the low-nutrient conditions, 5PTase13 is required to stabilize SnRKV5 protein and slow its degradation by the 26S proteasome pathway. Furthermore, the increased degradation of SnRKV5 seen with the low-nutrient conditions correlates well with the lower SnRK1 activity levels measured in 5ptase13 mutants under these same nutrient conditions (Fig. 6B). In contrast, 5PTase13 is not required to stabilize SnRKV5 when seedlings are grown with no nutrients, and this is consistent with the switch in SnRK1 activity levels we found in 5ptase13 mutants grown with no nutrients (Fig. 6A). However, since we did not observe an increase in SnRKV5 stability in 5ptase13 extracts prepared from the no-nutrient conditions, we speculate that there is an additional mechanism that influences the elevated SnRK1 activity in 5ptase13 mutants under these conditions.
We compared 5ptase13 mutants with SnRK1.1 mutants, which have a small reduction in root growth under the low-nutrient conditions and exhibit enhanced root growth with 1% to 3% Suc (Baena-Gonzalez et al., 2007
5PTase13 Alterations Change Plant Sensitivity to Sugars and ABA To test whether 5PTase13 is required for sugar and ABA responses, we analyzed age-matched seeds for germination in the presence of 0%, 1%, 3%, or 6% Glc or 0%, 1%, 2%, 3%, 6%, or 11% Suc. At low concentrations of sugar, there were no differences in the germination of wild-type and 5ptase13 seeds in the dark or light (data not shown). However, at a high exogenous sugar concentration (6% Glc and 11% Suc) in the dark, we found that 5ptase13 mutants were significantly less sensitive to sugar (Fig. 8D; Supplemental Figs. S3 and S4). We saw the same trend in sugar insensitivity when 5ptase13 mutant seeds were germinated in the presence of 6% Glc in the light, although the sugar insensitivity was less apparent (Supplemental Fig. S3). This sugar insensitivity was also noted for 5ptase13-1 and 5ptase13-2 mutant seeds germinated in the presence of 11% Suc, in which maximal increases of 2.5- and 2.2-fold for 5ptase13-1 and 5ptase13-2, respectively, were noted compared with wild-type seeds (Supplemental Fig. S4). Germination in the presence of mannitol, a nonmetabolizable sugar, was not altered, indicating that the sugar insensitivity of 5ptase13 mutants is not due to a general osmotic stress tolerance (Fig. 8D).
We also germinated 5ptase13 mutant seeds in the presence of 0, 1, 2, and 3 µM ABA and measured the impact on germination during a 6-d period. As expected, germination of wild-type seeds was delayed by ABA in a concentration-dependent manner during the 6-d period under light conditions (Fig. 8, E and F). In contrast, 5ptase13-1 and 5ptase13-2 mutant seeds were ABA insensitive, reaching 100% germination on day 2 in the presence of 1 µM ABA and 78% to 90% on day 6 in the presence of 3 µM ABA (Fig. 8, E and F). Since we did not find reduced seed dormancy in our mutants (data not shown), this ABA insensitivity of 5ptase13 mutants most likely does not correlate with changes in de novo synthesis of ABA (Gubler et al., 2005
To determine whether there are differences in the expression patterns of Glc- and/or ABA-regulated genes (RD29A, KIN1, and CAB1 genes; Price et al., 2004
Complementation of the 5ptase13-1 Mutant To ensure that the alterations noted in 5ptase13 mutants are due to loss of 5PTase13 expression, we expressed a 5PTase13:GFP fusion under the control of the 35S cauliflower mosaic virus promoter in wild-type and 5ptase13-1 plants (13:GFP and 13-1/13:GFP plants, respectively). We examined two lines of 13-1/13:GFP plants with good expression of the transgene (Fig. 9B) along with wild-type and 5ptase13-1 plants in ABA-sensitivity assays. Both 13-1/13:GFP lines exhibited more ABA sensitivity in germination assays compared with 5ptase13 mutants (Fig. 9C). Since 5ptase13-1 mutants contain decreased SnRK1 activity under low-nutrient conditions compared with wild-type plants (47%; Fig. 6B), we also measured SnRK1 activity in the 13-1/13:GFP-1 and 13-1/13:GFP-2 lines and found, as expected, similar or increased SnRK1 activity compared with wild-type plants (140% ± 10% for 13-1/13:GFP-1 and 102% ± 5% for 13-1/13:GFP-2). We conclude that expression of the 5PTase13:GFP transgene complements the 5ptase13-1 mutant.
We examined whether the Glc insensitivity of 5ptase13 mutants is accompanied by alterations in mass InsP3 levels by measuring mass InsP3 levels. The results in Figure 10 indicate that neither 5ptase13-1 nor 5ptase13-2 mutant seedlings differ significantly from wild-type seedlings in their InsP3 mass levels under control conditions. When wild-type seedlings are exposed to 6% Glc for 3 d, mass InsP3 levels increase 2.8- to 3.7-fold, which is a statistically significant elevation. However, when 5ptase13 mutants are grown for 3 d in the presence of 6% Glc, mass InsP3 level changes are smaller, with an increase of 1.6- to 2-fold over basal levels, and statistically significant only in the 5ptase13-2 mutant. More importantly, the Glc-stimulated InsP3 levels in both 5ptase13 mutants differ significantly from the Glc-stimulated InsP3 levels in wild-type seedlings. We conclude that 5ptase13 mutants are impaired in their ability to accumulate InsP3 in response to Glc and that this correlates with the sugar and ABA insensitivity noted in the germination assays.
Subcellular Localization of the 5PTase13:GFP Fusion Protein To investigate the subcellular location of the 5PTase13 protein, we performed imaging experiments with 5ptase13-1 mutants complemented with the 5PTase13:GFP construct (13-1/13:GFP plants) and wild-type plants containing the same 5PTase13:GFP construct. As the 5PTase13:GFP construct we used allowed for complementation of the ABA and sugar insensitivity phenotype in 5ptase13-1 mutants (Fig. 9, B and C), it is likely that this fusion protein undergoes the same posttranslational modifications and subcellular localization as the native 5PTase13 protein. We analyzed T2 progeny from two independent 13-1/13:GFP lines with fluorescence deconvolution microscopy and found a similar pattern in both lines. GFP fluorescence was associated with the nucleus in many, but not all, cells in cotyledon epidermis (Fig. 11, B and C ), hypocotyls (data not shown), and roots (Fig. 11, D–I). Nuclei from some but not all guard cells contained the 13:GFP protein (Fig. 11C). To confirm the nuclear localization, we stained 13-1/13:GFP seedlings with the nuclear dye 4',6-diamidino-2-phenylindole (DAPI) and imaged GFP and DAPI fluorescence simultaneously (Fig. 11, D–I). Once again, we found 13:GFP fluorescence in only a portion of root nuclei, while DAPI fluorescence was present in all nuclei. We conclude that 5PTase13 protein is located in the nucleus of seedlings and that its presence in the nucleus is restricted in some cells.
Since control of second messengers is critical for signaling, there is interest in determining how the plant cell regulates levels of second messengers such as InsP3. We identified a potential regulator of InsP3 signaling by isolating protein interactors of 5PTase13. We focused on 5PTase13 because it contains several conserved WD40 repeats (for review, see van Nocker and Ludwig, 2003 ) and the presence of WD40 repeats within 5PTase genes is unique to plants and certain fungi (Zhong and Ye, 2004). To date, the WD40 repeats of 5PTases have not been functionally characterized, and no potential protein partners have been identified. We show here an important and novel interaction between the WD40 repeat region of 5PTase13 and the central metabolic regulator, SnRK1.1. SnRK1.1 and its orthologues in yeast and mammals is one subunit of a complex that modulates cellular metabolism in response to nutrient and environmental conditions (Baena-Gonzalez et al., 2007; Hardie, 2007; Hue and Rider, 2007). Recent work in Arabidopsis has elegantly delineated the role of SnRK 1.1 in the transcriptional activation of genes from catabolic pathways that provide routes for alternative energy sources and coordinate repression of a large number of anabolic genes (Baena-Gonzalez et al., 2007). Our discovery of a 5PTase13:SnRK1.1 complex points to the novel interaction of this metabolic modulator and InsP3 signaling.
It has been shown previously that a loss of function in the 5PTase13 gene leads to defects in auxin-regulated development (Lin et al., 2005 To understand how a 5PTase13:SnRK1.1 complex might regulate nutrient and stress signaling, we measured SnRK1 activity in 5ptase13 mutants and wild-type seedlings grown under different nutrient conditions. Our data show that the presence of 5PTase13 affects SnRK1 activity and that nutrient availability is an important switch in 5PTase13 regulation of SnRK1.1. Specifically, 5PTase13 is required to maintain wild-type levels of SnRK1 activity when low nutrient or high sugar is present (Fig. 6, B and C). In contrast, under the greatest starvation conditions of no added nutrients, 5PTase13 appears to be a negative regulator of SnRK1, as 5ptase13 mutants contain significantly elevated SnRK1 activity (Fig. 6A). To further delineate the mechanism of 5PTase13 regulation of SnRK1.1, we used a cell-free degradation assay and showed that the decrease in SnRK1 activity in 5ptase13 mutants correlates with increased degradation of SnRK1.1 by the proteasome (Fig. 7D).
Interestingly, another WD40 repeat-containing protein called PRL1 (for PLEITROPIC REGULATOR LOCUS1) also interacts with SnRK1.1 and is able to inhibit the activity of SnRK1.1 and SnRK1.2 (Bhalerao et al., 1999
Our data reported here, along with data from other groups, support the model of SnRK1 as a sensor of low nutrient status and cellular stress. Under low-nutrient or sugar-stress conditions, 5PTase13 acts as a positive regulator of SnRK1.1 activity by reducing the amount of SnRK1 targeted for proteasomal destruction. This regulation most likely involves a 5PTase13-SnRK1.1 interaction within the nucleus, as that is where we find 5PTase13:GFP (Fig. 11), and SnRK1 is most likely nuclear as well (Pierre et al., 2007
While data support this model, we currently do not know how second messenger InsP3 affects the SnRK1 pathway. We have shown that InsP3 levels increase when seedlings are given a sugar stress, implicating InsP3 as a second messenger under high-sugar conditions (Fig. 10). Given that the 5PTase13 enzyme has been shown to hydrolyze InsP3 in vitro (Zhong and Ye, 2004
There have been previous reports that inositol signaling and sugar sensing/metabolism are linked. Im et al. (2007)
Together, the data presented here support a unique role for 5PTase13 functioning as a binding partner and regulator of the SnRK1.1 modulator of energy and stress signaling. Given the previously established role of 5PTase13 in blue light signaling (Chen et al., 2008
Phylogenetic Tree Construction BLASTp was used to identify 12 related WD40 regions. ClustalW and PAUP4.0 were used to generate phylogenetic trees using parsimony. One thousand bootstraps were performed.
The Matchmaker Two-Hybrid System 3 was used (BD Biosciences Clontech). The cDNA corresponding to amino acids 1 to 533 of 5PTase13 was amplified by PCR and ligated into pGBKT7 bait vector (BD Biosciences Clontech), verified by DNA sequencing, and transformed into yeast strain AH109. The yeast strain AH109 containing the 5PTase13 WD40 repeat domain was transformed with an Arabidopsis (Arabidopsis thaliana) 3-d-old etiolated seedling cDNA library (Thelogis). Screening for interactors was performed with SD/-Ade-His-Leu-Trp, 10 mM 3-amino-1,2,4-triazole, and 20 µg mL–1 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside plates. Candidate clones that grew were rescued from yeast and retested in the original bait and control strains. Prey plasmids that passed all tests were sequenced to identify the Arabidopsis gene insert.
Arabidopsis ecotype Columbia was used for all experiments. Growth conditions of soil-grown plants have been described (Berdy et al., 2001
Conditions for reverse transcription (RT)-PCR have been described previously (Ercetin et al., 2008
Age-matched seeds used for assays were harvested from plants grown in parallel on the same shelf in a growth room, and seeds were harvested on the same day and ripened for 6 weeks at room temperature. Seeds were surface sterilized and plated on no salts or 0.5x MS salts solution (pH 5.7) containing 0.8% agar. Seeds were stratified on plates at 4°C for 3 d and germinated at 23°C in the light or dark. For seed germination and root growth assays, seeds were plated on medium containing 0%, 1%, 3%, or 6% Glc or mannitol (Sigma-Aldrich) or 0%, 1%, 2%, 3%, 6%, or 11% Suc (Sigma-Aldrich) in the light or dark at 23°C. Germination was scored as positive when the radicle protruded through the seed coat. For ABA sensitivity experiments, ABA (Sigma-Aldrich) was dissolved in 100% ethanol and added to cooled, sterile medium at a final concentration of 1, 2, or 3 µM ABA. The germination assay was performed as before. Hormone/sugar treatment experiments were repeated two or three times.
The entire open reading frame (encoding 535 amino acids) of At3g01090 was amplified with gene-specific primers SnRK1.1pFor and SnRK1.1pRev (Supplemental Table S4) and ligated to pCRT7/CT-TOPO vector. This resulted in pSnRKV5, which directed expression of a 61.7-kD SnRK1.1 protein with a C-terminal 6xHis tag and V5 epitope tag (Invitrogen). The WD40 repeat region of 5PTase13 was amplified with gene-specific primers WD40For and WD40Rev (Supplemental Table S4) and ligated to pCRT7/NT-TOPO vector. This resulted in p13WDX, which directed expression of a 65.9-kD protein that corresponds to the WD40 region of 5PTase13 and an N-terminal 6xHis tag and Xpress epitope (Invitrogen). For expression, pSnRKV5 and p13WDX were transformed in Escherichia coli BL21(DE3) pLysS cells (Invitrogen), and the resulting recombinant proteins 13WDX and SnRKV5 were purified using nickel-nitrilotriacetic acid agarose columns (Invitrogen). SnRKV5 protein appeared on the SDS protein gel as two separate bands. These bands represented an active SnRKV5 recombinant protein as determined by the SnRK1 activity assay.
For the immunoprecipitation, between 40 and 90 ng of the purified and dialyzed proteins was used. The immunoprecipitation reactions were carried out as described by Ercetin et al. (2008)
Whole 7-d-old seedlings were ground in liquid nitrogen and resuspended in extraction buffer (50 mM Tris-HCl, pH 7.8, 1 mM EGTA, 1 mM EDTA, 2 mM dithiothreitol [DTT], 0.05% Nonidet P-40, 0.5 mM phenylmethylsulfonyl fluoride, 1 mM benzamide, 3 mM glycerophosphate, and plant protease cocktail; Sigma). After centrifugation at 13.2 rpm for 15 min at 4°C, ammonium sulfate was slowly added to the supernatant to 40% saturation while stirring for 10 min at 4°C and centrifuged for 15 min at 13.2 rpm and 4°C. Precipitated protein was resuspended in 50 µL of fractionation buffer (50 mM Tris-HCl, pH 7.8, 1 mM EGTA, 1 mM EDTA, 2 mM DTT, 0.05% Nonidet P-40, 0.5 mM phenylmethylsulfonyl fluoride, 1 mM benzamide, 3 mM glycerophosphate, 10% glycerol, and plant protease cocktail; Sigma). Protein concentration was determined according to the Bradford method (Bradford, 1976
Conditions described by Lee et al. (2008)
The 3,408-bp coding region of 5PTase13 minus the stop codon was amplified by high-fidelity PCR, confirmed by sequencing, cloned into the pENTR/D-TOPO vector (Invitrogen), and recombined via the Gateway system (Invitrogen) using the manufacturer's instructions into pK7FWG2. The resulting 35S cauliflower mosaic virus promoter:5PTase13:GFP construct was transformed into Agrobacterium tumefaciens by cold shock and was used in the transformation of 5ptase13-1 and wild-type plants as described (Bechtold et al., 1993
Filters containing whole 4-d-old dark-grown seedlings were floated on a 6% Glc solution (0.5x MS salts, pH 5.7) or on a control solution (0.5x MS salts, pH 5.7, only) in the dark for 3 d and then frozen in liquid nitrogen at the end of the treatment. Tissues were harvested and mass InsP3 measurements were made as described previously (Gunesekera et al., 2007
The histological analysis was performed as described (Carland et al., 1999 Sequence data from this article can be found in the GenBank/EMBL data libraries under the following accession numbers: 5PTase13, At1g05630, NP_172054; SnRK1.1, At3g01090, NP_001118546; 5PTase12, At2g43900, NP_181918; 5PTase14, At2g31830, NP_180742; FRA3, At1g65580, NP_176736.
The following materials are available in the online version of this article.
This paper is dedicated to the memory of Emily Jane Hilscher. We are grateful to SIGnAL and the Arabidopsis Biological Resource Center for supplying mutant seeds. We also thank Dr. P.J. Kennelly (Virginia Tech) for advice on the SnRK1 activity assay, Dr. Jae-Hoon Lee and Dr. Xing Wang Deng (Yale University) for advice on the cell-free degradation assay, and Janet Donahue (Virginia Tech) for assistance with InsP3 measurements. Received September 30, 2008; accepted October 14, 2008; published October 17, 2008.
1 This work was supported by the National Science Foundation (grant no. MCB–0641954 to G.E.G.) and the U.S. Department of Agriculture (grant no. 2003–35318–13690 to G.E.G.) and by Hatch program funds (grant no. VA–135583).  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.plantphysiol.org) is: Glenda E. Gillaspy (gillaspy{at}vt.edu).
[W] The online version of this article contains Web-only data.
[OA] Open Access articles can be viewed online without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.108.130575 * Corresponding author; e-mail gillaspy{at}vt.edu.
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ABSTRACT
RESULTS
-32P]ATP, and 5 µg of plant protein extract for 30 min. Validation that these conditions are in the linear range of product accumulation is shown in Supplemental Figure S2. 
40). The experiment was independently repeated two times. D, Comparison of germination of wild-type and 5ptase13 seeds grown on 0.5x MS salts and 0.8% agar that were untreated (control) or treated with 6% Glc, 6% mannitol, or 11% Suc after 7 d in the dark. Values are means ± SE (n = 50). The data are representative of three independent experiments. E and F, Wild-type and 5ptase13 mutant seeds were germinated in the light for 6 d on 0.5x MS salts, 0.8% agar, and either 1 µM ABA (E) or 3 µM ABA (F). The germination rate was scored for each group of seeds starting from day 1. Values are means ± SE (n = 50). Significant differences from wild-type germination were noted at days 1 to 5 for 5ptase13-1 (1 µM ABA) and days 4 to 6 for both mutants (3 µM ABA). The experiment was independently repeated two times. *, P < 0.05 compared with the wild type.
