- © 2018 American Society of Plant Biologists. All rights reserved.
Abstract
Inorganic pyrophosphate (PPi) is a phosphate donor and energy source. Many metabolic reactions that generate PPi are suppressed by high levels of PPi. Here, we investigated how proper levels of cytosolic PPi are maintained, focusing on soluble pyrophosphatases (AtPPa1 to AtPPa5; hereafter PPa1 to PPa5) and vacuolar H+-pyrophosphatase (H+-PPase, AtVHP1/FUGU5) in Arabidopsis thaliana. In planta, five PPa isozymes tagged with GFP were detected in the cytosol and nuclei. Immunochemical analyses revealed a high abundance of PPa1 and the absence of PPa3 in vegetative tissue. In addition, the heterologous expression of each PPa restored growth in a soluble PPase-defective yeast strain. Although the quadruple knockout mutant plant ppa1 ppa2 ppa4 ppa5 showed no obvious phenotypes, H+-PPase and PPa1 double mutants (fugu5 ppa1) exhibited significant phenotypes, including dwarfism, high PPi concentrations, ectopic starch accumulation, decreased cellulose and callose levels, and structural cell wall defects. Altered cell arrangements and weakened cell walls in the root tip were particularly evident in fugu5 ppa1 and were more severe than in fugu5. Our results indicate that H+-PPase is essential for maintaining adequate PPi levels and that the cytosolic PPa isozymes, particularly PPa1, prevent increases in PPi concentrations to toxic levels. We discuss fugu5 ppa1 phenotypes in relation to metabolic reactions and PPi homeostasis.
INTRODUCTION
Inorganic pyrophosphate (PPi) is a high-energy phosphate compound that is generated as a by-product of over 200 metabolic reactions, including fatty acid β-oxidation and DNA, RNA, aminoacyl-tRNA, and polysaccharide biosynthesis (Heinonen, 2001; Ferjani et al., 2014). In most organisms other than plants, PPi hydrolysis is catalyzed by the soluble form of inorganic pyrophosphatase (sPPase; EC 3.6.1.1). Genetic defects in sPPase lead to the intracellular accumulation of PPi, resulting in severe growth defects or cell death owing to thermodynamic inhibition of the abovementioned essential reactions (Chen et al., 1990; Serrano-Bueno et al., 2013). In addition to orthologs of sPPase, plants also contain vacuolar H+-pumping inorganic pyrophosphatase (H+-PPase). Arabidopsis thaliana contains six paralogs of sPPase, PPa1 to PPa6 (Schulze et al., 2004; Navarro-De la Sancha et al., 2007). sPPase isozymes are classified into two families: Mg2+-dependent enzymes in Family I and Mn2+-dependent enzymes in Family II. The six sPPase isozymes in Arabidopsis require Mg2+ for their activity and thus belong to Family I. Family II sPPases are found in only a limited number of bacteria and archaea (Shintani et al., 1998; Young et al., 1998).
Family I members are separated into two clades: prokaryotic and eukaryotic types (Schulze et al., 2004; Gómez-García et al., 2006). The prokaryotic type is represented by the hexameric enzyme ppa in Escherichia coli and the eukaryotic type by the dimeric enzyme IPP1 (INORGANIC PYROPHOSPHATASE1) in yeast (Saccharomyces cerevisiae). Arabidopsis sPPases PPa1 to PPa5 are of the prokaryotic type and are localized to the cytosol (Koroleva et al., 2005; Öztürk et al., 2014; Gutiérrez-Luna et al., 2016). PPa6, on the other hand, is of the eukaryotic type and is localized to plastids in Arabidopsis (Schulze et al., 2004). The subcellular localization of plant sPPases has been examined using several methods: immunogold labeling for potato (Solanum tuberosum) enzymes (Rojas-Beltrán et al., 1999), analysis of GFP fusion proteins for Arabidopsis enzymes (Koroleva et al., 2005; Öztürk et al., 2014; Gutiérrez-Luna et al., 2016), and an in vitro import assay for pea (Pisum sativum) PPa6 (to chloroplasts; Schulze et al., 2004). PPi hydrolysis activity has been detected for several sPPase isozymes in plants (Schulze et al., 2004; Gómez-García et al., 2006; Navarro-De la Sancha et al., 2007). Nonetheless, the physiological roles of these cytosolic sPPases are not fully understood.
In evaluating the physiological functions of sPPases, two points are worth considering: (1) the enzymatic activity of plant sPPases is very low and (2) cytosolic PPi concentrations are relatively high (0.2–0.3 mM) (Weiner et al., 1987). The enzymatic activity of plastid sPPases is markedly higher than that of the cytosolic isozymes, and PPi levels are lower in plastids than in the cytosol (Weiner et al., 1987). Gene silencing of plastid PPa6 in tobacco (Nicotiana tabacum) resulted in dynamic changes in chloroplast metabolism (George et al., 2010), suggesting that PPa6 is a major enzyme for PPi hydrolysis in plastids. Of the cytosolic isozymes, the overexpression of PPa1 driven by a seed-specific promoter reduced seed oil content and increased starch and soluble sugar contents in Arabidopsis (Meyer et al., 2012). The reduced PPi levels in PPa1-overexpressing plants are thought to stimulate both the β-oxidation of stored lipids and sugar biosynthesis. By contrast, seed-specific silencing of PPa1 and PPa4 led to a 1 to 4% increase in seed oil contents (Meyer et al., 2012). Taken together, these findings indicate that sPPases strongly influence lipid and sugar metabolism in plants.
We recently reported that AtVHP1/FUGU5/AVP1, encoding a vacuolar H+-PPase, regulates cytosolic PPi levels (Ferjani et al., 2011; Asaoka et al., 2016; Fukuda et al., 2016). Three independent VHP1 loss-of-function mutant lines (the fugu5 mutant series) have been identified based on their gross phenotypes (including oblong-shaped cotyledons) and their cellular phenotypes (namely, compensation; i.e., cell enlargement due to a decrease in cell number) (Ferjani et al., 2007). This phenotype is caused by an increase in PPi levels, which inhibits the gluconeogenesis of seed storage lipids to sucrose (Ferjani et al., 2011; Takahashi et al., 2017). At the postgermination stage of development, fugu5 mutants also exhibit moderately delayed growth, although they appear healthy (Asaoka et al., 2016; Fukuda et al., 2016). This contrasts with bacteria and yeast, in which loss of function of sPPase results in cell death (Chen et al., 1990; Serrano-Bueno et al., 2013). Therefore, it is important to evaluate the physiological balance between cytosolic sPPases and vacuolar H+-PPase in plants.
In this study, we examined the expression of sPPase isozymes and their roles in PPi hydrolysis in Arabidopsis. We also characterized loss-of-function mutants of PPa1, PPa2, PPa4, and PPa5, together with various double knockout mutants of these sPPase isozyme genes and fugu5. Among the double mutants, a few combinations, such as fugu5-1 ppa1-1, displayed severe phenotypes, whereas single knockout mutants exhibited rather moderate phenotypes. Finally, we determined the PPi contents in mutant plant tissues. Together, our findings provide important information about the relationships among loss-of-function mutations, phenotypes, and PPi levels. We discuss the physiological cooperation between vacuolar H+-PPase and sPPase isozymes in regulating cytosolic PPi levels.
RESULTS
Generation of Loss-of-Function Mutants of Cytosolic Soluble PPase Isozymes
To investigate the phenotypes of Arabidopsis mutants lacking each of the PPa isogenes, we obtained the following T-DNA insertion lines from the Nottingham Arabidopsis Stock Centre (http://arabidopsis.info): ppa1-1, SAIL_251_D07; ppa2-1, SAIL_618_H05; ppa4-1, SAIL_916_C08; and ppa5-1, SALK_014647. To confirm the insertion of T-DNA into the corresponding genes, we performed PCR using genomic DNA from each line with T-DNA- and gene-specific primers (Supplemental Figure 1). We also generated multiple knockout mutants through crosses: The quadruple mutant ppa1-1 ppa2-1 ppa4-1 ppa5-1 is referred to as ppa1,2,4,5 hereafter. The mutant lines were used for isozyme measurements and phenotypic analyses.
Detection of Cytosolic PPa Isozymes
To detect PPa isozymes in tissues, we prepared microsomal and soluble fractions by ultracentrifugation and performed immunoblotting with an anti-PPa antibody raised against a sequence conserved among the five isozymes. An immunostained band was clearly detected in the soluble fraction from the wild type (Supplemental Figure 2A). A faint band was observed in the wild-type microsomal fraction but not in the fraction from the quadruple mutant ppa1,2,4,5. These results suggest that the anti-PPa antibody is useful for detecting PPa proteins and that PPa is found in the soluble fraction. Together with previous reports on GFP fusion analysis (Koroleva et al., 2005; Öztürk et al., 2014; Gutiérrez-Luna et al., 2016), these results confirm the presence of PPas in the cytosol.
The amino acid sequences of the five PPa isozymes are only 69% identical (Schulze et al., 2004), although these proteins have similar molecular masses (Supplemental Figure 2B). These differences in primary amino acid sequences suggest that it is possible to distinguish between each isozyme. We therefore separated the proteins by two-dimensional polyacrylamide gel electrophoresis (2D-PAGE) and detected them by immunoblotting (Supplemental Figure 2C). We identified spots corresponding to each PPa isozyme by comparing them with those from the wild type with multiple knockout lines, although we observed several nonspecific spots larger than 28 kD on the immunoblots. For PPa1 and PPa5, we detected two spots with different isoelectric point (pI) values (Supplemental Figure 2C). The lower pI spots might represent phosphorylated proteins, as the phosphorylation of plant sPPases has been reported (de Graaf et al., 2006; Eaves et al., 2017). The pI values of each spot were comparable to the calculated pI values of each isozyme. We failed to detect a spot corresponding to PPa3, with a calculated pI of 5.63 and a molecular mass of 24.9 kD in the wild type (Supplemental Figures 2B and 2C), suggesting that PPa3 is absent in vegetative tissues.
Tissue Specificity of PPa Isozymes
To examine the tissue specificity of the PPa isozymes, we determined their relative abundance by immunoblotting with anti-PPa antibody. First, we compared the relative levels of PPa isozymes in roots and shoots at 16 d after sowing (DAS) and in other organs of 5-week-old plants (Figures 1A and 1B). PPa isozymes were detected at ∼27 kD in all wild-type tissues. In 5-week-old plants, the total isozyme content was highest in flowers. We also examined samples from the quadruple mutant ppa1,2,4,5 in parallel with the wild type but did not detect any bands corresponding to PPas (Figures 1A and 1B). These results point to the low abundance of PPa3 in these tissues.
Tissue Specificity of Arabidopsis PPa Isozymes Detected by Immunoblotting.
The soluble fractions prepared from various organs of individual knockout lines and the quadruple ppa1,2,4,5 mutant were immunoblotted with anti-PPa. The gels were also silver-stained to test the amount of protein loaded. Relative intensity of the immunoblot is shown as a percentage compared with the wild type ([A] and [C]) or leaves (B).
(A) Relative quantities of PPa isozymes in the supernatants of homogenates prepared from shoots and roots of the wild type and five individual mutants at 16 DAS. The molecular masses of marker proteins are given on the right (kD).
(B) Relative amounts of PPa isozymes in leaves, stems, buds, flowers, and fruits of 37-DAS plants. Soluble fractions were prepared from the wild type and quadruple mutant ppa1,2,4,5.
(C) Comparison of PPa levels in five organs. Soluble fractions were prepared from the wild type and ppa1-1, ppa2-1, ppa4-1, and ppa5-1 single mutants.
Next, we compared the abundance of each PPa in individual single knockout mutant lines with that in the wild type by immunochemical analysis. Wild-type leaves and flowers showed intense bands, but the ppa1 mutants exhibited faint bands (Figure 1C), confirming the abundance of PPa1 in leaves and flowers. PPa1 was also more abundant in shoot and root tissues than were the other isozymes.
PPa2 was abundant in buds and fruits, as indicated by the decreased immunoblot intensity for the ppa2 mutants (Figure 1C). Shoots, roots, leaves, stems, and flowers also contained considerable amounts of PPa2. On the other hand, stems and buds contained substantial amounts of PPa4. For PPa5, the intensity of stained bands was slightly reduced in all tissues examined except roots and leaves. A spot clearly corresponding to PPa5 was detected by 2D-PAGE analysis (Supplemental Figure 2). Thus, PPa5 appears to be broadly expressed in most plant tissues at rather low levels. We could not examine PPa3 content in this study because no corresponding knockout mutant was available.
Visualization of PPa Isozymes via the Expression of GFP Fusion Protein
To further investigate the spatiotemporal localization of PPa isozymes, we constructed GFP fusion proteins with PPa1, PPa2, PPa3, PPa4, and PPa5. To observe the natural endogenous expression patterns of these proteins, the constructs contained the promoters, introns, and 3′ untranslated regions (UTRs) of these genes. In this experiment, we used cfSGFP2 (hereafter, cfGFP), a cysteine-free GFP variant with strong green fluorescence (Suzuki et al., 2012). As discussed below, the expression of PPa1-cfGFP and PPa4-cfGFP rescued the fugu5-1 ppa1-1 and fugu5-1 ppa4-1 phenotypes, respectively, indicating that PPa-cfGFPs have enzymatic activity when expressed in seedlings and localize to the proper subcellular spaces (Supplemental Figure 3). All PPa-cfGFPs were detected in the cytosol and nucleus (Figures 2A to 2F). In the immunoblot analysis, no 27-kD band corresponding to free GFP, which can freely pass through nuclear pores, was detected, suggesting that the nucleus-localized signals were from genuine PPa-cfGFP (Figure 2G). The results of PPa-GFP spatiotemporal expression analysis are shown in Supplemental Figures 4 to 8 and are summarized in Table 1.
Subcellular Localization of PPa Isozymes.
(A) to (F) Roots expressing PPa1, PPa2, PPa3, PPa4, PPa5, or VHP1 fused with GFP under the control of their own promoters from plants grown for 6 to 21 d on 0.5× MS plates containing 1% sucrose were observed by CLSM. The root regions observed in the experiment were as follows: (A) PPa1-cfGFP line #14, elongation zone; (B) PPa2-cfGFP line #12, division zone; (C) PPa3-cfGFP line #16, root hairs; (D) PPa4-cfGFP line #11, differentiation zone; (E) PPa5-cfGFP line #5, differentiation zone; and (F) VHP1-mGFP line #wM-A9, differentiation zone. Arrows indicate nuclei. Green fluorescence from GFP-linked PPa isozymes was detected in the cytosol and nuclei. Bars = 20 μm.
(G) Immunochemical analysis of PPa-GFP protein. A 5-µg sample of the soluble fraction prepared from shoots of 20 DAS plants was immunoblotted with anti-GFP.
PPa Isozyme Activity
The five cytosolic PPa isozymes share high sequence similarity, except for their N-terminal regions. The five isozymes share a sequence identity of 69% (Schulze et al., 2004). This high identity suggests that these isozymes share functional similarity, acting as pyrophosphatases in planta. PPa1 and PPa4 possess PPi hydrolysis activity, as revealed in an E. coli heterologous expression system (Navarro-De la Sancha et al., 2007). The enzymatic functions of PPa2, PPa3, and PPa5, however, have not been confirmed. We attempted to measure pyrophosphatase activity in soluble fractions prepared from plants by ultracentrifugation but did not detect any differences between the wild type (0.105 ± 0.004 [se] units/mg protein, n = 3) and ppa1,2,4,5 (0.102 ± 0.002 [se] units/mg protein, n = 3). The detected activity was probably derived from vacuolar acid phosphatase (Veljanovski et al., 2006), and sPPase activity might be negligible compared with that of vacuolar acid phosphatase(s). Because the plant vacuole accounts for over 90% of the cell volume in mature tissues, cytosolic sPPase must be separated from the other abundant phosphatases, such as vacuolar acid phosphatases, prior to measurement. In addition, perhaps the PPa isozymes were inactivated during preparation of the soluble fractions from tissue homogenates.
Therefore, we analyzed the PPase activity of each PPa isozyme following expression in a transgenic yeast yTT1 strain in which the Gal1 promoter was replaced by the promoter of the yeast cytosolic sPPase gene IPP1. Arabidopsis PPa isogenes were introduced individually into yTT1 yeast cells under the control of the IPP1 promoter. Although PPa levels varied among transformants, all yeast strains expressing PPa isozymes exhibited normal growth in glucose-based culture medium in which the expression of endogenous IPP1 was strongly suppressed (Supplemental Figures 9A and 9B), indicating that these PPa isozymes possess PPase activity.
Phenotypic Properties of Single and Multiple PPa Mutants, Particularly the fugu5 ppa1 Double Mutant
The ppa1,2,4,5 mutant showed no obvious phenotype (Figures 3 and 4). There are two possible explanations for this: (1) Cytosolic PPa isozymes may not contribute to PPi homeostasis or (2) perhaps there are other redundant pyrophosphatases that hydrolyze cytosolic PPi. To examine the physiological roles of vacuolar H+-PPase and cytosolic PPa isozymes in PPi homeostasis, we crossed the PPa knockout mutants with VHP1-deficient lines, fugu5-1 and fugu5-3. The resulting double knockout mutants were all viable and fertile. Nonetheless, several double knockout mutants showed significant phenotypes in their roots and shoots (Figures 3 and 4; Supplemental Figure 10). In particular, the severe phenotype of fugu5-1 ppa1-1 was also detected in the hypocotyls of etiolated seedlings. Notably, fugu5-1 ppa1-1 showed severe defects in hypocotyl elongation in both the presence and absence of sucrose (Figures 5A and 5B).
Phenotypes of Multiple Knockout Mutants of PPa and H+-PPase in Shoots.
(A) and (B) Plants grown on 0.5× MS plates containing 1% sucrose or no sugar were captured with a digital camera (A) or a stereomicroscope (B) at 13 DAS. Arrowheads indicate leaf atrophy, and magnified images are also shown on the right. Bars = 10 mm in (A) and 2 mm in (B). Arrows indicate atrophied areas (B).
(C) Fresh weights of whole plants. The box plot was created using R version 3.1.2, and different letters above each bar indicate statistically significant differences (P < 0.05, Tukey’s HSD test; R version 3.1.2). n = 15 to 21 from three independent biological replicates.
Root Phenotypes of Multiple PPa and H+-PPase Knockout Mutants.
(A) Root lengths of seedlings grown on 0.5× MS plates with 2% sucrose for 13 d. Measurements taken after 5 d (from day 8 to day 13) are shown. Bars = 10 mm.
(B) fugu5-1 ppa1-1 was grown on a 0.5× MS/2% sucrose plate for 6 d, transferred to a sugar-free plates, and grown for 18 d.
(C) Stereomicroscopic analysis of roots, including the root tip. Bars = 1 mm.
(D) Comparison of root tip length and thickness of fugu5-1, fugu5-1 ppa1-1, and fugu5-1 ppa2-1 grown on 2% sucrose plates for 9 d. Dv, cell division zone; El, elongation zone. One border was set between the cell division and elongation zones where the first square cortex cell was observed, and the other border was set between the elongation and maturation zone where the first root hair bulge was observed. Error bars indicate se. Asterisks indicate significant differences at *P < 0.05, **P < 0.01, and ***P < 0.001 compared with fugu5-1 (Tukey’s HSD test; R version 3.1.2).
(E) and (F) Complementation of fugu5-1 ppa1-1 by yeast sPPase gene IPP1.
(E) Vertical-plate culture of fugu5-1 ppa1-1 harboring IPP1. The line numbers (#8-3 and #17-3) correspond to the transgenic IPP1 parental lines (AVP1pro:IPP1; Ferjani et al., 2011).
(F) Comparison of root length in mutants grown for 9 d. Asterisks indicate statistically significant differences at ***P < 0.001 (Tukey’s HSD test; R version 3.1.2).
Defect in Hypocotyl Elongation in Multiple PPase Knockout Mutants.
Seeds were grown in the light for 6 h and transferred to the dark.
(A) Image of 4-DAS etiolated seedlings. Bars = 10 mm.
(B) Hypocotyl length in 4-DAS etiolated seedlings grown without sugar (upper) or with 1% sucrose (lower). Different letters above each bar indicate statistically significant differences (P < 0.01, Tukey’s HSD test; R version 3.1.2). n = 13 to 15.
(C) PPi content in 3.5-DAS etiolated seedlings.
(D) Sucrose content in 3.5-DAS etiolated seedlings.
(E) TAG content in 3-DAS etiolated seedlings.
(C) to (E) Data are means and se from three independent experiments. Letters indicate groups divided by Tukey’s HSD test (R version 3.1.2). Asterisks indicate significant differences at *P < 0.05 and ***P < 0.001 compared with the wild type (Student’s t test; R version 3.1.2).
The fugu5s ppa1-1 lines (fugu5-1 ppa1-1, fugu5-3 ppa1-1) showed severe phenotypes (Figure 3; Supplemental Figure 10), although the other mutants grew as well as the wild type on medium containing 1% sucrose. The growth of fugu5s ppa1-1 was markedly suppressed on sucrose-free medium and did not totally recover upon sucrose supplementation (Figures 3A to 3C; Supplemental Figure 10). The rosette leaves of fugu5s ppa1-1 and fugu5s ppa4-1 plants showed symptoms of atrophy, a phenotype that did not recover upon the addition of sucrose.
The roots of fugu5-1 ppa1-1 and fugu5-1 ppa2-1 were short and thick, with an increased number of root hairs (Figure 4), as were the roots of fugu5-3 ppa1-1 and fugu5-3 ppa2-1 (Supplemental Figure 10). The meristematic and elongation zones of fugu5-1 ppa1-1 roots were extremely short; sucrose supplementation in the growth medium enhanced the root phenotype of this double mutant (Figures 4B and 4C). We performed a complementation assay and found that introducing the genomic sequences of PPa1, PPa2, and PPa4 completely rescued the phenotypes of the individual mutants (Supplemental Figure 11). The fugu5-1 ppa1-1 mutant was also rescued by the expression of yeast IPP1 (Figures 4E and 4F). These results suggest that PPa isozymes function as pyrophosphatases in vivo and that their loss of function in the fugu5 background caused the phenotypes mentioned above.
Marked Increases in PPi Accumulation in fugu5 ppa1 Double Mutants
We examined whether the lack of H+-PPase and PPa isozymes further increases PPi levels in plants. The PPi content in the single mutant, fugu5-1 (51.6 nmol/g fresh weight [FW]), was greater than that of the wild type (31.5 nmol/g FW) (Figure 5C), which is consistent with our previous finding (Ferjani et al., 2011). Although the PPi content in ppa1-1 (30.1 nmol/g FW) was comparable to that of the wild type, the PPi content was significantly higher in fugu5-1 ppa1-1 (197 nmol/g FW) than in fugu5-1. These results indicate that H+-PPase is essential for maintaining adequate PPi levels and that the cytosolic PPa isozymes, particularly PPa1, prevent increases in PPi concentrations to toxic levels.
The sucrose content in fugu5 seedlings is low due to suppressed gluconeogenesis (Ferjani et al., 2011; Takahashi et al., 2017). The sucrose content per seedling in fugu5-1 ppa1-1 was lower than in the wild type but slightly higher than in the fugu5-1 single mutant (Figure 5D). Based on fresh weight, the sucrose content was highest in fugu5-1 ppa1-1 and lowest in fugu5-1.
Triacylglycerol (TAG) is the main energy and carbon source in germinating seedlings. In dry seeds, all mutants had normal TAG contents; however, in 3-DAS etiolated seedlings, TAG levels were significantly higher in fugu5-1 ppa1-1 than in the other lines (Figure 5C). These results suggest that excess PPi levels delay the breakdown of TAG.
Morphological Defects in the Cell Walls of fugu5 ppa1 Roots
The severe growth suppression of fugu5s ppa1-1 roots prompted us to investigate morphological changes in mutant root tissues. In plants grown under normal conditions with sucrose, the epidermal and cortex cells of fugu5 ppa1-1 roots were short and swollen, particularly elongating cells (Figure 6E). These characteristics resembled those of wild-type roots treated with the cellulose synthase inhibitor isoxaben (Xu et al., 2008; Figure 6B).
fugu5-1 ppa1-1 Root Tips Exhibit Cell Swelling and Incomplete Cell Walls.
(A) to (F) CLSM analysis of the root elongation zone after staining cell walls with PI/0.25 M sorbitol. Bars = 20 µm.
(G) to (K) TEM images of the division zone in the root tip.
(H), (J), and (K) Enlarged images corresponding to the boxed regions in (G) and (I).
Asterisks indicate incomplete cell walls, and arrows indicate starch grains. CW, cell wall; Ptd, plastid; Nc, nucleus; Vac, vacuole.
Electron microscopy analysis revealed defects in cell wall morphology in fugu5-1 ppa1-1 roots. Parts of the horizontal cell walls in dividing cells appeared to be incomplete (Figures 6I and 6J). These incomplete cell walls were also detected in fugu5-1 ppa1-1 roots by confocal laser scanning microscopy (CLSM) (Figure 6E, asterisks).
In addition, the cell walls of fugu5-1 ppa1-1 roots showed a phenotype that could be interpreted as mechanical weakness. Indeed, when we immersed the roots of fugu5-1 ppa1-1 plants grown on 0.5× Murashige and Skoog (MS) culture medium in water containing propidium iodide (PI), their root tip cells swelled and burst within a few minutes (Figure 7A; Supplemental Movie 1). This bursting occurred after collapse of the inner root cells, which was clearly visible after PI staining (Supplemental Movie 2). When cellulose synthesis was inhibited in wild-type seedlings by 5 nM isoxaben treatment, the root tips of these seedlings were also swollen and damaged but did not burst (Figure 7B; Supplemental Movies 3 and 4). The fugu5-1 and ppa1-1 single mutants were not damaged by water immersion (Figure 7C). To examine whether the cells burst due to low osmotic pressure or other factors, we transferred fugu5-1 ppa1-1 roots from 0.5× MS to PI staining solution containing various concentrations of sorbitol. Damaged PI-stained cells were observed in solutions containing ≤75 mM sorbitol (Figure 7D). Under 50 mM sorbitol treatment, half the area of the root tip was damaged. The osmotic pressure of 0.5× MS medium is ∼80 mOsm. Thus, fugu5-1 ppa1-1 root tips were sensitive to small reductions in osmotic pressure. These results suggest that a defect in cell wall synthesis reduces osmotic stress tolerance in roots.
fugu5-1 ppa1-1 Root Tip Cells Burst Due to Low Osmotic Pressure.
(A) and (B) Time-lapse imaging of water-submerged root tips. Root tips of fugu5-1 ppa1-1 plants (A) grown on 0.5× MS medium with 1% sucrose and wild-type plants (B) treated with 5 nM isoxaben for 3 d on 0.5× MS medium without sucrose and transferred to water containing 10 µM PI. Arrowhead indicates a bulge caused by internal expansion. Arrow indicates a ruptured cell. After treatment, the root tip began to turn brown.
(C) Ratio of the injured versus noninjured area in the root division zone in plants transferred to water containing 10 µM PI.
(D) Ratio of the injured versus noninjured area in fugu5-1 ppa1-1 root tips transferred to various concentrations of sorbitol.
Changes in Cell Wall Composition in fugu5-1 ppa1-1
To further investigate the weakness of fugu5-1 ppa1-1 cell walls, we used three different dyes to stain various cell wall components. The first dye, Direct Fast Scarlet 4BS, also known as Pontamine Fast Scarlet 4B (S4B; CAS no. 3441-14-3), specifically stains cellulose (Anderson et al., 2010). Although S4B cannot penetrate the surfaces of living roots, fixed samples were evenly stained by this dye, including inner root cells. S4B staining was notably weaker in fugu5-1 ppa1-1 compared with the wild type, especially on the outside surface of the epidermis (Figures 8A and 8B), where thick cell walls were observed (Supplemental Figures 12D and 12E), compared with inner root cells (Figures 6G and 6I). Quantification of the intensity of staining showed that fugu5-1 ppa1-1 was stained by S4B at a level only 17.6% that of the wild type. In addition, in another mutant with defective roots, fugu5-1 ppa2-1, the intensity of S4B staining was only 57.1% that of the wild type (Figures 8C and 8D; Supplemental Figure 12).
Staining of Cell Wall Components in the Root Tip.
Plants were grown on 0.5× MS medium containing 1% sucrose.
(A) to (D) CLSM analysis of S4B-stained root elongation zone. Roots of 8 DAS seedlings were fixed and cleared with ClearSee.
(A) and (B) X-Y images of root elongation zones taken 14 μm below the surface.
(C) X-Z images of root elongation zones. Bars = 20 µm.
(D) Quantification of S4B signal intensity between the lateral root cap and epidermis. Values were determined from Figure 8C, Supplemental Figure 12C, and their biological replicates. Error bars indicate se (n = 6). Different letters above each bar indicate statistically significant differences (P < 0.01, Tukey’s HSD test; R version 3.1.2).
(E) and (F) Light microscopy of roots tips stained with Ruthenium Red. The 5-DAS roots were cut into 5-μm-thick sections. Bars = 50 µm.
(G) and (H) CLSM analysis of Aniline Blue-stained root division zones. Roots were fixed and cleared with ClearSee. The images show at the bottom are 3D constructions of the cell plates indicated by arrows. Configuration of the CLSM and staining conditions were the same among samples. Epi, epidermis; LR, lateral root cap; CW, cell wall.
The second dye, Ruthenium Red, specifically stains pectin (Sterling, 1970). In contrast to S4B, Ruthenium Red staining was not significantly reduced in fugu5-1 ppa1-1 versus the wild type (Figures 8E and 8F). The third dye, Aniline Blue, is used to stain β1,3-glucans including callose. Cell plates were clearly stained in the wild type (Figure 8G), whereas fewer stained cell plates, with lower signal intensity, were detected in fugu5-1 ppa1-1 compared with the wild type (Figure 8H). In addition, 3D reconstructions of cell plates showed that callose was unevenly deposited in fugu5-1 ppa1-1 (Figures 8G and 8H, bottom).
To further investigate differences in cell wall composition between fugu5-1 ppa1-1 and the wild type, we examined the monosaccharide composition in alcohol-insoluble residue (AIR) extracted from the tissue, which contains cell wall materials and starch. In this analysis, we used hydroponically grown etiolated seedlings to avoid contamination by agar in the medium. As shown in Figure 9, compared with the wild type, fugu5-1 ppa1-1 had lower ratios of non-starch Glc per total detected monosaccharides (found in cellulose and hemicellulose), GalA, and Gal (found in pectin), Xyl and Man (found in hemicellulose), and GlcA, Fuc, and mGlcA (found in hemicellulose and pectin). By contrast, the levels of Rha and Ara (found in pectin) were higher in futu5-1 ppa1-1 than in the wild type. The results support the finding that fugu5-1 ppa1-1 had less cellulose than the wild type and are consistent with the results of imaging analysis. Imaging analysis revealed no marked difference in pectin (mainly composed of GalA, Gal, Rha, and Ara) content between the wild type and mutant. However, the pectin composition in fugu5-1 ppa1-1 might be altered, with low homogalacturonan and high rhamnogalacturonan content, as indicated by the changes in monosaccharide contents in this mutant (Figure 9).
Monosaccharide Composition in 3-DAS Etiolated Seedlings.
The data are presented as monosaccharide content per total detected monosaccharides in AIR. Glucose (Glc), galacturonic acid (GalA), galactose (Gal), rhamnose (Rha), arabinose (Ara), xylose (Xyl), mannose (Man), glucuronic acid (GlcA), fucose (Fuc), and 4-O-methyl glucuronic acid (mGlcA). Error bars indicate se (n = 3). Different letters above each bar indicate statistically significant differences (P < 0.05, Tukey’s HSD test; R version 3.1.2).
Ectopic Accumulation of Starch in fugu5 ppa1 Cells
In addition to the abovementioned changes, an analysis of monosaccharide composition in AIR revealed a marked increase in starch levels in etiolated fugu5-1 ppa1-1 seedlings compared with the wild type (Figure 9). In addition, transmission electron microscopy (TEM) images of fugu5-1 ppa1-1 roots show small starch grains with high electron density in plastids (Figures 6I and 6K, arrows). To test whether these were starch grains, we stained fugu5-1 ppa1-1 roots with iodine. Columella cells in the root tip contain starch grains, which are involved in gravity sensing. We detected intense iodine staining in wild-type, fugu5-1, and ppa1-1 root tips, particularly in columella cells (Figure 10A). Interestingly, fugu5-1 ppa1-1 roots showed dense staining in the upper meristematic and transition zones, in addition to columella cells. These findings support the notion that the electron-dense grains in TEM images of fugu5-1 ppa1-1 roots are likely starch grains, and they reveal extensive ectopic accumulation of starch in the root cells of the double mutant.
Ectopic Accumulation of Starch Grains in fugu5-1 ppa1-1.
Samples were fixed, cleared with ClearSee (Kurihara et al., 2015), and stained with iodine. Starch grains were stained blue, purple, or red depending on the chain length of amylose (Bailey and Whelan, 1961). All plants were grown in the absence of sucrose.
(A) Root tips of 2-week-old plants subjected to 8 h of illumination prior to fixation.
(B) 3-DAS etiolated hypocotyls.
(C) Iodine staining of third or fourth leaves after 2 h of illumination.
We also observed ectopic starch accumulation in other organs of fugu5-1 ppa1-1. In the wild type, fugu5-1, and ppa1-1, etiolated hypocotyls were stained with iodine, especially in the hook region. In the wild type and ppa1-1, only endodermal cells were stained, whereas fugu5-1 showed starch grains in other cells as well, such as cortical cells. In fugu5-1 ppa1-1 hypocotyls, the staining intensity was extremely high (Figure 10B). By contrast, in the third to fourth leaves of wild-type, fugu5-1, and ppa1-1 plants grown in the dark and then illuminated for 2 h, only a few chloroplasts were slightly stained with iodine. In the third to fourth leaves of fugu5-1 ppa1-1, however, whole leaves, including petioles, were densely stained (Figure 10C). Together, these results reveal high starch synthesis activity in leaves and a shift in energy metabolism from cell proliferation and elongation, including cellulose synthesis, to starch synthesis in the hypocotyl and elongating root cells of fugu5-1 ppa1-1.
DISCUSSION
Tissue Specificity and Abundance of PPa Isozymes
In this study, we established the cytosolic PPa knockout mutants ppa1-1, ppa2-1, ppa4-1, and ppa5-1 and used them to examine the tissue specificity of each isozyme (Supplemental Figure 1). We investigated the presence of these isozymes in wild-type plants by immunoblotting (Supplemental Figure 2) and successfully detected the accumulation of GFP-tagged PPa isozymes expressed under the control of their own promoters (Table 1; Supplemental Figures 4 to 8). PPa1, PPa2, and PPa4 were found in cotyledons, hypocotyls, leaves, roots, and flowers, whereas PPa3 was only found in root hairs and pollen. CLSM analysis clearly revealed the presence of PPa5-GFP in mature epidermal cells in both shoots and roots (Supplemental Figures 5 and 6), but not in cotyledons (Supplemental Figure 4F), emerging rosette leaves (Supplemental Figure 4L), or the root elongation zone (Supplemental Figure 6). The low abundance of PPa5 in these developing tissues might be related to the observation that fugu5s ppa5-1 showed no morphological phenotypes (Figures 3 and 4; Supplemental Figure 10). Furthermore, heterologous expression of these isozymes in yeast cells confirmed the PPi hydrolysis activity of PPa2, PPa3, and PPa5 (Supplemental Figure 9) in addition to PPa1 and PPa4 (Navarro-De la Sancha et al., 2007).
Localization of PPa Isozymes in the Cytoplasm and Nuclei
The cytosolic localization of Arabidopsis PPa isozymes has been examined in plants expressing GFP-PPa (Koroleva et al., 2005; Öztürk et al., 2014; Gutiérrez-Luna et al., 2016). However, the results differ among reports. In our system, all cfGFP-fused PPa isozymes were detected in the cytosol and nucleus (Figure 2). We used a flexible linker, [Gly4Ser]x3, and a cysteine-free monomeric GFP, cfGFP, which prevents the artificial formation of disulfide bonds and noncovalent dimer formation (Suzuki et al., 2012). These PPa-GFP chimeric proteins had enzymatic activity and were localized to their original subcellular compartments, confirming that PPa1-cfGFP and PPa4-cfGFP rescued the fugu5-1 ppa1-1 and fugu5-1 ppa4-1 phenotypes, respectively (Supplemental Figure 3). The dual localization of PPa isozymes in the cytosol and nuclei makes them physiologically effective in hydrolyzing PPi in both intracellular compartments, since the nucleus generates large amounts of PPi during gene transcription and DNA replication.
We will discuss the physiological roles of PPa isozymes based on their tissue specificity and localization in the cytosol and nuclei.
Physiological Importance of PPa Isozymes, as Indicated by Their Knockout Mutants
The fugu5s ppa1-1 mutant showed a severe phenotype in roots, with extremely short main roots, a short, thick meristem region, and numerous root hairs (Figure 4; Supplemental Figure 10). Roots of the fugu5 ppa2-1 mutant showed a similar but milder phenotype compared with fugu5 ppa1-1. These phenotypes might reflect the relative abundance and physiological importance of PPa1 and PPa2 in roots (Figure 1, Table 1; Supplemental Figure 6). Cell type-specific phenotypes, such as abnormal swelling in fugu5-1 ppa1-1 root cells above the elongation zone (Figure 6), might reflect the abundance of PPa1 exclusively in this region (Table 1; Supplemental Figure 6). By contrast, PPa2 expression was limited to the root meristematic zone (Table 1; Supplemental Figure 6), which may explain the moderate phenotype of fugu5-1 ppa2-1 in the root elongation zone (Figure 4), with the absence of cell swelling (Figure 6F) and a gradual increase in S4B staining intensity from the division zone to the elongation zone (Supplemental Figure 12A).
On the other hand, the leaves of fugu5s ppa1-1 and fugu5s ppa4-1 were atrophied (Figure 3; Supplemental Figures 10 and 11). The severe phenotype of fugu5s ppa1-1 leaves points to the physiological importance of PPa1 in this organ. The leaf atrophy of fugu5s ppa4-1 might be related to the extensive expression of PPa4 in young, emerging leaves (Table 1; Supplemental Figure 4). Leaf atrophy has been reported in the fugu5-1, fugu5-3, and vhp1 single mutants grown on ammonium-free medium (Fukuda et al., 2016). In this study, we detected leaf atrophy in both fugu5-1 ppa1-1 and fugu5-1 ppa4-1 seedlings, even when grown on 0.5× MS medium with sufficient levels of ammonium (Figure 3; Supplemental Figure 10). Under the same conditions, fugu5s seedlings did not exhibit leaf atrophy. Therefore, fugu5s ppa1-1 and fugu5s ppa4-1 plants are more sensitive to conditions in the medium than fugu5s. The leaf atrophy in the mutants could be attributed to increased PPi levels, as this phenotype was recovered by the introduction of PPa1 or PPa4 (Supplemental Figure 11). Metabolomics studies of these mutants should provide important information about the biochemical mechanisms underlying leaf atrophy.
H+-PPase Is the Key Scavenging Enzyme of Cytosolic PPi
Previous and current results confirm that all five PPa isozymes have PPi hydrolysis activity. Nevertheless, the ppa1,2,4,5 quadruple mutant showed no visible phenotype (Figures 3 to 5), whereas single knockout mutants of H+-PPase genes (fugu5s and vhp1-1) exhibit clear phenotypes (Figures 3 and 5) (Ferjani et al., 2011; Asaoka et al., 2016; Fukuda et al., 2016). These results indicate that H+-PPase and the other cytosolic PPases fully complemented the lack of PPa1, PPa2, PPa4, and PPa5.
The cytosol of ppa1,2,4,5 seedlings contains at least five types of inorganic PPase: PPa3, VHP1, VHP2;1, VHP2;2, and PS2 (Segami et al., 2010; May et al., 2011). As mentioned above, the PPa3 content is negligible in vegetative tissues, except root hairs (Figure 1, Table 1; Supplemental Figures 6 and 7). PS2 is induced under phosphate-deficient conditions (May et al., 2011), and its physiological contribution may be negligible under normal conditions. The type II H+-PPases, VHP2;1 and VHP2;2, are localized to the Golgi apparatus, and their levels and activities are negligible compared with VHP1 (Segami et al., 2010). Hence, the vacuolar H+-PPase VHP1 might have sufficient activity to complement the lack of the four PPa isozymes.
Conversely, defects in VHP1 alone caused an increase in PPi content (Figure 5C) (Ferjani et al., 2011). The even higher levels of PPi in fugu5-1 ppa1-1 seedlings indicate that PPa1 functions in fugu5-1 to partially compensate for the lack of VHP1. Taken together, these findings indicate that Arabidopsis H+-PPase is the principal enzyme for maintaining adequate cytosolic PPi levels, at least under normal culture conditions.
Notably, the activities of the remaining PPa isozymes in the VHP1 knockout mutant were only partially complemented. The PPi hydrolysis activity of H+-PPase is coupled with H+ pumping; the reaction velocity is only moderate due to its complicated mechanism (Nakanishi et al., 2003). Mung bean (Vigna radiata) H+-PPase has a lower Km for PPi (4.6 μM for Mg-PPi2) and a higher kcat (14 s−1) (Nakanishi et al., 2003) than Arabidopsis PPa1 and PPa4 when expressed in E. coli (93 and 101 μM for PPi; 9.20 and 9.28 s−1, respectively) (Navarro-De la Sancha et al., 2007), indicating that H+-PPase can reduce cytosolic PPi concentrations to a level lower than that achieved by PPa isozymes. Indeed, H+-PPase is abundant, accounting for 10% of total tonoplast proteins in young tissues (Maeshima, 2001). Further experiments aimed at quantifying the levels and activities of PPa isozymes may provide a better understanding of their contributions.
H+-PPase has dual functions, namely, catalyzing PPi hydrolysis in the cytosol and facilitating active translocation of protons into the vacuole. Thus, an adequate concentration of PPi is essential for the full activity of H+-PPase and other PPi-utilizing enzymes. We speculate that plants might maintain cytosolic PPi at adequate levels through the reduction of PPa activity. Cytosolic PPa isozymes may serve as an emergency system for scavenging PPi when present in the cytosol at extremely high levels (Figure 11A).
Models of the Roles of H+-PPase and sPPase in Maintaining PPi Homeostasis and Sugar Metabolism for the Biosynthesis of Sucrose and Cell Wall Components.
(A) Model of the physiological roles of H+-PPase and sPPase in cytosolic PPi homeostasis. The expected cytosolic PPi concentrations in the wild type, ppa1,2,4,5, fugu5, and fugu5 ppa1 are indicated by dotted lines, a triangle (fugu5), and a square (fugu5 ppa1). H+-PPase and sPPases maintain adequate levels of PPi, as shown in the white zone. sPPases may help reduce PPi levels when H+-PPase activity is suppressed, as shown by the red arrow.
(B) Model of sugar metabolism in heterotrophic tissues of fugu5-1 ppa1-1.
Disrupting VHP1 and PPa1 causes a marked increase in cytosolic PPi levels, thereby biasing PPi-dependent bidirectional reactions in the PPi-degrading direction (light blue arrows). However, plastids have their own PPi environment; thus, PPi-dependent reactions including starch synthesis are not affected by increases in cytosolic PPi. Experimentally confirmed increases and decreases in the levels of components are indicated by different colors (red, increase; blue, decrease). 1, pyrophosphate-dependent phosphofructokinase; 2, fructose-1,6-bisphosphatase; 3, phosphofructokinase; 4, UGPase (UDP-Glc pyrophosphorylase); 5, sucrose phosphate synthase; 6, invertase; 7, sucrose synthase; 8, phosphoglucomutase; and 9, AGPase (ADP-Glc pyrophosphorylase).
PPi Accumulation Causes Weakness and Morphological Changes in the Cell Wall
The fugu5-1 ppa1-1 mutant exhibited severe phenotypes in both the shoot and root (Figures 3 and 4), as well as swollen cells with incomplete cell walls in the root elongation zone (Figures 6E, 6I, and 6J). This swelling phenotype mimics the cell wall defect observed in plants treated with isoxaben (Figure 6B) and in prc1/cesA6 (PROCUSTE1/CELLULOSE SYNTHASE6) and rsw1/cesA1 (RADIALLY SWELLEN1/CELLULOASE SYNTHASE1) mutants, which lack cellulose synthase (Baskin et al., 1992; Arioli et al., 1998; Fagard et al., 2000; Desprez et al., 2002). Furthermore, sucrose feeding reinforced the morphological changes in fugu5-1 ppa1-1 roots (Figures 4B and 4C), which also resembled cell wall-deficient mutants fei1 fei2, sos5, prc1, and cob, as well as wild-type plants treated with isoxaben (Xu et al., 2008). Incomplete cell walls (referred to as stubs) have been observed in both the cellulose synthase mutant prc1 and the callose synthase mutant massue/gsl8 (Fagard et al., 2000; Chen et al., 2009; Thiele et al., 2009). The stubs in prc1 form during cell elongation due to the tearing of weak cell walls. The formation of stubs in gsl8 is due to a deficiency in callose, which is essential for cell plate formation during cell division. Young fugu5-1 ppa1-1 cells possess stubs and lack cell plate-forming vesicles (Figures 6I and 6J), indicating they are defective in callose synthesis and cell plate formation. Consistently, Aniline Blue staining revealed poor, incomplete deposition of callose in the cell plates of fugu5-1 ppa1-1 (Figure 8H).
Cell walls provide structural strength to plant cells and determine cell shape. Although plant cells are normally resistant to low osmotic conditions, fugu5-1 ppa1-1 root tips burst under hyperosmotic shock (Figures 7A and 7D; Supplemental Movies 1 and 2), indicating that the cell walls in the double mutant were very weak in the face of increasing turgor pressure. This cell wall weakness was partially mimicked by treatment with isoxaben, an inhibitor of CESA3 and CESA6 trafficking to the plasma membrane (Fisher and Cyr, 1998; Paredez et al., 2006; Tateno et al., 2016) (Figure 7B; Supplemental Movies 3 and 4). The damage was more severe in fugu5-1 ppa1-1 than in isoxaben-treated wild-type plants, and fugu5-1 ppa1-1 roots burst, whereas the root tips of isoxaben-treated wild-type plants did not. Indeed, we detected a notable decrease in cellulose levels in the epidermis of fugu5-1 ppa1-1 (Figures 8A to 8D). The decrease in the non-starch Glc ratio detected in etiolated seedlings also points to reduced cellulose levels in fugu5-1 ppa1-1 (Figure 9). By contrast, isoxaben-treated roots exhibited normal cellulose levels, at least in the epidermis (Figure 8D; Supplemental Figure 12) and therefore did not burst in response to hypotonic conditions (Figure 7B; Supplemental Movies 3 and 4).
Cell wall polysaccharides including callose, cellulose, hemicellulose, and pectin are synthesized from sugar nucleotides. Reactions leading to the biosynthesis of sugar nucleotides from sugar phosphates and nucleotides generate PPi and are mainly detected in the cytosol (Seifert, 2004). Therefore, high cytosolic PPi levels should inhibit sugar nucleotide biosynthesis, particularly in these actively growing cells. However, fugu5-1 ppa1-1 root tips were stained normally by Ruthenium Red (Figures 8E and 8F). TEM analysis revealed that the cell walls of the epidermis and lateral root cap were quite thick (Supplemental Figures 12D and 12E). The changes in monosaccharide composition in the cell walls of etiolated seedlings also indicate that the inhibitory effect of PPi differed among polysaccharides (Figure 9). The substrates of hemicellulose side chains and pectin are transported into the Golgi (Seifert, 2004; Reyes and Orellana, 2008). Therefore, the biosynthesis of these cell wall components is also affected by their affinity for sugar nucleotide transporters. In addition, these substances are concentrated in the Golgi lumen, which may affect their tolerance to high cytosolic PPi concentrations.
Starch Accumulates in Source and Sink Organs in Response to Increased PPi Levels
Ectopic starch grain formation occurred in fugu5-1 ppa1-1 (Figure 10). The relationship between starch and PPi was previously examined by overexpressing sPPase in plants (Sonnewald, 1992; Lerchl et al., 1995; Geigenberger et al., 1998; Meyer et al., 2012; Osorio et al., 2013); however, the starch contents in transgenic plants generated in these studies varied. Theoretically, overexpressing sPPase should reduce starch biosynthesis and increase soluble sugar content due to a decrease in PPi and glucose 6-phosphate (Glc-6P) levels. Specifically, decreased cytosolic PPi levels stimulate a reaction catalyzed by UGPase: Glc-6P + UTP → UDP-Glc + PPi. Reduced Glc-6P levels suppress starch biosynthesis because Glc-6P is essential for this process in plastids (Kammerer et al., 1998). Simultaneously, sucrose is maintained at high levels.
The high starch accumulation in fugu5-1 ppa1-1 can be explained as follows. The metabolic pathways involving sucrose, starch, triacylglycerol, pectin, and cellulose at the heterotrophic phase and in heterotrophic tissues are summarized in Figure 11B. Starch is synthesized from ADP-Glc in plastids. ADP-Glc is synthesized by AGPase: Glc-1P + ATP → ADP-Glc + PPi (Streb and Zeeman, 2012). PPi generated in plastids is hydrolyzed by plastidial PPa6 (Schulze et al., 2004). A deficiency in the genes for either plastidial phosphoglucomutase (pgm) or plastidial AGPase (adg1) results in a starchless or low-starch phenotype in Arabidopsis (Caspar et al., 1985; Lin et al., 1988; Streb et al., 2009). In contrast to plants overexpressing sPPase, the high PPi levels in the cytosol of fugu5-1 ppa1-1 might stimulate the production of Glc-6P by UGPase. Increased Glc-6P levels stimulate starch synthesis in plastids (Figure 11B).
The sucrose concentration was markedly elevated in fugu5-1 ppa1-1, whereas it was reduced in fugu5-1 (Figure 5). High sucrose levels may result from severe growth suppression. High PPi levels suppress plant growth by inhibiting macromolecule biosynthesis (Maeshima, 2001; Ferjani et al., 2011; Takahashi et al., 2017). Under these conditions, sucrose consumption might be markedly reduced in fugu5-1 ppa1-1. This reduced sucrose consumption may exceed the decline in sucrose supply due to the inhibition of gluconeogenesis that occurs in fugu5 (Ferjani et al., 2011; Takahashi et al., 2017). This is a plausible scenario for why the sucrose levels were elevated in fugu5-1 ppa1-1 plants. Sucrose feeding failed to sufficiently restore fugu5s ppa1-1 growth and cellulose levels, which also supports this hypothesis (Figures 3, 5, and 8, Supplemental Figure 10). Taken together, carbohydrate metabolism is suppressed in fugu5s ppa1-1, and increased sucrose levels might stimulate metabolic flow to starch synthesis in plastids, in which PPa6 regulates PPi levels.
In conclusion, cytosolic PPa isozymes are expressed in a tissue- and growth stage-specific manner, with variable abundance. The physiological roles of PPa isozymes are masked by H+-PPase, which is abundant in most plant cells. Our analysis of the severe phenotypes of fugu5 ppa1 mutants unveiled the hidden contribution of PPa isozymes in plants. Finally, dysfunction in the cooperative regulation of PPi by H+-PPase and the PPa isozymes led to a marked increase in PPi levels, which in turn triggered ectopic starch accumulation in both heterotrophic and photosynthetic tissues, as well as incomplete cell wall formation.
METHODS
Plant Materials
Arabidopsis thaliana (accession Columbia-0) seeds, which were provided by the Riken Bioresource Center, were surface-sterilized, placed in the dark at 4°C for 2 d and sown in plates on 0.5× MS medium containing 2.5 mM MES-KOH, pH 5.7, sucrose, and gellan gum (Wako Pure Chemical Industries). To avoid the buildup of excess moisture, the concentration of the gellan gum varied according to sucrose concentration: 0% sucrose with 1% gellan gum, 1% sucrose with 0.75% gellan gum, and 2% sucrose with 0.5% gellan gum.
The plates were incubated at 22°C for 2 to 3 weeks under long-day conditions (16/8-h light/dark cycle with white light fluorescent lamps at ∼90 μmol m−2 s−1). For further analysis, the plants were transferred to soil in pots and grown in a greenhouse at 22°C under long-day conditions. The pots were irrigated twice weekly with Molecular Genetics Research Laboratory (MGRL) medium (Naito et al., 1994) supplemented with 0.5 mM NH4NO3. Prior to analysis, ppa1-1, ppa2-1, ppa4-1, and ppa5-1 were backcrossed three times with the wild type. The T-DNA insertion sites were determined by DNA sequencing (Supplemental Figure 1). To complement fugu5-1 ppa1-1 with IPP1, fugu5-1 ppa1-1 was crossed with AVP1pro:IPP1 (Ferjani et al., 2011), and IPP1+/+fugu5-1−/−ppa1-1−/− plants were selected in the F2 generation.
Vector Construction
The PCR primers used in this study are listed in Supplemental Table 1. To clone PPa isogenes from the Arabidopsis genome, PPa1 (−1985 to +1707 bp), PPa2 (−1876 to +1681 bp), PPa3 (−2008 to +1527 bp), PPa4 (−2047 to +2382 bp), and PPa5 (−884bp to +1685 bp) were amplified from genomic DNA using KOD-plus DNA polymerase (Toyobo). The resulting DNA fragments were subcloned into the EcoRV site of the pZErO 2.1 plasmid (Thermo Fisher Scientific). To construct PPa-cfGFP, the GFP variant cfSGFP2 (Suzuki et al., 2012) was constructed by site-specific mutation of C48S, F64L, C70M, S72A, M153T, V163A, and S175G toward the [Gly4Ser]×2-mGFP (M1A, A207K) of pGWB501-VHP1-mGFP (Segami et al., 2014).
To elongate the N terminus linker, an additional 5-amino acid sequence was added to cfSGFP2_set1_Fw. Three sets of DNA fragments were amplified from [Gly4Ser]x2-mGFP using primers including the mutation shown above and fused by PCR to obtain [Gly4Ser]x3-cfGFP.
To construct PPa1-cfGFP, PPa2-cfGFP, and PPa3-cfGFP, the coding region with its promoter and 3′UTR with the GFP sequence were amplified from pZErO2-PPa individually. [Gly4Ser]x3-cfGFP was conjugated with 20 bp of each PPa genomic fragment by adding the extra sequence to the forward primer. The promoter + coding region, 3′UTR, and pZErO2.1 were digested with the following sets of restriction enzymes: KpnI, XbaI, and SpeI + NotI for PPa1-cfGFP; KpnI, XbaI, and BamHI + XhoI for PPa2-cfGFP and PPa3-cfGFP. These four DNA fragments were fused using an In-fusion HD Cloning Kit (Clontech Laboratories) to obtain each pZErO2-PPa-GFP. The PPa-GFP fusions were excised with KpnI and NotI and subcloned into the pENTR-2B vector (Thermo Fisher Scientific).
To construct PPa4-cfGFP and PPa5-cfGFP, vector conversion of pZErO2-PPa from pZErO2 to pENTR-2B was performed using KpnI and NotI. The coding region, 3′UTR, and linker-cfGFP were amplified and fused by PCR, and the PPa-GFP-3′UTR fragment was inserted into pENTR-PPa using HindIII or ScaI and NotI. The vector portions of the five entry vectors listed above were replaced with the pGWB501 binary vector (Nakagawa et al., 2007) using LR Clonase II (Invitrogen).
Construction of Complementation Vectors
To complement fugu5-1 ppa1-1, fugu5-1 ppa2-1, and fugu5-1 ppa4-1 with PPa1, PPa2, and PPa4 genomic DNA, respectively, the DNA was excised from pZErO2-sPPases with KpnI and NotI as described in the Methods and subcloned into the pENTR-2B vector (Thermo Fisher Scientific). The vector portions of the entry vectors were replaced with pGWB501 binary vectors using LR Clonase II (Invitrogen) to obtain pGWB501-PPa1, pGWB501-PPa2, and pGWB501-PPa4.
Generation of Transgenic Plants
Plants were transformed with Agrobacterium radiobacter strain C58C1 carrying the pGWB501-based binary vectors by the floral dip method (Clough and Bent, 1998). The transformants were grown on 0.5× MS/gellan gum plates containing 7.5 μg mL−1 hygromycin and 0.2 mg mL−1 Cefotax (Chugai Pharmaceutical).
Genotyping
To select sPPase knockout mutants, genomic DNA was extracted from cotyledons (Thomson and Henry, 1995). Specific primer pairs used for selection are shown in Supplemental Table 1. DNA was amplified using KOD-FX Neo (Toyobo).
To select fugu5-1−/− from a population of heterozygous plants of the fugu5-1 genotype, genomic DNA was purified from transgenic plants and amplified using r-Taq DNA polymerase (Toyobo) with specific primers (fugu5-1-Fw and fugu5-1-Rv; Table 1). The amplified DNA was treated with SphI, and fugu5-1−/− lines were selected based on the sizes of the SphI products.
Iodine Staining
Root and hypocotyl samples were fixed in 4% (w/v) paraformaldehyde in PBS for 30 to 120 min under a vacuum at room temperature. Leaf samples were fixed overnight in acetic acid: ethanol at a 1:3 ratio. The solvent was gradually replaced with PBS in an ethanol series. The fixed tissues were washed twice for 1 min in PBS and cleared with ClearSee version 1 (10% [w/v] xylitol, 15% [w/v] sodium deoxycholate, and 25% [w/v] urea) (Kurihara et al., 2015). Sodium deoxycholate has weak decolorizing activity for iodine staining, so the samples were transferred to 10% (w/v) xylitol and 25% (w/v) urea. Finally, the tissues were stained in 2 mM iodine (Wako), 10% xylitol, and 25% urea.
Microscopy
Bright-field and epifluorescence images were obtained using an upright BX51 microscope (Olympus) and captured with a DP72 or DP73 camera (Olympus). For epifluorescence imaging, a U-MNIBA2 or U-MWIG2 filter cube equipped for GFP or PI, respectively, was used. Macro images were observed under a SZ61 stereomicroscope (Olympus) and captured with a DP50 camera (Olympus). CLSM observations were conducted with an upright FV1000-D CLSM (Olympus). The excitation wavelength and transmission range for emission were 473/485 to 560 nm for GFP, 473/650 to 750 nm for chlorophyll, 473/617 to 717 nm for PI, 559/570 to 670 nm for S4B, and 405/425 to 525 nm for Aniline Blue. The images were obtained using Olympus FluoView software and a UPLSAPO60XW water immersion objective or a UPLSAPO 10× objective lens (Olympus). To quantify S4B intensity, the free line region of interest was set between the lateral root cap and the epidermis excluding crushed cells, and the intensity value was recorded. The intensity values were quantified and 3D images of the cell plate were constructed using FluoView software (Olympus).
Expression of GAL1-Driven IPP1 in Yeast
An inducible/repressive IPP1 gene in yeast driven by a GAL1 promoter was constructed as previously reported (Drake et al., 2010). The His3 and GAL1 promoters were amplified from pRS413 (Brachmann et al., 1998) and pYES2 (Thermo Fisher Scientific), respectively. The genomic sequences of the IPP1 upstream region and the IPP1 CDS were amplified from yeast genomic DNA prepared from BY4741 yeast cells (Brachmann et al., 1998). The four DNA fragments were connected by PCR using their complementary sequences with amplifying primers to obtain IPP1upstream-His3-GAL1pro-IPP1. After subcloning into pBluescript-KS(−) and checking the sequences, the DNA was excised with SalI and SacI and transformed into BY4741. The homologous recombinant yTT1, in which the endogenous IPP1 promoter was replaced with the GAL1 promoter, was selected on medium containing galactose without histidine or uracil (−histidine, −uracil, and +galactose media).
Yeast Transformation
The IPP1 promoter (1171 bp) and IPP1 terminator (370 bp) were amplified from yeast DNA and subcloned into pRS416 using SacI and XbaI, and XhoI and KpnI, respectively, to obtain pRS-IPP1pro. The CDS of PPa1-5 and IPP1 were amplified by PCR from Arabidopsis cDNA or yeast genomic DNA and cloned into pRS-IPP1pro using the restriction enzyme pair EcoRI and SalI for PPa1 and IPP1; EcoRI and XhoI for PPa2, PPa3, and PPa5; and EcoRI and HindIII for PPa4. After checking the sequences, the pRS-IPP1pro:sPPases were introduced into yTT1 competent cells by lithium acetate/single-stranded DNA/polyethylene glycol transformation (Gietz and Schiestl, 2007).
SDS-PAGE and Immunoblotting
Crude membrane and soluble fractions were prepared from Arabidopsis plants as described previously (Segami et al., 2014). After centrifugation at 100,000g for 10 min at 4°C, the supernatant was used as the soluble fraction and the pellet was resuspended in a solution of 20 mM Tris-acetate, pH 7.5, 0.25 M sorbitol, 1 mM EGTA-Tris, 1 mM MgCl2, and 2 mM DTT and used as the crude membrane fraction.
Proteins were separated by SDS-PAGE and transferred to an Immobilon-P membrane (EMD Millipore). After treatment with 1% ECL blocking reagent (GE Healthcare) or 3% skim milk, the membrane filter was incubated with primary antibody (1:4000 dilution), followed by horseradish peroxidase-conjugated goat anti-rabbit IgG (1:5000 dilution) (catalog no. 65-6120, lot QE215051; Thermo Fisher Scientific). H+-PPase was detected using a previously prepared antibody against the substrate binding motif of H+-PPase (Segami et al., 2010). To detect PPa isozymes, peptide-specific antibodies were prepared against the common region of Arabidopsis PPa1-PPa5 (C+MPMIDQGEKDDKII) and used as anti-PPa antibodies. Commercially available anti-GFP (catalog no. ab290; lot GR3184825-1; Abcam) and anti-IPP1 (catalog no. AP21326BT-N; lot 6002; Acris Antibodies) antibodies were used for detection of GFP and IPP1, respectively. The chemiluminescence reagent ECL or ECL-prime (GE Healthcare) was used for antigen detection. Chemiluminescence was detected with a Light-Capture II imaging device with a cooled CCD camera (Atto).
2D-PAGE
For isoelectric-focusing, samples were frozen in liquid nitrogen and homogenized with a mortar and pestle. After the addition of chilled acetone (1 mL) containing 10% trichloroacetate and 20 mM DTT, the homogenate was chilled at −20°C for 1 h. After centrifugation for 10 min, the precipitate was washed twice with acetone containing 20 mM DTT and incubated at −20°C for 1 h. The precipitate was dried in an evaporator for 20 min and melted in a 50-fold volume of 7 M urea, 2 M thiourea, 3% CHAPS, 1% Triton X-100, and 20 mM DTT. The sample was subjected to ultracentrifugation at 40,000 rpm and 4°C for 8 min in an SA45 rotor (Hitachi). After adding a one-tenth volume of 1 M Tris-HCl, pH 9.5, and 1 M acrylamide, the supernatant was subjected to A-M58 IEF agarGEL (pH range 5−8; Atto) electrophoresis at a constant voltage of 360 V for 160 min. After separation, the gel was fixed in 10% trichloroacetic acid for 3 min, washed three times with distilled water, and incubated in distilled water for 120 min. The gel was equilibrated with 50 mM Tris-HCl, pH 6.8, 2% SDS, and 0.001% bromophenol blue for 10 min with gentle agitation, followed by SDS-PAGE.
Quantification of PPi and Sucrose
Approximately 200 to 300 etiolated seedlings at 3.5 DAS were homogenized in 0.2× volume of 3 M HClO4 on ice with a BioMasher II (Nippi). The homogenate was incubated on ice for 15 min to precipitate proteins and centrifuged at 20,000g for 10 min at 4°C. The supernatant was transferred to a fresh tube, along with 25 mg of the metal scavenger SiliaMetS TAAcOH (SiliCycle) and gradually neutralized with KOH. The sample was centrifuged at 20,000g for 10 min at 4°C to remove KClO4 and SiliaMetS. The supernatant was passed twice through a Monospin C18 column (GL Sciences). The flow-through fraction was diluted with water and filtered through a MILLEX-GV 0.22-μm filter (Millipore). PPi and sucrose levels were measured by ion chromatography with a DX-500 system (Dionex) using an IonPac AS11 column (Dionex) for PPi and a CarboPac PA1 column (Dionex) for sucrose.
Quantification of Total TAG
Total TAG levels were quantified as described previously, with minor modifications (Ferjani et al., 2011). Either 20 dry seeds or 20 etiolated seedlings at 3 DAS were homogenized in a mortar or BioMasher II homogenizer (Nippi) with 50 μL distilled water. The homogenates were combined with 0.5 mL triglyceride E-Test (Wako) and incubated at 37°C for 5 min. The supernatants (200 μL) were transferred to 96-well plates, and absorbance at 600 nm was quantified with an Enspire 2300 plate reader (Perkin-Elmer).
Cell Wall Staining
Arabidopsis plants were grown on MS plates containing 1% sucrose for 5 to 8 d, and the roots were fixed overnight in FAA (50% ethanol, 5% acetic acid, and 4% formaldehyde) at room temperature. The samples were gradually rehydrated through an ethanol series, washed with PBS, and cleared with ClearSee (Kurihara et al., 2015). For S4B staining, samples were stained with 0.01% S4B in xylitol/urea/NaCl (10% xylitol, 25% urea, and 150 mM NaCl) for 30 min, washed with xylitol/urea/NaCl, and observed via CLSM. For Aniline Blue staining, the rehydrated samples were stained with 0.05% decolorized Aniline Blue, 54 mM K3PO4 (pH 11), 10% xylitol, and 25% urea for 1 h and observed by CLSM. Aniline Blue decolorization was performed according to Mori et al. (2006). For Ruthenium Red staining, samples fixed in FAA were dehydrated through an ethanol series, embedded in Technovit 7100 (Kulzer and Co.) following the manufacturer’s protocol, and sectioned with a microtome (RM2125 RTS; Leica Microsystems) to a thickness of 5 μm as described previously (Maeda et al., 2014). The histological sections were stained with 0.02% Ruthenium Red for 30 min, washed with water for 30 s, and examined using a light microscope (DM6 B; Leica Microsystems).
Sugar Composition Analysis
AIR and monosaccharide composition analyses were performed as described previously (Sakamoto and Mitsuda, 2015; Sakamoto et al., 2015). Starch content was determined using the modified protocol of the Resistant Starch Assay Kit (Megazyme). Starch in powdered AIR (2.00‒3.00 mg) was digested with 500 units mL−1 α-amylase from porcine pancreas (Megazyme) and 0.33 units mL−1 amyloglucosidase from Aspergillus niger (Megazyme) at 37°C for 18 h. Liberated glucose, reflecting starch levels, was measured using a Glucose C II Test Kit (Wako).
Statistical Analysis
To quantify fresh weight, length, thickness, and concentration, data from at least three independent experiments were averaged, and the values were subjected to statistical analyses using Student’s t test, or Tukey’s honestly significant difference (HSD) test (R version 3.1.2; R Core Team, 2014). For ratio quantification, the values were subjected to statistical analysis using the prop test (R version 3.1.2; R Core Team, 2014).
Accession Numbers
Sequence data from this article can be found in the GenBank/EMBL/DDBJ libraries under the following accession numbers: Arabidopsis VHP1 (AT1G15690.1), PPa1 (AT1G01050.1), PPa2 (AT2G18230.1), PPa3 (AT2G46860.1), PPa4 (AT3G53620.1), PPa5 (AT4G01480.1), PPa6 (AT5G09650.1), VHP2;1 (AT1G78920), VHP2;2 (AT1G16780), and PS2/PPSPASE1 (AT1G73010).
Supplemental Data
Supplemental Figure 1. Identification of T-DNA insertional mutants.
Supplemental Figure 2. Immunological detection of PPase isozymes.
Supplemental Figure 3. GFP-fused pyrophosphatases complement the double mutant phenotypes.
Supplemental Figure 4. Tissue-specific accumulation of PPa-GFP in seedlings.
Supplemental Figure 5. Intracellular localization of PPa-GFP in leaves.
Supplemental Figure 6. Cell-specific and intracellular localization of PPa-GFP in roots.
Supplemental Figure 7. Epifluorescent microscopy images of 2-DAG etiolated seedlings expressing PPa1, PPa2, PPa4, PPa3, PPa5, and VHP1 fused with GFP.
Supplemental Figure 8. Distribution of PPa isozymes in flowers.
Supplemental Figure 9. Complementation of IPP1-defective yeast strain yTT1 by Arabidopsis PPa.
Supplemental Figure 10. Growth phenotypes of double mutants of fugu5-3 with ppa1-1, ppa2-1, ppa4-1, and ppa5-1.
Supplemental Figure 11. Complementation of fugu5-1 ppa1-1, fugu5-1 ppa2-1, and fugu5-1 ppa4-1 by PPa1, PPa2, and PPa4.
Supplemental Figure 12. Complementary data for Figure 8.
Supplemental Table 1. List of oligonucleotide primers used in this study.
Supplemental Movie 1. Time-lapse imaging of a water-submerged fugu5-1 ppa1-1 root tip.
Supplemental Movie 2. Time-lapse imaging of PI staining.
Supplemental Movie 3. Time-lapse imaging of a water-submerged wild-type root tip treated with 5 nM isoxaben for 3 d.
Supplemental Movie 4. Time-lapse imaging of PI staining.
Acknowledgments
We thank Yoichi Nakanishi and Miki Kawachi (Nagoya University, Japan) for their valuable advice. This work was supported by JSPS KAKENHI Grants 26252011 and 26113506 to M.M. and 2816K18687 to S. Segami and by a Grant-in-Aid for Scientific Research (B) (16H04803 to A.F.).
AUTHOR CONTRIBUTIONS
S. Segami and M.M. designed the experiments and wrote and finalized the manuscript. S. Segami established the knockout mutant lines and conducted the phenotyping and construction of PPa4-cfGFP and PPa5-cfGFP, microscopy observations, quantification of PPi and sucrose, and biochemical and physiological analyses. T.T. constructed PPa1-cfGFP, PPa2-cfGFP, and PPa3-cfGFP and performed the yeast experiments and IPP1 complementation assays. M.F. performed the physiological analyses. S. Segami and S.G. performed sectioning and cell wall staining. S. Sakamoto and N.M. performed the sugar composition analysis. S.K. improved the iodine staining method. A.F. provided the fugu5s mutants and AVP1pro:IPP1 transgenic lines and finalized the manuscript.
- Received December 1, 2017.
- Revised April 5, 2018.
- Accepted April 23, 2018.
- Published April 24, 2018.
References
