Constitutive and Companion Cell-Specific Overexpression of AVP1, Encoding a Proton-Pumping Pyrophosphatase, Enhances Biomass Accumulation, Phloem Loading, and Long-Distance Transport^,

H⁺-pumping pyrophosphatases appear to function in companion cells to increase phloem loading and long-distance transport, leading to more robust plants with enhanced growth.

Abstract

Plant productivity is determined in large part by the partitioning of assimilates between the sites of production and the sites of utilization. Proton-pumping pyrophosphatases (H⁺-PPases) are shown to participate in many energetic plant processes, including general growth and biomass accumulation, CO₂ fixation, nutrient acquisition, and stress responses. H⁺-PPases have a well-documented role in hydrolyzing pyrophosphate (PPi) and capturing the released energy to pump H⁺ across the tonoplast and endomembranes to create proton motive force (pmf). Recently, an additional role for H⁺-PPases in phloem loading and biomass partitioning was proposed. In companion cells (CCs) of the phloem, H⁺-PPases localize to the plasma membrane rather than endomembranes, and rather than hydrolyzing PPi to create pmf, pmf is utilized to synthesize PPi. Additional PPi in the CCs promotes sucrose oxidation and ATP synthesis, which the plasma membrane P-type ATPase in turn uses to create more pmf for phloem loading of sucrose via sucrose-H⁺ symporters. To test this model, transgenic Arabidopsis (Arabidopsis thaliana) plants were generated with constitutive and CC-specific overexpression of AVP1, encoding type 1 ARABIDOPSIS VACUOLAR PYROPHOSPHATASE1. Plants with both constitutive and CC-specific overexpression accumulated more biomass in shoot and root systems. ¹⁴C-labeling experiments showed enhanced photosynthesis, phloem loading, phloem transport, and delivery to sink organs. The results obtained with constitutive and CC-specific promoters were very similar, such that the growth enhancement mediated by AVP1 overexpression can be attributed to its role in phloem CCs. This supports the model for H⁺-PPases functioning as PPi synthases in the phloem by arguing that the increases in biomass observed with AVP1 overexpression stem from improved phloem loading and transport.

Plant productivity is largely determined by the acquisition and partitioning of organic carbon and other nutrients among the various organs of the plant body. Suc is the principal form of organic carbon transported long distance via the phloem vascular system. In the initial steps of carbon partitioning, Suc enters the phloem companion cell (CC)-sieve element (SE) complex to generate a hydrostatic pressure gradient that pushes water and dissolved compounds from source to recipient sink organs. Phloem loading from the apoplasm is the best described method for accumulating solute in to the CC-SE complex. Suc generated in mesophyll cells moves cell to cell, presumably via plasmodesmata, to enter the vascular bundle (Giaquinta, 1983). Efflux to the apoplasm occurs from phloem parenchyma cells through the SWEET transporters AtSWEET11 and AtSWEET12 (Chen et al., 2012). From the apoplasmic space, Suc is then actively accumulated into the CC-SE complex by Suc transporter/carrier proteins (SUTs or SUCs) energized by the proton motive force (pmf; Lalonde et al., 2004; Ayre, 2011).

Although phloem loading from the apoplasm is mechanistically well understood, little is known about how the process is regulated. SUT genes that participate in phloem loading are transcriptionally regulated by Suc concentrations (Vaughn et al., 2002), leaf maturation (Riesmeier et al., 1993; Truernit and Sauer, 1995), diurnal cycles (Kühn et al., 1997), and possibly pressure (Smith and Milburn, 1980; Aloni et al., 1986). Posttranslational control may be mediated by protein sorting and protein-protein interactions (Krügel et al., 2008; Liesche et al., 2010). Overexpression of SUT genes specifically in CCs to enhance loading and carbon partitioning to sink organs has been proposed as a mechanism to improve plant productivity (Ainsworth and Bush, 2011; Braun et al., 2014). In a test of this hypothesis, evidence for enhanced loading and partitioning was obtained in Arabidopsis (Arabidopsis thaliana), but plants were stunted, apparently because improved Suc transport disrupted nutrient homeostasis in the sink organs (Dasgupta et al., 2014).

Further regulation of phloem loading and partitioning may come from Suc availability in the apoplasm or from the strength of the pmf required for Suc uptake, which is generated by plasma membrane (PM)-localized, P-type ATPases. The CCs of species that load from the apoplasm have high levels of ATPase gene expression (DeWitt and Sussman, 1995) and greater pmf than the surrounding phloem parenchyma (Hafke et al., 2005). To generate the ATP required by the ATPases, a portion of the loaded Suc must be oxidized. A morphological hallmark of CCs is that they are cytoplasmically dense with many mitochondria, which may be cell-specific adaptations to accommodate this metabolism (van Bel and Knoblauch, 2000). In principle, complete oxidation of a single molecule of Suc can generate approximately 60 ATPs and thus provide pmf to load approximately 60 molecules of Suc.

During the initial steps of Suc oxidation in CCs, sucrose synthase (SUS) splits Suc into Fru and UDP-Glc, and then UDP-Glc pyrophosphorylase (UGPase) catalyzes UDP-Glc + pyrophosphate (PPi) ↔ Glc-1-P + UTP (Yang and Russell, 1990; Nolte and Koch, 1993). Genetic evidence for this pathway derives in part from strong SUS expression in CCs. Biochemical support derives from CC-specific expression of an Escherichia coli pyrophosphatase, ppaI, which hydrolyzes PPi to prevent the production of Glc-1-P and UTP and results in the stunted growth characteristic of deficient phloem transport. Importantly, phloem-specific expression of a yeast invertase (rolC_pro:SUC2) restored wild-type phenotypes by allowing Suc to enter glycolysis through a SUS- and PPi-independent pathway (Lerchl et al., 1995).

In addition to P-type ATPases, plants possess membrane-bound H⁺-pumping pyrophosphatases (H⁺-PPases) that have a well-established role in capturing the energy of PPi hydrolysis by pumping protons across the tonoplast or other endomembrane to contribute to pmf (Zhen et al., 1997; Baltscheffsky et al., 1999; Maeshima, 2000). PPi is a by-product in the synthesis of many biopolymers, including proteins, nucleic acids, starch, and cell wall carbohydrates, and is thus a convenient source of energy in metabolically active cells. Prior work established that type I H⁺-PPase is broadly expressed, with highest levels in rapidly growing and vascular tissues (Li et al., 2005; Yang et al., 2007; Pizzio et al., 2015).

While the activity of an E. coli soluble PPase in the cytoplasm results in debilitated plants, overexpression of H⁺-PPases results in plants with greater salt and drought tolerance, improved nutrient uptake, and greater biomass accumulation (Li et al., 2005, 2008, 2010; Park et al., 2005; Yang et al., 2007; Bao et al., 2008; Lv et al., 2008, 2009; Schilling et al., 2014). In Arabidopsis, ARABIDOPSIS VACUOLAR PYROPHOSPHATASE1 (AVP1) overexpression resulted in 40% to 60% greater leaf area and up to 8.4-fold increase in root growth, relative to the wild type (Li et al., 2005; Gonzalez et al., 2010). Similar increases in biomass were observed with AVP1 and orthologs from different species overexpressed in a diversity of plants, including rice (Oryza sativa), cotton (Gossypium hirsutum), maize (Zea mays), alfalfa (Medicago sativa), wheat (Triticum aestivum), barley (Hordeum vulgare), and creeping bentgrass (Agrostis stolonifera; Brini et al., 2007; Bao et al., 2008; Li et al., 2008, 2010; Lv et al., 2008, 2009; Pasapula et al., 2011; Schilling et al., 2014). Overexpressing H⁺-PPase genes also enhances nutrient uptake by affecting the abundance and activity of the PM H⁺-ATPase in a manner consistent with apoplasmic pH alterations and rhizosphere acidification (Li et al., 2005; Yang et al., 2007). Consequently, plants overexpressing AVP1 outperform controls when grown under phosphate and nitrate limitations and accumulate more potassium (Yang et al., 2007; Undurraga et al., 2012; Paez-Valencia et al., 2013). In addition, the metabolite profile in Arabidopsis overexpressing AVP1 is shifted toward nitrogen metabolism and changes in carbon signaling (Gonzalez et al., 2010).

It is proposed that H⁺-PPase overexpression improves this diversity of physiological attributes because it localizes to different membranes and has different functions depending on the cells in which it is being expressed (Paez-Valencia et al., 2011; Gaxiola et al., 2012; Pizzio et al., 2015). Although first characterized as a vacuolar H⁺-PPase (Rea et al., 1992; Maeshima, 2000), studies with gold-conjugated H⁺-PPase-specific antibodies and proteomic approaches show a dual localization at the tonoplast and the PM (Long et al., 1995; Robinson et al., 1996; Langhans et al., 2001; Alexandersson et al., 2004; Paez-Valencia et al., 2011; Regmi et al., 2015). Although dual localization is well supported, a mechanistic model for this alternative targeting has not been put forward.

It was suggested that PM-localized H⁺-PPase could contribute to phloem pmf to help energize phloem loading and enhance long-distance transport. Although this is an attractive model, H⁺-PPases on the CC PM cannot thermodynamically operate hydrolytically to pump H⁺ into the apoplasm (Davies, 1997), based on estimations of the free energy of the H⁺-PPase pump action for in vivo conditions. Instead, it is argued that the reverse reaction, in which the PM pmf is used to synthesize PPi, is thermodynamically feasible (Davies, 1997). Therefore, it is highly noteworthy that in vivo data obtained with the H⁺-PPase from the gram-negative proteobacterium Rhodospirillum rubrum are consistent with the capacity of this enzyme to play two distinct roles depending on location and conditions: it can act as an intracellular H⁺ pump in the acidocalcisomes (Seufferheld et al., 2004) or as a PPi synthase in the chromatophore membranes during illumination (Baltscheffsky et al., 1966, 1999). Furthermore, Rocha Facanha and de Meis (1998) presented in vitro evidence with tonoplast fractions of maize coleoptiles and seeds consistent with the reverse function of the H⁺-PPase and suggested that the H⁺-PPase could operate in the maintenance of cytosolic PPi levels. Supported by these studies, it was proposed that H⁺-PPases localized to the PM of CCs could use pmf to maintain the cytosolic PPi levels required for Suc respiration via SUS, UGPase, and PPi-dependent phosphofructokinase (see figure 1 in Gaxiola et al., 2012). A recent report showed evidence consistent with a PPi synthase function for PM-localized AVP1 in Arabidopsis CCs, with direct implications in phloem function (Pizzio et al., 2015).

To directly test for an AVP1 function in phloem loading and transport, transgenic Arabidopsis plants were generated with constitutive and CC-specific overexpression of AVP1. Plants with both expression patterns accumulated more biomass in shoot and root systems. ¹⁴C-labeling experiments showed enhanced photosynthesis, phloem loading, phloem transport, and delivery to sink organs. Changes in the levels of key metabolites that report the carbon status of the plants were sought but not identified, suggesting that steady-state levels are not altered. Since constitutive and CC-specific expression patterns gave very similar results, this work supports the model for a noncanonical H⁺-PPase function in the phloem by arguing that the increases in biomass observed with AVP1 overexpression stem from improved phloem loading and transport.

RESULTS

Overexpression of AVP1 Enhances Plant Growth

To dissect the roles of AVP1 in enhancing biomass accumulation and carbohydrate distribution, transgenic Arabidopsis plants were generated with constitutive AVP1 complementary DNA (cDNA) overexpression from the cauliflower mosaic virus 35S promoter (35S_pro) and phloem CC-specific expression from a promoter element derived from Commelina yellow mottle virus (CoYMV_pro). CoYMV_pro has been shown to have strong CC-specific expression (Medberry et al., 1992; Matsuda et al., 2002). For each construct, 25 to 30 independently transformed lines were characterized for vegetative rosette growth at 16 d post germination. Three lines for each construct that showed increases in rosette area, relative to the wild type, were selected for further study. From those constitutively expressing AVP1 cDNA from 35S_pro, these were 35S-9, 35S-30, and 35S-31; and among the lines with CC-specific overexpression from CoYMV_pro, these were CoY-41, CoY-50, and CoY-68. These lines were briefly described previously (Pizzio et al., 2015), and their enhanced growth on potting mix is elaborated in Supplemental Figure S1 (for shoot and root growth in hydroponic conditions, see Fig. 6, A and B, below). In addition, AVP1-1, which is a well-described 35S_pro:AVP1-overexpressing line (Li et al., 2005; Yang et al., 2007; Pizzio et al., 2015), was included in all analyses as an additional control (Fig. 1). Growth rate analysis showed that the transgenic lines grew faster (Supplemental Fig. S1C) than the wild type.

Figure 6.

Biomass accumulation and steady-state analysis of prominent metabolites of central metabolism in rosettes and roots grown hydroponically. A and B, Fresh weights of shoots (A) and roots (B) from hydroponically grown 22-d-old plants of the indicated lines. C and D, Principal soluble sugars, Glc, Fru, and Suc, as indicated in the key, from shoots (C) and roots (D). E and F, Malate and fumarate, as indicated in the key, from shoots (E) and roots (F). G and H, Glc-6-P (G6P) from shoots (G) and roots (H) of the indicated lines. I to L, Total amino acids from shoots (I) and roots (J), and total protein from shoots (K) and roots (L), of the indicated lines. M, Starch from the rosettes of the indicated lines; starch in roots was below the level of detection. The y axis indicates the units and concentration of each metabolite, relative to fresh weight. All metabolites were measured from the same extracts; n = 6 pools, each consisting of four rosettes or four root systems, and variation is expressed as se. Significant differences from the wild type (WT) are based on Student’s t test: ^, P ≤ 0.07; *, P ≤ 0.05; **, P ≤ 0.01; and ***, P ≤ 0.001. FW, Fresh weight.

Figure 1.

Immunolocalizations of AVP1 source-leaf phloem, and AVP1 genomic and transgene transcript abundance. Top, Reverse transcription (RT)-quantitative PCR (qPCR) results of AVP1 expression from the genomic locus (white portion of the bars) and cDNA driven by 35S_pro (dark gray) and CoYMV_pro (light gray), relative to GAPDH3 expression from shoots of potting mix-grown samples; n = three biological repetitions, each averaged from two technical repetitions; variation is expressed as se of the biological repetitions. Transgene levels in 35S_pro lines (dark gray bars) were reported previously (Pizzio et al., 2015). WT, Wild type. Bottom, Immunolocalization of AVP1 in leaf sections of an empty-vector control line and two representative CoYMV_pro:AVP1 lines. a, Empty vector control; b, CoYMVpro:AVP1-54 (CoY-54); c, CoYMV_pro:AVP1-68 (CoY-68); 1 and 2 are midrib large veins, and 3 and 4 are leaf-lamina small veins; 1 and 3 odd numbers are preimmune sera negative controls, and 2 and 4 even numbers are H⁺-PPase sera experimentals. Xylem (yellow arrows) and phloem (red arrows) regions of the veins are labeled. Compared with the empty vector control, CoY-54 and CoY-68 have higher AVP1 levels in leaf midrib and lamina veins. Bars in 1 and 2 = 50 μm, and bars in 3 and 4 = 10 μm.

To establish the AVP1 transcript levels, RT-qPCR was conducted on total RNA isolated from the wild type and each transgenic line. Oligonucleotides were designed for specific quantification of total AVP1 transcript and transgene-derived transcript and were standardized relative to the housekeeping gene GAPDH3 (Fig. 1). In the representative lines, expression levels varied as expected, but in all cases they exceeded wild-type levels. Relative to AVP1 transcript levels in wild-type plants, endogenous AVP1 transcript declined approximately 50% in the 35S_pro:AVP1 lines and increased moderately in the CoYMV_pro:AVP1 lines. Immunohistochemistry was conducted to confirm phloem-specific overexpression in two of the CoYMV_pro:AVP1 lines (Fig. 1). AVP1 protein was detected in the phloem and other cells of the vascular bundle of wild-type controls as expected (Paez-Valencia et al., 2011) but was enhanced substantially in the phloem of transgenic lines in both the midrib and the smaller veins of the leaf.

Overexpression of AVP1 Influences Carbon Partitioning

We hypothesize that AVP1 overexpression influences carbon flux through primary metabolism and partitioning through the plant. To assess the effect of AVP1 overexpression on Suc synthesis, the major nonstructural carbohydrates (Glc, Fru, Suc, and starch) were quantified in rosettes of the wild type and the representative transgenic lines at 21 d post germination (Fig. 2). All plants were harvested into liquid nitrogen 6 to 8 h into the 12-h illuminated period. Relative to the wild type, the representative 35S_pro lines did not show consistent alterations in nonstructural carbohydrates, but line AVP1-1 did accumulate more starch, consistent with prior reports (Pizzio et al., 2015). In contrast, the CoYMV_pro:AVP1 lines consistently had more hexose and more starch (Fig. 2, A and B). Soluble carbohydrates were measured enzymatically and by high-performance anion-exchange chromatography (data not shown), and the results were consistent with the two techniques.

Figure 2.

Steady-state analysis of prominent metabolites of central metabolism in rosettes. A, Principal soluble sugars, Glc, Fru, and Suc, as indicated in the key, for the indicated lines. B, Starch levels (expressed as Glc equivalents) in the indicated lines. C, Chlorophyll a and b, as indicated in the key, in the indicated lines. D, Malate and fumarate, as indicated in the key, for the indicated lines. E and F, Total amino acids (E) and total protein (F) in each line as indicated. G and H, Inorganic phosphate (P_i; G) and total phosphate (H) in each line as indicated. I, Glc-6-P (G6P) in the indicated lines. The y axis indicates the units and concentration of each metabolite, relative to fresh weight. All metabolites were measured from the same extracts; n = 6 pools, each consisting of four rosettes, and variation is expressed as se. Significant differences from the wild type are based on Student’s t test: ^, P ≤ 0.07; *, P ≤ 0.05; **, P ≤ 0.01; and ***, P ≤ 0.001. FW, Fresh weight.

In addition, a broader collection of prominent carbon-containing compounds were measured in rosettes to assess the overall carbon status of the plants. Levels of chlorophyll A and B were measured as metrics for overall plant health and photosynthetic capacity, and no alterations were observed (Fig. 2C). No significant and consistent changes were observed in the major organic acids malate and fumarate or in protein and total amino acid levels (Fig. 2, D–F). It was shown previously that AVP1 overexpression can enhance nutrient uptake, including the uptake of phosphate (Yang et al., 2007, 2014; Gaxiola et al., 2012; Pizzio et al., 2015). Thus, inorganic phosphate and total phosphate were measured. Some lines had significantly more or less phosphate, but a consistent pattern was not evident (Fig. 2, G and H). Phosphorylated intermediates, represented by Glc-6-P, showed moderate increases in two representative 35S_pro:AVP1 lines but not the third; levels were also increased in the CoYMV_pro:AVP1 line that was tested (Fig. 2I).

Since 35S_pro:AVP1 and CoYMV_pro:AVP1 plants showed increases in rosette biomass and area (Supplemental Fig. S1), photosynthetic rates of 21-d-old plants were quantified by infrared gas analysis using a Li-Cor Li-6400XT photosynthesis monitoring system (Fig. 3). Average photosynthetic rates were determined relative to rosette surface area. Measurements were done 4 to 6 h into the light period under conditions similar to the growth environment. Average photosynthetic rates were greater among the transgenic lines tested, relative to the wild type, but only AVP1-1 was statistically significant at a P = 0.05 cutoff (Fig. 3A). Infrared gas analysis provides instantaneous photosynthetic rates, and rate increases not exceeding sample variation may account for the larger rosette size and biomass of the transgenic plants. In an alternative approach to measure primary productivity, photosynthetic labeling with ¹⁴CO₂ was conducted over a 20-min period among seedlings growing on vertical plates in sterile medium, with results standardized relative to shoot mass. These experiments showed enhanced assimilation among the transgenic lines relative to the wild type (Fig. 3, B and C). Since CoYMV_pro is specific to CCs, enhanced ¹⁴C assimilation most likely stems from increased photosynthesis indirectly triggered by improved phloem transport.

Figure 3.

Photosynthetic carbon fixation in wild-type (WT) and transgenic lines. A, Net photosynthesis rates as measured by infrared gas analysis on 21-d-old whole rosettes; n = 12 plants for each line. B and C, Photosynthetic labeling with ¹⁴CO₂ for a 20-min pulse and a 40-min chase of wild-type control and 35S_pro:AVP1 (B) and CoYMV_pro:AVP1 (C) lines as indicated; n = 3 to 6 pools of two plants, and variation is expressed as se. Significant differences from the wild-type are based on Student’s t test: *, P ≤ 0.05. FW, Fresh weight.

AVP1 Overexpression Enhances Phloem Loading and Long-Distance Transport

We previously put forward a model in which AVP1 in CC localizes to the PM and uses pmf to function as a synthase to increase pyrophosphate levels (Fuglsang et al., 2011; Paez-Valencia et al., 2011; Gaxiola et al., 2012; Pizzio et al., 2015). In this model, increased pyrophosphate levels promote Suc oxidation for ATP synthesis, which in turn enhances pmf across the PM to energize phloem loading. Complete oxidation of one molecule of Suc may yield up to approximately 60 ATPs, and a small increase in Suc oxidation can dramatically increase CC membrane potential for improved loading. Earlier work showed that AVP1 overexpression enhances pmf, since 35S_pro:AVP1 lines exude more H⁺ to the rhizosphere than the wild type (Pizzio et al., 2015). Earlier work also showed that the initial velocity and steady state of the pmf H⁺ gradients generated by the P-type H⁺-ATPase (i.e. ATP dependent and sensitive to 0.1 mm Na₃VO₄) exhibited values significantly greater (approximately 50%–70%; P = 0.05) in 35S_pro:AVP1-overexpressing lines compared with the wild type (Undurraga et al., 2012).

Several experiments were conducted to test for enhanced phloem loading and long-distance transport of Suc. To directly test the phloem-loading capacity of plants overexpressing AVP1, the uptake of [¹⁴C]Suc into the leaf veins was studied (Fig. 4). Leaf discs from source leaves were vacuum infiltrated with a buffered solution containing [¹⁴C]Suc, incubated for 20 min, and then washed thoroughly to remove [¹⁴C]Suc from the apoplasm. The washed discs were blotted dry, frozen on dry ice, and lyophilized. Autoradiography confirmed that the majority of ¹⁴C label was concentrated in the veins (Supplemental Fig. S2). Scintillation counting showed that all the 35S_pro:AVP1 and CoYMV_pro:AVP1 lines absorbed significantly more [¹⁴C]Suc, indicating that the transgenic lines have enhanced phloem loading compared with the wild type (Fig. 4A).

Figure 4.

Phloem loading and transport in excised plants. A, Uptake of [¹⁴C]Suc into source leaf discs of wild-type (WT), 35S_pro:AVP1, and CoYMV_pro:AVP1 plants to test phloem-loading capacity; n = 8 pools of three discs, randomized from different plants. B, Phloem exudation from detached rosettes of soil-grown plants into EDTA-containing solution after photosynthetic labeling with ¹⁴CO₂ shows improved phloem transport of ¹⁴C relative to the wild type. The first 20 min of exudate was discarded, since this would contain the contents of cut cells. The black bars represent cpm mg⁻¹ fresh weight (FW) h⁻¹ in the exudate of the first subsequent hour, and the white bars represent exudate from the second subsequent hour; n = 9 exudations for each line (three of each were labeled in three labeling chambers; see “Materials and Methods”). C, Day and night total Suc phloem exudation rates from mature source leaves of 40-d-old potting mix-grown wild-type (white bars) and AVP1-1 (black bars) plants; n = 4 exudations from 15 pooled leaves per line per time point. Variation is expressed as se. Significant differences from the wild type are based on Student’s t test: #, P ≥ 0.07; *, P ≤ 0.05; **, P ≤ 0.01; and ***, P ≤ 0.001.

Enhanced phloem loading mediated by AVP1 expression in CCs should lead to greater phloem transport. To test for this, phloem exudation rates from cut stems of transgenic and wild-type control plants were determined. Rosettes were first photosynthetically labeled in the middle of the illuminated period with ¹⁴CO₂, and then the stems below the rosettes were cut and maintained in an EDTA solution, such that less than 2 mm of the cut stem was submerged. The first 20 min of exudation, which would contain the contents of cut cells, was discarded, and ¹⁴C exudation over the next two 1-h intervals was measured and expressed as counts per minute exuded per hour per milligram of fresh weight. To minimize EDTA entering the rosettes via the xylem and transpiration, exudations were conducted in darkened, sealed chambers to promote stomatal closure and maximize humidity. AVP1-overexpressing lines had phloem exudation rates 2- to 5-fold higher than the wild type (Fig. 4B), with the greatest enhancement in the CoYMV_pro:AVP1 lines, indicating more efficient loading and transport of carbon in the phloem. In a related experiment, the rate of unlabeled Suc exuded from cut petioles of AVP1-1 was established relative to the wild type. Throughout the diurnal cycle, transgenic line AVP1-1 had enhanced Suc transport (Fig. 4C). The results presented in Figure 4, A and B, show that plants overexpressing AVP1 from either a constitutive promoter or a phloem-specific promoter load and transport more carbon. Figure 4C argues that these enhancements occur throughout the diurnal cycle and are not directly dependent on photosynthesis. This is consistent with results showing that AVP1-overexpressing plants have more starch throughout the diurnal cycle and, consequently, have more carbon available for transport at night (Pizzio et al., 2015). Diurnal transport in CoYMV_pro:AVP1 lines was not tested, but the same effect is expected.

In the exudation experiments described above, phloem transport is not sink limited, since the stem is cut and the heterotrophic roots are removed. Thus, the experiment addresses Suc that can enter the phloem transport stream but does not speak to whether more photoassimilate is unloaded into sink organs. To assess the efficiency of phloem transport out of leaves and into sink organs, source leaves were photosynthetically labeled for 20 min with ¹⁴CO₂ and ¹⁴C transport into roots was measured by scintillation counting. Wild-type plants transported 10% to 15% of the photoassimilated label out of shoots and into roots, 35S_pro:AVP1 plants transported more than 15% of the assimilated ¹⁴C to roots, and CoYMV_pro:AVP1 plants transported 25% to 35% of the incorporated label to roots (Fig. 5). This shows that long-distance transport and unloading in the root sink tissues is enhanced by constitutive and especially CC-specific AVP1 expression. Furthermore, regions of cell division and elongation constitute the strongest sinks; thus, the roots were cut into 1-cm sections, starting at the root tip, to establish where the majority of the photoassimilate was partitioning. The AVP1-overexpressing lines partitioned a greater proportion of the label to the growing tip of the primary root (lowest 1 cm in Fig. 5) and to the uppermost part of the root (remainder in Fig. 5), which contained emerging lateral roots, indicating that AVP1 overexpression enhances long-distance transport and unloading.

Figure 5.

Constitutive and phloem-specific AVP1 overexpression enhances phloem transport and unloading in heterotrophic roots. A, Ratio of cpm in shoots to total incorporated cpm of wild-type (WT) and 35S_pro:AVP1 lines. B, Ratio of cpm in shoots to total incorporated cpm of wild-type and CoYMV_pro:AVP1 lines. C, Ratio of cpm in roots to total incorporated cpm of wild-type and 35S_pro:AVP1 lines. D, Ratio of cpm in roots to total incorporated cpm of wild-type and CoYMV_pro:AVP1 lines. In C and D, roots were further subdivided into three 1-cm sections from the root tip and the remainder of the root, as indicated in the key. The remainder section has emerging lateral roots. Averages and se are represented; n = 12, and each replicate consisted of a pool of two plants. Shoots of 35S_pro:AVP1 and CoYMV_pro:AVP1 lines have a lower ratio of total label incorporated in the shoots compared with the wild type, whereas the terminal 1-cm distal tip mainly has a significantly higher ratio of label compared with the wild type, indicating increased unloading in the growing region.

Since the CoYMV_pro lines were equivalent or superior to the 35S_pro lines in these loading and transport studies, we conclude that AVP1 overexpression primarily affects plant growth by improving phloem transport. AtSUC2 transcript levels in transgenic lines were not significantly different from the wild-type levels (Supplemental Fig. S3). AVP1-mediated increases in phloem pmf for the H⁺-Suc symport activity of AtSUC2 would account for enhanced loading and transport without a corresponding increase in AtSUC2 levels.

Metabolite and Transcript Analysis in Shoots and Roots of Hydroponically Grown Plants

Enhanced transport from source to sink organs among the transgenic lines relative to the wild type suggests that the heterotrophic roots of the various lines may have different levels of primary metabolites (Fig. 6). Wild-type and transgenic lines were thus grown hydroponically in one-quarter-strength Hoagland solution, such that both shoots and roots were readily accessible. Consistent with the increased rosette growth observed among transgenic lines on potting mix, both shoot and root biomass of the transgenics significantly exceeded that of the wild type when grown hydroponically (Fig. 6, A and B). In addition, the general pattern of biomass accumulation among the various lines was consistent between growth on potting mix and in hydroponics (compare Fig. 6A and Supplemental Fig. S1B), indicating that growing medium did not differentially impact plant growth. Transcript abundance from the genomic copy and transgene copies of AVP1 was measured in shoots and roots of hydroponically grown plants and was consistent with that measured in plants grown on potting mix (data not shown).

Roots and shoots from hydroponically grown plants were independently harvested and analyzed for levels of Glc, Fru, and Suc (Fig. 6, C and D), malate and fumarate (Fig. 6, E and F), Glc-6-P (Fig. 6, G and H), and total amino acid and total protein (Fig. 6, I–L). Starch was measured in rosettes but was below detection limits in roots (Fig. 6M). Similar to the results obtained with rosettes from potting mix-grown plants, a clear pattern of statistically significant differences in metabolite levels did not emerge among the wild type and the lines overexpressing AVP1 either constitutively or specifically in the phloem. Considering the evidence for more reduced carbon reaching the roots, this metabolite analysis argues that steady-state metabolite levels are maintained among all the lines but that the AVP1-overexpressing lines may have more carbon flux through central metabolism.

RT-qPCR was used to measure the transcript abundance of genes that may be influenced by constitutive or CC-specific AVP1 overexpression in shoots and roots of representative hydroponically grown plants (Supplemental Figs. S4 and S5). These included genes encoding SUSs (SUS1, AT5G20830; SUS2, AT5G49190; SUS3, AT4G02280; SUS4, AT3G43190; SUS5, AT5G37180; and SUS6, AT1G73370), UDP-Glc pyrophosphatases (UGP1, AT3G03250; and UGP2, AT5G17310), PM H⁺-ATPases (AHA1, AT2G18960; AHA2, AT4G30190; AHA3, AT5G57350; and AHA4, AT3G47950), cytosolic alkaline invertases (CIN1, At1g35580; and CIN2, AT4G09510); phosphofructokinase α-subunits (PFKα1, AT1G20950; and PFKα2, AT1G20950), phosphofructokinase β-subunits (PFKβ1, AT1G12000; and AtPFKβ2, AT4G04040), and SWEETs involved in phloem loading (SWEET11, AT3G48740; and SWEET12, AT5G23660). Strong changes in gene expression were not observed. AtSUS1 was moderately repressed, while AtSUS2 was moderately enhanced, in the shoot of the representative CoYMV_pro:AVP1 line. AtUGP2 transcript was moderately reduced in the roots of 35S_pro:AVP1 lines.

DISCUSSION

Enhancing phloem loading and long-distance partitioning is a potential means to improve plant productivity through two principal mechanisms: (1) decreasing feedback inhibition on photosynthesis in source leaves, and (2) mobilizing more resources to sink organs for growth, storage, general metabolism, and nutrient uptake (Ainsworth and Bush, 2011; Braun et al., 2014; Yadav et al., 2015). Overexpressing the H⁺-PPase AVP1 and orthologs in numerous plant species increased biomass transfer to sink organs and overall whole-plant biomass accumulation (Li et al., 2005; Lv et al., 2008; Gonzalez et al., 2010). It also increased salt tolerance and drought resistance (Gaxiola et al., 2001; Gao et al., 2006; Brini et al., 2007; Bao et al., 2008; Li et al., 2008, 2010; Lv et al., 2009; Pasapula et al., 2011; Schilling et al., 2014) and improved growth in nutrient-limited conditions (Yang et al., 2007; Pei et al., 2012; Paez-Valencia et al., 2013; Li et al., 2014). Consistent with improved nutrient use and uptake, plants overexpressing AVP1 acidified the rhizosphere and showed greater expression of transporters involved in nitrogen, potassium, and phosphate uptake than wild-type controls (Yang et al., 2007; Undurraga et al., 2012; Pizzio et al., 2015). Enhanced growth and energization of the rhizosphere are outcomes expected for plants with increased phloem loading providing more Suc to sink organs.

A model was developed previously that proposes that H⁺-PPases contribute to phloem loading by functioning as PPi synthase at the CC PM to promote the oxidation of a portion of the Suc loaded into the CC (Fig. 7; Gaxiola et al., 2012). Additional PPi functioning as substrate instead of product favors Suc oxidation at steps mediated by UDP-Glc pyrophosphorylase and PPi-dependent phosphofructokinase (Fig. 7B). This oxidation results in additional ATP, which is hydrolyzed by P-type PM H⁺-ATPases to create more pmf to further energize the phloem system. AVP1 is strongly expressed in source-leaf phloem (Pizzio et al., 2015), and PM localization of H⁺-PPases in the SE-CC complex is well documented (Long et al., 1995; Langhans et al., 2001; Paez-Valencia et al., 2011); the ability of H⁺-PPases to function as PPi synthases energized by pmf is also well documented (Rocha Facanha and de Meis, 1998; Seufferheld et al., 2004).

Figure 7.

A model for different membrane localizations and functions for AVP1 in mesophyll cells and phloem companion cells. A, AVP1 H⁺-PPase localized to the tonoplast of mesophyll cells promotes the hydrolysis of PPi for enhanced Suc synthesis and to energize the vacuole by pumping protons into the lumen. B, Suc/H⁺ symporters on the PM of CCs load Suc from the apoplasm; most is destined for long-distance transport through sieve elements, but a portion is oxidized to energize the phloem network (top of schematic). AVP1 H⁺-PPase localized to the PM of CCs utilizes the pmf to synthesize PPi. More PPi substrate promotes Suc oxidation to create more ATP (middle of schematic). More ATP is hydrolyzed by P-type ATPases on CC PMs to create more pmf (bottom of schematic). Thus, increased Suc oxidization promoted by AVP1-mediated PPi synthesis results in an overall more energized phloem network.

In this study, we used constitutive and CC-specific AVP1 overexpression to directly test for improved phloem loading and long-distance partitioning. Both constitutive overexpression with 35S_pro:AVP1 and CC-specific overexpression with CoYMV_pro:AVP1 resulted in plants with larger rosettes and root systems (Figs. 1 and 6). Both constructs also resulted in improved photosynthesis (Fig. 3), consistent with expectations for enhanced phloem transport. A series of complementary experiments with ¹⁴C labeling directly demonstrated improved phloem loading and long-distance transport to sink organs (Figs. 4 and 5). Loading of [¹⁴C]Suc into the veins of mature source leaves was greater than that in wild-type controls. After photosynthetic labeling with ¹⁴CO₂, up to five times more ¹⁴C label was transported through the phloem and out of cut stems. In intact plants, significantly more ¹⁴C label was transported out of shoots and into roots systems, with CoYMV_pro:AVP1 plants consistently showing the greatest transport levels. Measuring biochemical and transport processes directly in the phloem CCs of source and sink tissues in living plants is notoriously difficult (Turgeon and Wolf, 2009). However, by using CoYMV_pro to overexpress AVP1, we have genetically isolated the biochemical and physiological impacts to CCs. Since CoYMV_pro:AVP1 plants perform equivalently, and by some metrics better than 35S_pro:AVP1 plants (Fig. 5), we conclude that most, if not all, of the beneficial impact of AVP1 overexpression arises from its activity in phloem.

In an effort to identify how AVP1 overexpression influences primary metabolism, steady-state levels of numerous prominent compounds of central metabolism were measured. With constitutive AVP1 overexpression, line AVP1-1, which was used here as a previously characterized control, showed increases in midday rosette starch of plants grown on potting mix, consistent with earlier results (Pizzio et al., 2015), but the other 35S_pro:AVP1 lines did not (Fig. 2). The lines with AVP1 overexpression specifically in CCs had elevated Glc and starch in plants grown on potting mix. These results suggest that, in addition to loading and transporting more carbon (Figs. 4 and 5), the higher photosynthesis in these lines (Fig. 3) results in greater storage reserves, with independent transgenic lines fluctuating moderately in this capacity.

It is not reasonable to compare metabolite levels between rosettes from plants grown in potting mix and those grown hydroponically because of the different conditions, but hydroponic growth allows ready access to the heterotrophic roots and permits comparisons between source and sink organs. Rosettes of the 35S_pro lines had more hexose, particularly Glc (Fig. 6C). Gonzalez et al. (2010) also noted higher levels of hexose at midday in AVP1-1 plants grown in vitro with one-half-strength Murashige and Skoog medium. The other rosette metabolites we measured did not differ consistently and significantly from the wild type when grown hydroponically (Fig. 6), and this too is consistent with prior results (Gonzalez et al., 2010). In roots, AVP1-1 showed increases from the wild type in Glc, Fru, Suc, and, to a lesser extent, malate, indicating a general trend to having more reduced carbon in roots. Line 35S-9 showed slightly lower amino acid levels in rosettes and slightly higher levels in roots, suggesting improved amino acid transport, but this was not consistently observed in the other transgenic lines.

Although prominent changes in steady-state levels of measured metabolites in most lines were not observed, the higher levels of carbon fixation, transport, and growth make it clear that carbon flux through the central metabolism is increased in both source and sink organs. Flux analysis with isotopic ¹³C or ¹⁴C labeling will be necessary to better determine the impact of AVP1 overexpression on Suc oxidation and/or synthesis (i.e. PPi as a substrate during Suc oxidation and as a product during Suc synthesis; Fig. 7; Heise et al., 2014; Ma et al., 2014).

There is substantial interest in metabolic engineering of source/sink relationships to understand and increase plant productivity (Yadav et al., 2015). Overexpressing Suc-H⁺ symporters specifically in CCs resulted in more loading and transport but, unexpectedly, resulted in stunted growth rather than more robust growth. This effect was linked to a potential disruption in homeostasis between transported carbon and nutrient availability in sink organs, particularly phosphate (Dasgupta et al., 2014). In the work presented here, AVP1 overexpression constitutively and specifically in CCs similarly increased phloem loading and carbon transport (Fig. 4), but unlike SUT overexpression, the desired effect of enhanced biomass was achieved. The reason why one method to enhance phloem transport resulted in stunted growth and the other resulted in enhanced growth may relate to the nutrient balances required in the growing sink organs. When phloem loading and transport were increased by SUT overexpression, Suc was presumably the only metabolite for which transport was increased. The proposal that this caused a nutrient imbalance in sinks is supported by SUT overexpression lines showing increased expression of phosphate transporters, purple acid phosphatases, and increased acidification of the rhizosphere, all of which indicate that the plants are perceiving a phosphate limitation. In addition, the stunted phenotype of the SUT overexpression lines was alleviated by supplementing the growth medium with additional phosphate (Dasgupta et al., 2014).

With AVP1 overexpression in the phloem, increased Suc loading and transport were achieved indirectly by promoting Suc oxidation in CCs, leading ultimately to increased ATP production and stronger pmf via PM P-type H⁺-ATPases. AVP1-mediated stimulation of PM ATPase activity is documented (Yang et al., 2007; Undurraga et al., 2012). Interestingly, although Suc transport was enhanced, increased expression of AtSUC2, the Suc transporter responsible for phloem loading in Arabidopsis, was not observed in an earlier study (Gonzalez et al., 2010) or in our own analysis. Similarly, the expression of AtSWEET11 and AtSWEET12, which are involved in Suc release to the apoplasm, did not increase, nor did transcript levels of several steps in Suc metabolism (Supplemental Figs. S3–S5). This implies that enhanced Suc transport resulted from more flux through the pathway and stronger pmf rather than more metabolic enzymes and membrane-bound transporters. In addition to increasing Suc loading, this stronger pmf around the phloem CCs likely also increased the phloem-mediated transport of other nutrients to maintain balance in the phloem sap. Supporting this, and in contrast to the situation with AtSUC2, transcriptome analysis of AVP1-1 rosettes showed more than 2-fold up-regulation of several amino acid transporter genes (Gonzalez et al., 2010), and our analysis showed up-regulation of transporters for nitrate, potassium, and phosphate (Paez-Valencia et al., 2013; Pizzio et al., 2015). Enhanced transport of a balanced nutrient supply to sink organs may account for the larger and more energized root systems with higher water and nutrient uptake efficiencies observed among AVP-overexpressing plants (Gaxiola et al., 2012). In turn, these enhanced root systems likely account for the overall more robust phenotypes and enhanced productivity of plants overexpressing AVP1.

CONCLUSION

A model put forward previously (Gaxiola et al., 2012) proposes that H⁺-PPase AVP1, and its orthologs, has opposite functions depending on cell type and intracellular localization (Fig. 7). In nonphloem cells, the model proposes localization on endomembranes and canonical function as proton-pumping pyrophosphatase to energize the endomembrane compartments (principally, but not exclusively, the tonoplast; Fig. 7A). In phloem CCs, however, localization to the PM is well documented. Here, the H⁺-PPase runs backward as a synthase to synthesize PPi at the expense of the pmf. Increased levels of PPi in CCs promote ATP synthesis by Suc oxidation, which in turn is used by the PM P-type ATPase to create more pmf around the phloem, resulting ultimately in more phloem loading and transport of Suc and presumably other nutrients loaded into the phloem by symport with protons (Fig. 7B). Here, we show directly by physiological and ¹⁴C-labeling studies that overexpression of AVP1 results in enhanced phloem loading and transport of Suc to sink organs. We show that a CC-specific promoter, CoYMV_pro, for overexpression has the same impact as the constitutive promoter, 35S_pro. This indicates that most, if not all, growth enhancements and increases in tolerance to water and nutrient deficiencies observed in both model and crop plant systems may be attributable to a previously unrecognized function for H⁺-PPases in controlling phloem loading and transport. By comparing these results with other efforts to manipulate phloem transport, our findings indicate that a more holistic approach of increasing the pmf around the phloem may be more effective than efforts to engineer transport systems for individual metabolites. The simultaneous overexpression of AVP1 and genes encoding transporters for specific nutrients may be additive.

MATERIALS AND METHODS

Plasmid Construction and Plant Transformation

AVP1 cDNA was amplified by PCR from pRT103-AVP1 (Gaxiola et al., 2001) using forward oligonucleotide AVP1-F (5′-CACCATGGTGGCGCCTGCTTTGTTAC-3′) and reverse oligonucleotide AVP1-R (5′-GAAGTACTTGAAAAGGATACCACC-3′). Phusion Hot start polymerase (New England Biolabs) was used according to the manufacturer’s instructions. The PCR product was cloned with a pENTR/D-TOPO Cloning Kit (Invitrogen), resulting in pENTR:AVP1. pENTR:AVP1 was recombined with pMDC32 (Curtis and Grossniklaus, 2003) using the Gateway LR Clonase II enzyme mix (Invitrogen) to generate pMDC32:AVP1. pMDC32 contains a dual 35S enhancer. pENTR:AVP1 was recombined with pGPTV:CoYMV_pro:cmr:ccdB (Dasgupta et al., 2014) to get pGPTV:CoYMV_pro:AVP1. These vectors were electroporated into Agrobacterium tumefaciens strain GV3101mp90.

Wild-type Columbia-0 (Arabidopsis Biological Resource Center stock no. CS-70000) was transformed with the above constructs by the floral dip method (Clough and Bent, 1998). T1 generation seedlings with pMDC32:AVP1 were selected on sterile one-half-strength Murashige and Skoog medium (Phytotechnology Laboratories) containing 40 mg L⁻¹ hygromycin. Seeds were germinated in the dark and grown for 5 d; resistant seedlings were selected based on their longer hypocotyls (Harrison et al., 2006) and transferred to Fafard 3B potting mix (Sun Gro Horticulture). For selecting plants transformed with pGPTV:CoYMV_pro:AVP1, T1 seeds were germinated on Fafard 3B potting mix and sprayed with 20 mg L⁻¹ glufosinate ammonia (Finale; Farnam) post germination. Twenty-five or more independent lines were identified for both CoYMV_pro:AVP1 and 35S_pro:AVP1. Lack of segregation of the resistance marker in the T3 generation was used to identify homozygous lines. From these, three representative lines for each construct were selected for further experiments based on cDNA expression and growth morphology.

Plant Material and Growth Conditions

For all plant growth, Arabidopsis (Arabidopsis thaliana) seeds were stratified for 72 h at 4°C, and plants were grown in a controlled environment chamber (Percival AR95L; Percival Scientific) with 12 h of 180 µmol photons m⁻² s⁻¹ light at 22°C and 12 h of dark at 18°C. All experiments were conducted with T3 or T4 generation seeds. For the determination of fresh weight, metabolite analysis, and analysis of transcript abundance, seeds were sown on 9-cm square pots, and seedlings were thinned to four well-spaced plants per pot. At 28 d post germination, plants were harvested randomly to create six pools with four plants in each pool. Each pool was frozen in liquid nitrogen and cryogenic ground in 15-mL polycarbonate vials (catalog no. PCRV 15-100-23; OPS Diagnostics) with 8-mm (5/16th inch) stainless steel balls (approximately 2 g each) in a GenoGrinder 2010 (SPEX Sample Prep). The vials, steel balls, and aluminum cryo block for supporting the vials in the GenoGrinder were all prechilled in liquid nitrogen; samples were maintained in liquid nitrogen during processing and otherwise stored at −80°C. For growth rate analysis (Fig. 1C) and photosynthesis measurements, seeds were sown in 65-mm round pots, and seedlings were thinned to one plant per pot. Plants were photographed 16, 20, 22, and 24 d after germination. Rosette area was measured with ImageJ version 1.42f (Rasband, 2014). Growth on vertically oriented square culture dishes for photosynthetic labeling with ¹⁴CO₂ was described previously (Dasgupta et al., 2014). Growth on potting mixture for photosynthetic labeling with ¹⁴CO₂ was described previously (Dasgupta et al., 2014). Plants were grown hydroponically as described (Guo et al., 2008) except that one-quarter-strength Hoagland solution (Phytotechnology Laboratories) was used and replaced every 7 d. Medium trays and black expanded polyvinylcarbonate sheets to cover the trays and support the shoots were obtained from US Plastics (catalog nos. 49275 and 42500, respectively). Holes were drilled in the polyvinylcarbonate sheets for 12 plants to be grown in each tray. For each line, six pools of four plants, randomized from different trays, were harvested 22 d after germination: shoots, cut at the hypocotyl, were weighed and frozen directly, and the roots were quickly blotted dry by gently pressing between paper towels prior to weighing and freezing. The frozen samples were cryo-ground and stored as described above. All tissues were harvested 5 to 6 h into the light period.

AVP1 Immunolocalization

Arabidopsis source leaves from plants grown in potting mix were fixed in 3.7% (v/v) formaldehyde, 50% (v/v) ethanol, and 5% (v/v) acetic acid overnight at 4°C. Fixed material was dehydrated in a graded ethanol series (50%, 60%, 70%, 80%, 90%, and 100%, all v/v), and absolute ethanol was replaced by histological clearing agent (xylene; Sigma-Aldrich). Tissues were embedded in Paraplast at 60°C. The embedded tissue was sliced into 10- to 15-μm sections and placed onto poly-Lys-coated slides. Sections were deparaffinized in xylene, hydrated in graded alcohol (100%, 90%, 80%, 70%, 60%, 50%, and 30%, with deionized water), and pretreated with Antigen Retrieval Solution (BioGenex) under a steamer at 65°C for 20 min. Endogenous peroxidase activity was quenched with 3% hydrogen peroxide, and the slides were treated with Power Universal Reagent (BioGenex) to reduce nonspecific binding and incubated with polyclonal anti H⁺-PPase antibody (Park et al., 2005). Controls were done with preimmune serum at a 1:1,000 dilution in phosphate-buffered saline for all samples. Then, three rinses with phosphate-buffered saline-Tween (0.1%, v/v) were followed by incubation with peroxidase-labeled polymer conjugated to goat anti-rabbit immunoglobulins as provided by the EnVision system (DAKO), following the manufacturer’s instructions.

Transcript Analysis

Total RNA was isolated from 50 mg of cryo-ground rosette tissues (see above) using Trizol (Life Technologies). Total RNA was further treated with Turbo RNase-free DNase-I (Life Technologies) or with the RTS DNase Kit (Mo Bio Laboratories). One milligram of total RNA was reverse transcribed with 50 µm oligo(dT) (Life Technologies) and either SuperScript III reverse transcriptase (Life Technologies) or GoScript Reverse Transcriptase (Promega) according to the manufacturer’s instructions. The gene-specific primers used for qPCR are described in Supplemental Table S1. Real-time qPCR was carried out on an Applied Biosystems ViiA 7 device (Life Technologies) with GO Taq qPCR Mix (Promega) using the following protocol: 2 min of denaturation at 95°C followed by 40 cycles of 95°C for 10 s, 65°C for 30 s, and 70°C for 30 s; or with JumpStart Taq Ready Mix (Sigma-Aldrich) using the following thermal profile: 2 min of denaturation at 94°C followed by 40 cycles of 94°C for 20 s, 60°C for 30 s, and 65°C for 60 s. A final melting curve confirmed a single PCR product. Three biological and two technical replicates were used for analysis, and transcript levels were normalized against GAPDH3 transcript abundance.

Extraction and Measurement of Metabolites

Aliquots of 20 mg of cryo-ground tissue (see above) were extracted with 80% ethanol or TCA (Gibon et al., 2002). The major transient soluble carbohydrates and starch were extracted with ethanol and quantified enzymatically (Stitt et al., 1989) using a Synergy H Reader (Bio Tek) or by high-performance anion-exchange chromatography with pulsed amperometric detection (Dasgupta et al., 2014). Starch was measured enzymatically from the insoluble material after ethanolic extraction of soluble carbohydrates (Hendriks et al., 2003) or by a commercially available starch assay kit (Megazyme). Total protein was measured using the Bradford assay (Bradford, 1976). Glc-6-P was determined according to Gibon et al. (2004). Malate and fumarate were determined according to Cross et al. (2006). Inorganic phosphate and total phosphate were measured using an ammonium molybdate assay (Chiou et al., 2006). Total amino acids were measured using a fluorescamine method (Gibon et al., 2009).

Photosynthesis Measurement

Photosynthesis per unit of rosette surface area was measured with a Li-Cor Li-6400XT infrared gas analyzer equipped with a whole-plant Arabidopsis chamber and red, green, and blue light source according to the equipment manufacturer’s instructions. Plants were grown for 21 d in 65-mm-diameter round pots that fit in the whole-plant Arabidopsis chamber. The surface of potting mix was sealed with pottery clay to avoid gas exchange with the potting medium. Gas-exchange measurements were performed with 400 µmol s⁻¹ CO₂, a gas flow rate of 500 mmol s⁻¹, and photosynthetically active radiation at 225 µmol m⁻² s⁻¹ to reasonably mimic environmental conditions used during growth. Chamber block temperature and relative humidity were established by ambient air conditions and were 23°C and approximately 60% respectively. Plants were allowed to acclimate for 120 s in the chamber before taking measurements. The acclimatization time was determined empirically. Two measurements were taken per plant in 30-s intervals. Plants were digitally photographed, and rosette area was determined with ImageJ software (Rasband, 2014) to standardize photosynthetic rates per unit area. Saturating light levels were not used for infrared gas analysis, since this would not have represented the physiological state during growth.

¹⁴C Labeling to Measure Phloem Loading and Transport

Phloem loading of [¹⁴C]Suc into source leaf discs was conducted as described (Dasgupta et al., 2014), except that 5-mm leaf discs were excised from source leaves rather than using whole leaves. Six replicates, each consisting of six leaf discs randomized from six plants, were used for each control and experimental line. Photosynthetic labeling of plants grown on potting mix for phloem exudation and the EDTA exudation method was described previously (Dasgupta et al., 2014). Photosynthetic labeling of seedlings grown in vertically oriented square culture dishes was described previously (Cao et al., 2013; Dasgupta et al., 2014). Shoots and roots were collected separately, and roots were cut at 1-cm intervals from the tip of the root. ¹⁴C levels (cpm) in shoots and root sections were determined by scintillation counting as described (Cao et al., 2013; Dasgupta et al., 2014). Two plants were pooled for analysis. Label in shoots was standardized relative to shoot mass, and label in roots was standardized relative to segment length. All experiments in this study were conducted with 12 h of light and 12 h of dark rather than 14 h of light and 10 h of dark.

To measure unlabeled Suc exuded from cut petioles at different times during the diurnal cycle (Fig. 4C), 14 to 16 mature rosette source leaves were harvested from 40-d-old adult plants for each time point. Phloem exudation was done as reported by Tetyuk et al. (2013). In brief, leaves were cut and immediately placed in 20 mm K₂-EDTA. Then, all leaves were stacked on top of each other, petioles were recut under 20 mm K₂-EDTA, and cut ends were placed in reaction tubes with 1 mL of fresh 20 mm K₂-EDTA solution for 1 h at 21°C. Suc content in exudates was determined by using the Suc/d-Glc/d-Fru assay kit (Megazyme).

Sequence data for the genes studied in this article are listed in Supplemental Table 1.

Supplemental Data

The following supplemental materials are available.

• Supplemental Figure S1. Growth characteristics of AVP1 lines.

• Supplemental Figure S2. Autoradiograms of representative leaf disks.

• Supplemental Figure S3. AtSUC2 expression in wild-type and AVP1 lines.

• Supplemental Figure S4. Expression of genes relating to carbon flux through central metabolism in shoots.

• Supplemental Figure S5. Expression of genes relating to carbon flux through central metabolism in roots.

• Supplemental Table S1. List of gene sequences and primers used in RT-qPCR analysis.

Glossary

CC

companion cell

SE

sieve element

pmf

proton motive force

PM

plasma membrane

cDNA

complementary DNA

qPCR

quantitative PCR

RT

reverse transcription

PPi

pyrophosphate