Grain Unloading of Arsenic Species in Rice

Rice (Oryza sativa) is the staple food for over half the world''s population yet may represent a significant dietary source of inorganic arsenic (As), a nonthreshold, class 1 human carcinogen. Rice grain As is dominated by the inorganic species, and the organic species dimethylarsinic acid (DMA). To investigate how As species are unloaded into grain rice, panicles were excised during grain filling and hydroponically pulsed with arsenite, arsenate, glutathione-complexed As, or DMA. Total As concentrations in flag leaf, grain, and husk, were quantified by inductively coupled plasma mass spectroscopy and As speciation in the fresh grain was determined by x-ray absorption near-edge spectroscopy. The roles of phloem and xylem transport were investigated by applying a ± stem-girdling treatment to a second set of panicles, limiting phloem transport to the grain in panicles pulsed with arsenite or DMA. The results demonstrate that DMA is translocated to the rice grain with over an order magnitude greater efficiency than inorganic species and is more mobile than arsenite in both the phloem and the xylem. Phloem transport accounted for 90% of arsenite, and 55% of DMA, transport to the grain. Synchrotron x-ray fluorescence mapping and fluorescence microtomography revealed marked differences in the pattern of As unloading into the grain between DMA and arsenite-challenged grain. Arsenite was retained in the ovular vascular trace and DMA dispersed throughout the external grain parts and into the endosperm. This study also demonstrates that DMA speciation is altered in planta, potentially through complexation with thiols.Paddy rice (Oryza sativa) is particularly effective, compared to other cereals, at accumulating arsenic (As) in shoot and grain (Williams et al., 2007b). Rice is the staple food for over half the world''s population (Fageria, 2007) and rice represents a significant dietary source of inorganic As, a class 1, nonthreshold carcinogen, particularly in Southeast Asia (Meharg et al., 2009). Inorganic As levels in rice grain are problematic even where soil As is at background levels, derived from geogenic sources (Lu et al., 2009; Meharg et al., 2009). However, widespread pollution of paddy soils with As, leading to further elevation of grain As, has occurred in some regions due to base and precious mining (Liao et al., 2005; Zhu et al., 2008), irrigation of paddies with As-elevated groundwaters (e.g. Meharg and Rahman, 2003; Williams et al., 2006), and the use of arsenical pesticides (Williams et al., 2007a). Unlike other cereal grains, paddy rice cultivation is dependent of soils being anaerobic, and it is this anoxia that gives rise to elevated As concentrations in the plant. Anaerobic soil conditions lead to the mobilization of As as arsenite, where under aerobic systems arsenate dominates (Xu et al., 2008). Arsenite is efficiently assimilated by rice roots through silicic acid transport pathway (Ma et al., 2008).Knowledge of As metabolism and partitioning within plants, particularly rice, is still developing rapidly (Zhao et al., 2009). Several studies have now shown that As in rice vegetative tissue and grain is predominantly speciated as inorganic As and the methylated species dimethylarsinic acid (DMA), with variable, though low, levels of monomethyl arsonic acid (MMA; Abedin et al., 2002a; Williams et al., 2005, 2006; Norton et al., 2009). Arsenate is an analog of phosphate and competes with phosphate for rice root uptake (Abedin et al., 2002a) while arsenite is taken up by rice roots via silicic acid transporters (Ma et al., 2008). Abedin et al. (2002b) demonstrated that the methylated species DMA and MMA are also taken up by rice plants although at a much slower rate than inorganic As, with the protonated neutral forms also transported through silicic acid pathway (Li et al., 2009). Arsenate is reduced to arsenite within the rice root (Xu et al., 2008; Zhao et al., 2009), which then enters the xylem via a silicic acid/arsenite effluxer (Ma et al., 2008; Zhao et al., 2009). Arsenite may be detoxified through complexation with thiol-rich peptides including phytochelatins (PCs) and glutathione followed by sequestration into vacuoles (Bleeker et al., 2006; Raab et al., 2007b; Zhao et al., 2009). Raab et al. (2007a) found that while methylated As species are taken up by rice roots much less efficiently than inorganic species, they appear to be translocated within the plant more efficiently. The comparative contributions of xylem and phloem transport, in translocation of As to the grain, are unknown.The main species within rice grain, along with DMA, are inorganic As, particularly arsenite, which may be complexed with thiols (Williams et al., 2005; Lombi et al., 2009). Nutrients are unloaded into the grain from the ovular vascular trace (OVT) into the nucellar tissue and from there are uploaded, via the apoplast into the filial tissue (the aleurone and the endosperm; Krishnan and Dayanandan, 2003). Lombi et al. (2009) recently suggested that this may represent a physiological barrier that As species cross with differential efficiency. However, the transport and unloading of As to/into the grain, which are key processes in terms of human exposure to this contaminant, are far from being fully understood.This study investigated the differential efficiency with which important As species are translocated and unloaded into the rice grain and the comparative contributions of phloem and xylem transport. Rice panicles were excised below the flag leaf node during grain development, 10 DPA, and treated to a hydroponically administered 48-h pulse of arsenite, arsenate, arsenite glutathione, or DMA. Total As concentrations in flag leaf, grain, and husk samples for each treatment were quantified by inductively coupled plasma mass spectroscopy (ICP-MS), and As speciation in the fresh grain was determined by x-ray absorption near-edge spectroscopy (XANES) analysis. To evaluate the contributions of phloem versus xylem transport, a stem-girdling treatment was applied, using steam to destroy phloem cells in a second set of panicles prior to a pulse of either DMA or arsenite. The spatial unloading of As species into the developing grain was examined by synchrotron x-ray fluorescence (XRF) mapping, and fluorescence microtomography for the DMA and arsenite treatments.     

Arsenic Speciation in Phloem and Xylem Exudates of Castor Bean

How arsenic (As) is transported in phloem remains unknown. To help answer this question, we quantified the chemical species of As in phloem and xylem exudates of castor bean (Ricinus communis) exposed to arsenate [As(V)], arsenite [As(III)], monomethylarsonic acid [MMA(V)], or dimethylarsinic acid. In the As(V)- and As(III)-exposed plants, As(V) was the main species in xylem exudate (55%–83%) whereas As(III) predominated in phloem exudate (70%–94%). The ratio of As concentrations in phloem to xylem exudate varied from 0.7 to 3.9. Analyses of phloem exudate using high-resolution inductively coupled plasma-mass spectrometry and accurate mass electrospray mass spectrometry coupled to high-performance liquid chromatography identified high concentrations of reduced and oxidized glutathione and some oxidized phytochelatin, but no As(III)-thiol complexes. It is thought that As(III)-thiol complexes would not be stable in the alkaline conditions of phloem sap. Small concentrations of oxidized glutathione and oxidized phytochelatin were found in xylem exudate, where there was also no evidence of As(III)-thiol complexes. MMA(V) was partially reduced to MMA(III) in roots, but only MMA(V) was found in xylem and phloem exudate. Despite the smallest uptake among the four As species supplied to plants, dimethylarsinic acid was most efficiently transported in both xylem and phloem, and its phloem concentration was 3.2 times that in xylem. Our results show that free inorganic As, mainly As(III), was transported in the phloem of castor bean exposed to either As(V) or As(III), and that methylated As species were more mobile than inorganic As in the phloem.Arsenic (As) is an environmental and food chain contaminant that has attracted much attention in recent years. Soil contamination with As may lead to phytotoxicity and reduced crop yield (Panaullah et al., 2009). Food crops are also an important source of inorganic As, a class-one carcinogen, in human dietary intake, and there is a need to decrease the exposure to this toxin (European Food Safety Authority, 2009). Paddy rice (Oryza sativa) is particularly efficient in As accumulation, which poses a potential risk to the population based on a rice diet (Meharg et al., 2009; Zhao et al., 2010a). Other terrestrial food crops generally do not accumulate as much As as paddy rice; however, where soils are contaminated, relatively high concentrations of As in wheat (Triticum aestivum) grain have been reported (Williams et al., 2007; Zhao et al., 2010b). On the other hand, some fern species in the Pteridaceae family are able to tolerate and hyperaccumulate As in the aboveground part to >1,000 mg kg⁻¹ dry weight (e.g. Ma et al., 2001; Zhao et al., 2002); these plants offer the possibility for remediation of As-contaminated soil or water (Salido et al., 2003; Huang et al., 2004). A better understanding of As uptake and long-distance transport, metabolism, and detoxification is needed for developing strategies for mitigating As contamination, through either decreased As accumulation in food crops or enhanced As accumulation for phytoremediation.The pathways of As uptake by plant roots differ between different As species; arsenate [As(V)] enters plant cells via phosphate transporters, whereas arsenite [As(III)] is taken up via some aquaporins (for review, see Zhao et al., 2009). In rice, a silicic acid efflux protein also mediates As(III) efflux toward stele for xylem loading (Ma et al., 2008). Methylated As species, such as monomethylarsonic acid [MMA(V)] and dimethylarsinic acid [DMA(V)], which may be present in the environment as products of microbial or algal methylation of inorganic As or from past uses of methylated As pesticides, are taken up by rice roots partly through the aquaporin NIP2;1 (for nodulin 26-like intrinsic protein; also named Lsi1; Li et al., 2009). Once inside plant cells, As(V) is reduced to As(III), possibly catalyzed by As(V) reductase(s) such as the plant homologs of the yeast (Saccharomyces cerevisiae) ACR2 (Bleeker et al., 2006; Dhankher et al., 2006; Ellis et al., 2006; Duan et al., 2007). As(III) has a high affinity to thiol (-SH) groups and is detoxified by complexation with thiol-rich phytochelatins (PCs; Pickering et al., 2000; Schmöger et al., 2000; Raab et al., 2005; Bluemlein et al., 2009; Liu et al., 2010). As(III)-PC complexation in roots was found to result in reduced mobility for efflux and for long-distance transport, possibly because the complexes are stored in the vacuoles (Liu et al., 2010). Excess As(III) causes cellular toxicity by binding to the vicinal thiol groups of enzymes, such as the plastidial lipoamide dehydrogenase, which has been shown to be a sensitive target of As toxicity (Chen et al., 2010). The As hyperaccumulating Pteris species differ from nonhyperaccumulating plants because of enhanced As(V) uptake (Wang et al., 2002; Poynton et al., 2004), little As(III)-thiol complexation (Zhao et al., 2003; Raab et al., 2004), and efficient xylem loading of As(III) (Su et al., 2008). Recently, an As(III) efflux transporter, PvACR3, has been found to play an important role in As(III) detoxification by transporting As(III) into vacuoles in Pteris vittata (Indriolo et al., 2010).With the exception of As hyperaccumulators, most plant species have a limited root-to-shoot translocation of As (Zhao et al., 2009). The chemical species of As in xylem exudate have been determined in a number of plant species. As(III) was found to be the predominant species (80%–100%) in the xylem sap of rice, tomato (Solanum lycopersicum), cucumber (Cucumis sativus), and P. vittata even when these plants were fed As(V) (Mihucz et al., 2005; Xu et al., 2007; Ma et al., 2008; Su et al., 2010), suggesting that As(V) is reduced in roots before being loaded into the xylem. In other plant species, such as Brassica juncea (Pickering et al., 2000), wheat, and barley (Hordeum vulgare; Su et al., 2010), As(V) accounted for larger proportions (40%–50%) of the total As in the xylem sap. Studies using HPLC-inductively coupled plasma (ICP)-mass spectrometry (MS) coupled with electrospray (ES)-MS showed no evidence of As(III)-thiol complexation in the xylem sap of sunflower (Helianthus annuus; Raab et al., 2005). When rice plants were exposed to MMA(V) or DMA(V), both As species were found in the xylem sap (Li et al., 2009). Generally, methylated As species are taken up by roots at slower rates than inorganic As, but they are more mobile during the xylem transport from roots to shoots (Marin et al., 1992; Raab et al., 2007; Li et al., 2009).It has been shown that phloem transport contributes substantially to As accumulation in rice grain (Carey et al., 2010). However, little is known about how As is transported in phloem (Zhao et al., 2009). There are no reports on the chemical species of As in phloem exudate. The speciation of As in phloem is important because it dictates how As is loaded in the source tissues and unloaded in the sink tissues, such as grain. Questions with regard to the oxidation state, methylation, and complexation of As in phloem sap remain to be answered. Unlike xylem sap, phloem sap is much more difficult to obtain in sufficient quantities for analysis. In this study, we investigated As speciation in phloem and xylem exudates of castor bean (Ricinus communis), which is widely used as a model plant to investigate phloem transport of solutes (e.g. Hall et al., 1971; Hall and Baker, 1972; Allen and Smith, 1986; Bromilow et al., 1987).     

Most Water in the Tomato Truss Is Imported through the Xylem,Not the Phloem: A Nuclear Magnetic Resonance Flow Imaging Study

In this study, we demonstrate nuclear magnetic resonance flow imaging of xylem and phloem transport toward a developing tomato (Solanum lycopersicum) truss. During an 8-week period of growth, we measured phloem and xylem fluxes in the truss stalk, aiming to distinguish the contributions of the two transport tissues and draw up a balance between influx and efflux. It is commonly estimated that about 90% of the water reaches the fruit by the phloem and the remaining 10% by the xylem. The xylem is thought to become dysfunctional at an early stage of fruit development. However, our results do not corroborate these findings. On the contrary, we found that xylem transport into the truss remained functional throughout the 8 weeks of growth. During that time, at least 75% of the net influx into the fruit occurred through the external xylem and about 25% via the perimedullary region, which contains both phloem and xylem. About one-half of the net influx was lost due to evaporation. Halfway through truss development, a xylem backflow appeared. As the truss matured, the percentage of xylem water that circulated into the truss and out again increased in comparison with the net uptake, but no net loss of water from the truss was observed. The circulation of xylem water continued even after the fruits and pedicels were removed. This indicates that neither of them was involved in generating or conducting the circulation of sap. Only when the main axis of the peduncle was cut back did the circulation stop.Fruits are terminal organs that depend completely on long-distance transport to supply them with sugars and water for growth. Water is imported by means of both the xylem and the phloem, whereas sugars are only imported by means of the phloem. Fruits have to compete for water with the rest of the plant, and for that reason, xylem influx is expected to be sensitive to changes in plant water potential. Xylem influx into fruits may thus be lower during the day and higher during the night. When in the apoplast the water potential is especially low, for instance, when the plant is transpiring a lot of water during a hot day, fruits may even experience a xylem efflux and lose water to the vegetative parts of the plant (Johnson et al., 1992; Guichard et al., 2005). It has been suggested that in several species, in order to reduce the sensitivity of fruits to changes in plant water status, during fruit development the xylem connection between fruit and plant is reduced or even severed (Findlay et al., 1987; Lang, 1990; Creasy et al., 1993; Lang and Ryan, 1994; van Ieperen et al., 2003; Drazeta et al., 2004). In contrast to the xylem, the phloem is expected to be relatively insensitive to diurnal changes in water potential (Ehret and Ho, 1986; Ho et al., 1987). For instance, in the main stem of a number of plants, the phloem was found not to respond to diurnal differences in plant water status, whereas the xylem did (Peuke et al., 2001; Windt et al., 2006).The tomato (Solanum lycopersicum) plant has been the subject of many studies dealing with long-distance transport to fruits and has been chosen as a model system in this study as well. It has been estimated that in tomato fruits, about 80% to 90% of the influx of sap takes place by means of the phloem (Ho et al., 1987; Plaut et al., 2004; Guichard et al., 2005). It has been proposed that the low xylem contribution is due to the presence of some form of restriction in the xylem connection between plant and fruit, possibly in the knuckle (Lee, 1989; van Ieperen et al., 2003). Despite the expected low xylem contribution and limited conductivity of the xylem connection between plant and fruit, fruits have been shown to exhibit a diurnal pattern of growth. In most cases, fruits have been observed to grow fastest at night (Lee, 1989; Grange, 1995; van de Sanden and Uittien, 1995; Guichard et al., 2005). The opposite has been found to occur as well (Ehret and Ho, 1986; Pearce et al., 1993), but in these cases, the faster daytime growth was probably caused by a low diurnal stress environment. In a number of studies, even an efflux of xylem sap and fruit shrinkage during the day was reported (Johnson et al., 1992; Leonardi et al., 1999, 2000). It has been proposed that, if the phloem and xylem operate under different diurnal cycles or if their relative contributions can be modified in any way by adjusting the environmental conditions in a greenhouse, it might become possible to control and regulate fruit yield as well as fruit quality and taste.Considering the importance of fruit for the world''s food production, surprisingly little is known about the dynamics of sap flow to fruits. Since the conception of the cohesion tension theory (Dixon and Joly, 1894) and the Munch pressure flow hypothesis (Münch, 1930), there has been a decent theoretical understanding of the basic forces that govern phloem and xylem flow. It has already been attempted to apply this understanding to model fruit growth for a variety of fruits and applications (e.g. Daudet et al., 2002). However, many of the parameters that are needed to model long-distance transport to fruits are currently outside of experimental reach. First, little is known about the pressure and water potential gradients that drive flow to fruits. The xylem and the phloem are extremely sensitive to invasive experimentation and are easily disturbed, and the water potentials in the fruits’ symplast and apoplast are difficult to assess. Second, it is not clear whether xylem and phloem sap only enters the fruit (unidirectional flow), or if return flow is possible as well, and if it is, under which conditions it may occur. As the results of this study show, NMR flow imaging can provide answers to these important questions.

Estimating Long-Distance Transport to Fruits

So far, the most important methods to estimate xylem and phloem influx in fruits have been the subtractive method (Lang and Thorpe, 1989) and the mineral accumulation method (Ho et al., 1987). In the subtractive method, the contribution of xylem and phloem are estimated by heat girdling the pedicel (fruit stalk) of a fruit. Heat girdling destroys the sieve tubes, stopping phloem influx, while the xylem is assumed to remain intact and functional. By comparing the growth of nongirdled fruits to that of girdled fruits, the phloem contribution can be estimated. The most critical assumption in this method is that the xylem sap flow is not affected by heat girdling. However, the validity of this assumption is not evident. First, because xylem and phloem flow to fruits are coupled. Xylem influx is driven by a water potential difference between the xylem and the fruit symplast, which is maintained by osmotically active compounds (sugars), which in turn are imported by means of the phloem. Fishman et al. (2001) showed that the coupling between phloem and xylem influx could give rise to significant errors when using the pedicel girdling technique. A second reason is that heat girdling may profoundly affect xylem function. The xylem tissue may apparently escape heat girdling unscathed, as demonstrated by Guichard et al. (2005), but if the surrounding cells are damaged, it is not unlikely that functional damage will occur. For instance, it has been proposed that the cells that surround the xylem protect it against embolisms by preventing the entry of air (Hacke and Sperry, 2001). Van Ieperen et al. (2003) found that in the tomato pedicel, the abscission zone is the site of highest xylem resistance and that only a few xylem conduits traverse it. If an obstruction would occur in these conduits, either by embolisms or by particles of debris, it could significantly affect xylem resistance and have large implications for xylem transport to the fruit.In the second method, the mineral composition of the fruit is used to estimate the relative xylem and phloem contribution. Ho et al. (1987) measured calcium accumulation, net water import, and fruit respiration in tomato fruits. The xylem contribution to fruit growth was then estimated based on a number of assumptions: (1) the calcium content of phloem sap can be neglected compared to that of xylem sap; (2) the calcium content of xylem sap is similar to that measured in root stump exudate; and (3) xylem backflow from fruits does not occur. However, in view of current knowledge, the first and third assumptions are questionable. Calcium is used in signal transduction and as such is known to be present in the phloem. The question is, in what concentration. In phloem sap exudate of castor bean (Ricinus communis) and eucalyptus (Eucalyptus globulus), calcium concentrations have been found that were about 66% and 25% of the concentration in the root stump exudate, respectively (Pate et al., 1998; Peuke et al., 2006). In Banksia prionotes, the calcium concentration in phloem exudate was even found to be 10 times higher than that of the xylem sap (Pate and Jeschke, 1995). It should be noted that in these studies phloem sap was harvested by cutting. This may have elicited a wounding response, causing elevated calcium levels in the phloem (Knoblauch et al., 2001). Still, we argue that these findings illustrate that the calcium concentration in the phloem cannot be assumed to be negligible, especially when the majority of influx of sap is thought to take place via the phloem. The assumption that backflow does not take place also may not hold. In a number of studies, backflow from tomato fruits has already been observed, especially under summer conditions or high vapor deficit (Johnson et al., 1992; Leonardi et al., 1999, 2000; Guichard et al., 2005). The subtractive and the mineral accumulation method thus are likely to be subject to large systematic errors. Better methods to estimate or measure long-distance transport to fruits are needed.

NMR Flow Imaging

Over the last 10 years, it has been demonstrated that NMR flow imaging can provide an excellent tool to measure xylem and phloem transport (Van As, 2007). NMR flow imaging does not only give information about the average flow velocity, such as heat pulse based methods do, but gives access to all properties of the flowing water, such as the flow conducting area, the distribution of flow velocities, and the volume flow, all on a per pixel basis (Scheenen et al., 2000b). So far, studies have been conducted measuring flow in the stem of a variety of plants, ranging from castor bean seedlings (Köckenberger et al., 1997) to fully developed tomato, castor bean, and tobacco (Nicotiana tabacum) plants, and a small poplar tree (Populus spp.; Windt et al., 2006). The technique has been used to study the diurnal variation in long-distance transport (Peuke et al., 2001; Windt et al., 2006), the effects of cold girdling (Peuke et al., 2006), and xylem embolism repair (Scheenen et al., 2007) and has been used as a reference technique to provide detailed velocity maps for comparison with different heat pulse methods (e.g. Helfter et al., 2007; D. Chavarro, C.W. Windt, M.W. Lubczynski, J. Roy, and H. Van As, unpublished data). These studies have in common that flow was only measured in the main stem of the plant. This is a convenient place to do flow imaging for a variety of reasons. In comparison with other flow-conducting structures in the plant, the stem is large, sturdy, and stable. It conducts the largest fluxes, and the xylem and phloem can be easily distinguished on the basis of their direction of flow. These properties make imaging xylem and phloem transport relatively easy.

Aims and Research Questions

In this study, we used NMR flow imaging to measure long-distance transport to fruits. As a model plant, tomato was chosen. The anatomy of the tomato truss, as well as the dimensions of the magnetic resonance imaging (MRI) device and its components, made it impossible to image the pedicel of a single fruit. The pedicels were too short and too close together to fit them with the radio frequency (RF) coil that is needed for MRI. For this reason, we chose to perform flow imaging on the peduncle of tomato, measuring the transport toward the entire developing truss. After fitting the plant in the imager, it was impossible to remove the plant without damaging it. The plant was therefore left in the imager and allowed to grow there for 8 weeks. In this period, we continuously monitored long-distance transport into the truss, aiming to answer the following questions: (1) can xylem and phloem flow into the truss be visualized and distinguished; (2) what transport tissues conduct sap into the truss during truss development; (3) is phloem and xylem transport into the truss unidirectional, or does backflow occur; and (4) can NMR flow imaging be used to draw up a quantitative balance of xylem and phloem influx into the truss?     

Grain setting defect1, Encoding a Remorin Protein,Affects the Grain Setting in Rice through Regulating Plasmodesmatal Conductance

Effective grain filling is one of the key determinants of grain setting in rice (Oryza sativa). Grain setting defect1 (GSD1), which encodes a putative remorin protein, was found to affect grain setting in rice. Investigation of the phenotype of a transfer DNA insertion mutant (gsd1-Dominant) with enhanced GSD1 expression revealed abnormalities including a reduced grain setting rate, accumulation of carbohydrates in leaves, and lower soluble sugar content in the phloem exudates. GSD1 was found to be specifically expressed in the plasma membrane and plasmodesmata (PD) of phloem companion cells. Experimental evidence suggests that the phenotype of the gsd1-Dominant mutant is caused by defects in the grain-filling process as a result of the impaired transport of carbohydrates from the photosynthetic site to the phloem. GSD1 functioned in affecting PD conductance by interacting with rice ACTIN1 in association with the PD callose binding protein1. Together, our results suggest that GSD1 may play a role in regulating photoassimilate translocation through the symplastic pathway to impact grain setting in rice.Grain filling, a key determinant of grain yield in rice (Oryza sativa), hinges on the successful translocation of photoassimilates from the leaves to the fertilized reproductive organs through the phloem transport system. Symplastic phloem loading, which is one of the main pathways responsible for the transport of photoassimilates in rice, is mediated by plasmodesmata (PD) that connect phloem companion cells with sieve elements and surrounding parenchyma cells (Kaneko et al., 1980; Chonan et al., 1981; Eom et al., 2012). PD are transverse cell wall channels structured with the cytoplasmic sleeve and the modified endoplasmic reticulum desmotubule between neighboring cells (Maule, 2008). A number of proteins affect the structure and functional performance of the PD, which in turn impacts the cell-to-cell transport of small and large molecules through the PD during plant growth, development, and defense (Cilia and Jackson, 2004; Sagi et al., 2005; Lucas et al., 2009; Simpson et al., 2009; Stonebloom et al., 2009). For example, actin and myosin, which link the desmotubule to the plasma membrane (PM) at the neck region of PD, are believed to play a role in regulating PD permeability by controlling PD aperture (White et al., 1994; Ding et al., 1996; Reichelt et al., 1999). Callose deposition can also impact the size of the PD aperture at the neck region (Radford et al., 1998; Levy et al., 2007) and callose synthase genes such as Glucan Synthase-Like7 (GSL7, also named CalS7), GSL8, and GSL12 have been shown to play a role in regulating symplastic trafficking (Guseman et al., 2010; Barratt et al., 2011; Vatén et al., 2011; Xie et al., 2011). Other proteins that have been shown to impact the structure and function of the PD include glycosylphosphatidylinositol (GPI)-anchored proteins, PD callose binding protein1 (PDCB1), which is also associated with callose deposition (Simpson et al., 2009), and LYSIN MOTIF DOMAIN-CONTAINING GLYCOSYLPHOSPHATIDYLINOSITOL-ANCHORED PROTEIN2, which limits the molecular flux through the PD by chitin perception (Faulkner et al., 2013). Changes in PD permeability can have major consequences for the translocation of photoassimilates needed for grain filling in rice. However, the genes and molecular mechanisms underlying the symplastic transport of photoassimilates remain poorly characterized.Remorins are a diverse family of plant-specific proteins with conserved C-terminal sequences and highly variable N-terminal sequences. Remorins can be classified into six distinct phylogenetic groups (Raffaele et al., 2007). The functions of most remorins are unknown, but some members of the family have been shown to be involved in immune response through controlling the cell-to-cell spread of microbes. StREM1.3, a remorin that is located in PM rafts and the PD, was shown to impair the cell-to-cell movement of a plant virus X by binding to Triple Gene Block protein1 (Raffaele et al., 2009). Medicago truncatula symbiotic remorin1 (MtSYMREM1), a remorin located at the PM in Medicago truncatula, was shown to facilitate infection and the release of rhizobial bacteria into the host cytoplasm (Lefebvre et al., 2010). Overexpression of LjSYMREM1, the ortholog of MtSYMREM1 in Lotus japonicus, resulted in increased root nodulation (Lefebvre et al., 2010; Tóth et al., 2012). Although a potential association between remorins and PD permeability has been proposed (Raffaele et al., 2009), the diversity observed across remorins, plus the fact that remorin mutants generated through different approaches fail to show obvious phenotypes (Reymond et al., 1996; Bariola et al., 2004), have made it challenging to characterize the function of remorins in cell-to-cell transport.In this study, we identified a rice transfer DNA (T-DNA) insertion mutant (grain setting defect1-Dominant [gsd1-D]), with a grain setting-deficient phenotype caused by overexpression of GSD1, a remorin gene with unknown function. GSD1 is expressed specifically in phloem companion cells and is localized in the PD and PM. We provide evidence to show that overexpression of GSD1 leads to deficient grain setting in rice, likely as a consequence of reduced sugar transport resulting from decreased PD permeability in phloem companion cells.     

Tyloses and Phenolic Deposits in Xylem Vessels Impede Water Transport in Low-Lignin Transgenic Poplars: A Study by Cryo-Fluorescence Microscopy

Of 14 transgenic poplar genotypes (Populus tremula × Populus alba) with antisense 4-coumarate:coenzyme A ligase that were grown in the field for 2 years, five that had substantial lignin reductions also had greatly reduced xylem-specific conductivity compared with that of control trees and those transgenic events with small reductions in lignin. For the two events with the lowest xylem lignin contents (greater than 40% reduction), we used light microscopy methods and acid fuchsin dye ascent studies to clarify what caused their reduced transport efficiency. A novel protocol involving dye stabilization and cryo-fluorescence microscopy enabled us to visualize the dye at the cellular level and to identify water-conducting pathways in the xylem. Cryo-fixed branch segments were planed in the frozen state on a sliding cryo-microtome and observed with an epifluorescence microscope equipped with a cryo-stage. We could then distinguish clearly between phenolic-occluded vessels, conductive (stain-filled) vessels, and nonconductive (water- or gas-filled) vessels. Low-lignin trees contained areas of nonconductive, brown xylem with patches of collapsed cells and patches of noncollapsed cells filled with phenolics. In contrast, phenolics and nonconductive vessels were rarely observed in normal colored wood of the low-lignin events. The results of cryo-fluorescence light microscopy were supported by observations with a confocal microscope after freeze drying of cryo-planed samples. Moreover, after extraction of the phenolics, confocal microscopy revealed that many of the vessels in the nonconductive xylem were blocked with tyloses. We conclude that reduced transport efficiency of the transgenic low-lignin xylem was largely caused by blockages from tyloses and phenolic deposits within vessels rather than by xylem collapse.Secondary xylem in woody plants is a complex vascular tissue that functions in mechanical support, conduction, storage, and protection (Carlquist, 2001; Tyree and Zimmermann, 2002). The xylem must provide a sufficient and safe water supply throughout the entire pathway from roots to leaves for transpiration and photosynthesis. It is well established that enhanced water conductivity of xylem can increase total plant carbon gain (Domec and Gartner, 2003; Santiago et al., 2004; Brodribb and Holbrook, 2005a). According to the Hagen-Poiseuille equation, xylem conductivity should scale with vessel lumen diameter to the fourth power (Tyree and Zimmermann, 2002). Indeed, xylem conductivity largely depends on anatomical features, including conduit diameters and frequencies (Salleo et al., 1985; McCulloh and Sperry, 2005). However, there are hydraulic limits to maximum vessel diameters, because xylem conduits have to withstand the strong negative pressures of the transpiration stream that could cause cell collapse or embolisms within vessels that are structurally inadequate to withstand these forces (Tyree and Sperry, 1989; Lo Gullo et al., 1995; Hacke et al., 2000). To some extent, stomatal regulation of transpiration limits the negative pressures that the xylem experiences (Tardieu and Davies, 1993; Cochard et al., 2002; Meinzer, 2002; Brodribb and Holbrook, 2004; Buckley, 2005; Franks et al., 2007; Woodruff et al., 2007). Nevertheless, plants rely on an array of structural reinforcements of xylem to ensure the safety of water transport. The size of xylem elements, vessel redundancy, intervessel pit and membrane geometries, and the thickness, microstructure, and chemical composition of cell walls are among the features that regulate tradeoffs between efficiency and safety of xylem water transport (Baas and Schweingruber, 1987; Hacke et al., 2001; Domec et al., 2006; Ewers et al., 2007; Choat et al., 2008).The xylem cell wall is made up of cellulose bundles that are hydrogen bonded with hemicelluloses, which are in turn embedded within a lignin matrix (Mansfield, 2009; Salmén and Burgert, 2009). Besides providing this matrix for the cell wall itself, lignin is thought to contribute to many of the mechanical and physical characteristics of wood as well as conferring passive resistance to the spread of pathogens within a plant (Niklas, 1992; Boyce et al., 2004; Davin et al., 2008). Lignin typically represents 20% to 30% of the dry mass of wood and therefore is among the most abundant stores of carbon in the biosphere (Zobel and van Buijtenen, 1989). The complex molecular structure and biosynthetic pathway of various types of lignins have been studied extensively (Boerjan et al., 2003; Ralph et al., 2004, 2007; Higuchi, 2006; Boudet, 2007; Davin et al., 2008). The monomeric composition of lignin varies between different cell types of the same species depending on the functional specialization of the cell (Yoshinaga et al., 1992; Watanabe et al., 2004; Xu et al., 2006). The composition and amount of lignin in wild plants varies in response to climatic conditions (Donaldson, 2002) or gravitational and mechanical demands (Pruyn et al., 2000; Kern et al., 2005; Rüggeberg et al., 2008). It is clear that plants are capable of regulating the lignification pattern in differentiating cells, which provides them with flexibility for responding to environmental stresses (Donaldson, 2002; Koehler and Telewski, 2006; Ralph et al., 2007; for review, see Vanholme et al., 2008).Whereas some level of lignin is a requisite for all vascular plants, it is often an unwanted product in the pulp and paper industry because it increases the costs of paper production and associated water treatments necessary for environmental protection (Chen et al., 2001; Baucher et al., 2003; Peter et al., 2007). Reducing the lignin content of the raw biomass material may allow more efficient hydrolysis of polysaccharides in biomass and thus facilitate the production of biofuel (Chen and Dixon, 2007). With the ultimate goal of development of wood for more efficient processing, much research has been aimed at the production of genetically modified trees with altered lignin biosynthesis (Boerjan et al., 2003; Boudet et al., 2003; Li et al., 2003; Halpin, 2004; Ralph et al., 2004, 2008; Chiang, 2006; Coleman et al., 2008a, 2008b; Vanholme et al., 2008; Wagner et al., 2009). It is now technically possible to achieve more than 50% reductions of lignin content in xylem of poplar (Populus spp.; Leplé et al., 2007; Coleman et al., 2008a, 2008b), but the consequences of such reduction on plant function have received relatively little attention (Koehler and Telewski, 2006). In-depth studies on the xylem structure and functional performance of transgenic plants with low lignin are limited, despite the need to assess their long-term sustainability for large-scale production (Anterola and Lewis, 2002; Hancock et al., 2007; Coleman et al., 2008b, Voelker, 2009; Horvath et al., 2010).Genetically modified plants are suitable models for studying fundamental questions of the physiological role of lignin because of the possibility of controlling lignification without the confounding effects encountered when comparing across plant tissues or stages of development (Koehler and Telewski, 2006; Leplé et al., 2007; Coleman et al., 2008b). Research on Arabidopsis (Arabidopsis thaliana) and tobacco (Nicotiana tabacum) has shown that down-regulation of lignin biosynthesis can have diverse effects on plant metabolism and structure, including changes in the lignin amount and composition (p-hydroxyphenyl/guaiacyl/syringyl units ratio) as well as the collapse of xylem vessel elements (Lee et al., 1997; Sewalt et al., 1997; Piquemal et al., 1998; Chabannes et al., 2001; Jones et al., 2001; Franke et al., 2002; Dauwe et al., 2007). Among temperate hardwoods, poplar has been established as a model tree for genetic manipulations because of its ecological and economic importance, fast growth, ease of vegetative propagation, and its widespread use in traditional breeding programs (Bradshaw et al., 2001; Brunner et al., 2004). The question of how manipulation of lignin can affect the anatomy and physiological function of xylem in poplar has been addressed in part by several research groups (Anterola and Lewis, 2002; Boerjan et al., 2003; Leplé et al., 2007; Coleman et al., 2008b). Some studies that involved large lignin reductions reported no significant alterations in the xylem anatomy (Hu et al., 1999; Li et al., 2003). However, in many other experiments, reduced total lignin content was associated with significant growth retardation, alterations in the lignin monomer composition, irregularities in the xylem structure (Anterola and Lewis, 2002; Leplé et al., 2007; Coleman et al., 2008b), and the patchy occurrence of collapsed xylem cells (Coleman et al., 2008b; Voelker, 2009). Furthermore, severely down-regulated lignin biosynthesis has resulted in greatly reduced xylem water-transport efficiency (Coleman et al., 2008b; Lachenbruch et al., 2009; Voelker, 2009). It is generally assumed that the reduced water transport ability of xylem with very low lignin contents is caused by collapsed conduits and/or increased embolism due to the entry of air bubbles into the water-conducting cells (Coleman et al., 2008b; Wagner et al., 2009), but detailed anatomical investigations of the causes of impaired xylem conductivity of low-lignin trees are lacking. Analysis of the anatomical basis for the properties of xylem conduits in plants with genetically manipulated amounts and composition of lignin can provide a deeper understanding of the physiological role of lignin as well as the lower limit of down-regulation of lignin biosynthesis at which trees can still survive within natural environments.One of the approaches for the suppression of lignin biosynthesis is down-regulation of 4-coumarate:coenzyme A ligase (4CL), an enzyme that functions in phenylpropanoid metabolism by producing the monolignol precursor p-coumaroyl-CoA (Kajita et al.,1997; Allina et al., 1998; Hu et al., 1998; Harding et al., 2002; Jia et al., 2004; Costa et al., 2005; Friedmann et al., 2007; Wagner et al., 2009). In a 2-year-long field trial on the physiological performance of poplar (Populus tremula × Populus alba) transgenic clones, out of 14 genotypes with altered lignin biosynthesis (down-regulated 4CL), five showed dramatically reduced wood-specific conductivity (k_s) compared with that of control trees (Voelker, 2009). Those mutants with the severely reduced k_s were also characterized by having the lowest wood lignin contents (up to an approximately 40% reduction) in the study. Trees with transgenic events characterized by the formation of abnormally brown wood exhibited regular branch dieback at the end of the growing season, despite having been regularly watered (Voelker, 2009). Our objective was to identify the structural features responsible for reduced transport efficiency in the xylem of transgenic poplars with extremely low lignin contents. We employed fluorescence and laser scanning confocal microscopy for anatomical analyses of xylem structure as well as dye-flow experiments followed by cryo-fluorescence microscopy to visualize the functioning water-conductive pathways in xylem at the cellular level. We report the frequent occurrence of tyloses and phenolic depositions in xylem vessels of strongly down-regulated trees that may be the cause of their reduced xylem conductivity.     

Phloem as Capacitor: Radial Transfer of Water into Xylem of Tree Stems Occurs via Symplastic Transport in Ray Parenchyma

The transfer of water from phloem into xylem is thought to mitigate increasing hydraulic tension in the vascular system of trees during the diel cycle of transpiration. Although a putative plant function, to date there is no direct evidence of such water transfer or the contributing pathways. Here, we trace the radial flow of water from the phloem into the xylem and investigate its diel variation. Introducing a fluorescent dye (0.1% [w/w] fluorescein) into the phloem water of the tree species Eucalyptus saligna allowed localization of the dye in phloem and xylem tissues using confocal laser scanning microscopy. Our results show that the majority of water transferred between the two tissues is facilitated via the symplast of horizontal ray parenchyma cells. The method also permitted assessment of the radial transfer of water during the diel cycle, where changes in water potential gradients between phloem and xylem determine the extent and direction of radial transfer. When injected during the morning, when xylem water potential rapidly declined, fluorescein was translocated, on average, farther into mature xylem (447 ± 188 µm) compared with nighttime, when xylem water potential was close to zero (155 ± 42 µm). These findings provide empirical evidence to support theoretical predictions of the role of phloem-xylem water transfer in the hydraulic functioning of plants. This method enables investigation of the role of phloem tissue as a dynamic capacitor for water storage and transfer and its contribution toward the maintenance of the functional integrity of xylem in trees.Physiological and hydraulic functioning of the two long-distance transport systems in trees, xylem and phloem, have intrigued plant researchers for more than a century. Since the pioneering work of Dixon and Joly (1895; cohesion-tension theory for xylem) and Münch (1930; pressure flow hypothesis for phloem), the majority of studies have investigated these systems independently of each other. Although the work of Stout and Hoagland (1939) as well as Biddulph and Markle (1944) laid the foundation for the physiological nexus between xylem and phloem, it is only recently that we have begun to understand the importance of the hydraulic nexus (Hölttä et al., 2006, 2009; Sevanto et al., 2011, 2014). Processes related to both nexus occur in parallel, and here the term physiological nexus covers all metabolite exchange, including the bidirectional flow of amino acids, minerals, and carbohydrates (Wardlaw, 1974; Ferrier et al., 1975; Pfautsch et al., 2009, 2015; De Schepper et al., 2013; for review, see van Bel, 1990, 2003). The term hydraulic nexus refers to the function of phloem as a capacitor, where water stored in phloem moves into xylem vessels to maintain the integrity of the transpiration stream (Zweifel et al., 2000, and refs. therein). Throughout this article, we use the term phloem collectively for cells that make up the transport phloem of woody plants (including sieve element/companion cell complexes, parenchyma cells, etc.), as opposed to collection and release phloem tissue, which differ in structure and function. Transport phloem is characterized by the retention of “high hydrostatic pressure by retrieval of leaked osmotica accompanied by water flux” (Patrick, 2013).According to the cohesion-tension theory, water in xylem vessels is constantly under tension and moves in a metastable state from roots to leaves along a hydrostatic pressure gradient. Depending on both the availability of soil moisture and the vapor pressure deficit of the atmosphere, this tension can exceed the cohesive forces that bind water molecules, resulting in the formation of a gas void that, after expanding, can lead to rupture of the water column inside individual vessels (termed cavitation; Zimmermann, 1983). Once cavitated, vessels become dysfunctional, and the transport of water and nutrients to leaves declines. However, water stored in woody tissues of trees can be mobilized to alleviate the risk of cavitation, and recent theory suggests that both water and carbohydrates from phloem may aid in the reversal of vessel embolism (i.e. air intrusion), although the evidence is indirect (Salleo et al., 2009; Brodersen et al., 2010; Nardini et al., 2011).All parts of plants have a water storage capacity (symplastic and apoplastic), and this capacitance increases with tree size and age (Phillips et al., 2003). The ability to mobilize stored water varies according to the force required to drag it out of storage (Holbrook, 1995). One-half century ago, Reynolds (1965) highlighted the importance of the volume of internally stored water to support the transpiration of trees. Since then, studies have quantified the fraction of stored water in total daily transpiration for a range of tree species. This fraction varies between 2% and 20% (Tyree and Yang, 1990; Čermák et al., 2007, and refs. therein; Barnard et al., 2011; Pfautsch and Adams, 2013) and is generally smaller in angiosperms compared with gymnosperms, where a maximum fraction of 50% was reported for Pinus sylvestris (Waring et al., 1979). Given that the daily water use of large adult trees can easily reach 260 to 380 L (Čermák et al., 2007; Pfautsch et al., 2011), considerable volumes of stored water must be mobilized from and restored back into capacitors on a daily basis. Remobilization of stored water also can prolong stomatal opening and thus increase carbon gain (Goldstein et al., 1998).The volume of stored water released depends on the elasticity of the storage tissue, its connectivity to xylem vessels, and the gradient of water potential (ψ) between the storage tissue and vessels. Cells with elastic walls represent ideal capacitors because they can change their volume as a consequence of small changes in ψ. Thus, phloem, cambium, and juvenile xylem cells are well suited for water storage and release (Yang and Tyree, 1992; Zweifel et al., 2014). The magnitude of release and refill of stored water in trees can be approximated by separately measuring the change in thickness of phloem and xylem during a diel cycle using high-precision dendrometers (Zweifel et al., 2014). Whitehead and Jarvis (1981) have calculated that around 90% of the diurnal change in stem radius can be attributed to changes in the water content of cambial and phloem tissues. To date, it is commonly accepted that tree bark, independent of the wood below, swells during the night and shrinks during the day (Zweifel et al., 2000), reflecting the water flow dynamics that characterize the dynamic exchange of water between phloem and xylem.Phloem and xylem tissues are separated by rows of intermediary cambial cells. However, depending on the species, phloem and xylem are connected through uniseriate or multiseriate strands of radially aligned ray parenchyma cells, commonly termed wood rays. These rays have been shown to be capable of symplastic water transport through plasmodesmata (Höll, 1975). Based on measurements of radial conductance, Sevanto et al. (2011) suggested that aquaporins also might be involved in the radial transfer of water. Both theoretical and experimental approaches have been developed to better understand the dynamic exchange of water between xylem and phloem. Hölttä et al. (2006, 2009) developed a model based on Münch’s hypothesis and included a term that represents the hydraulic connection between the two tissue types. Through incorporating this term, model outputs suggest the occurrence of a constant exchange of water between xylem and phloem along gradients of ψ. However, some authors suggested that changes in ψ of xylem alone might be insufficient to account for the observed diurnal shrinkage and swelling of bark (Sevanto et al., 2003). Along this line of argument, loading and unloading of carbohydrates in phloem tissue has been suggested to further impact the radial transfer of water and associated changes in bark thickness (Mencuccini et al., 2013).Nevertheless, to date, all approaches remain indirect, and the routes of water transfer between phloem and xylem have yet to be determined. Here, we present a technique that enables the visualization of water transfer from phloem to xylem tissues and resolves the apoplastic and symplastic pathways and cell types. The method involves the injection of an aqueous solution that contains fluorescent dye into phloem followed by analyses of woody tissues using confocal laser scanning microscopy. We assess the effectiveness of three different dyes to stain possible transfer pathways. We also introduce dye during different time intervals of the diel transpiration cycle to test the effect of predicted dynamic changes in ψ between phloem and xylem on the transfer processes. We hypothesized that radial transfer of water would be most pronounced during periods where conditions of the hydraulic nexus between phloem and xylem differ the most. These differences are expected during high rates of transpiration that cause a steep decline in xylem ψ, commonly observed during morning hours. We use leaf water potential (ψ_L) and high-precision dendrometer measurements to identify relevant time intervals. The simultaneous assessment of ψ_L and the independent diameter fluctuation of phloem and xylem may provide empirical evidence for the role of phloem as a water storage capacitor that helps mitigate increasing tension in the transpiration stream.     

The HvNramp5 Transporter Mediates Uptake of Cadmium and Manganese,But Not Iron

The Natural Resistance Associated Macrophage Protein (Nramp) represents a transporter family for metal ions in all organisms. Here, we functionally characterized a member of Nramp family in barley (Hordeum vulgare), HvNramp5. This member showed different expression patterns, transport substrate specificity, and cellular localization from its close homolog in rice (Oryza sativa), OsNramp5, although HvNramp5 was also localized to the plasma membrane. HvNramp5 was mainly expressed in the roots and its expression was not affected by Cd and deficiency of Zn, Cu, and Mn, but slightly up-regulated by Fe deficiency. Spatial expression analysis showed that the expression of HvNramp5 was higher in the root tips than that in the basal root regions. Furthermore, analysis with laser microdissection revealed higher expression of HvNramp5 in the outer root cell layers. HvNramp5 showed transport activity for both Mn²⁺ and Cd²⁺, but not for Fe²⁺ when expressed in yeast. Immunostaining with a HvNramp5 antibody showed that this protein was localized in the root epidermal cells without polarity. Knockdown of HvNramp5 in barley resulted in a significant reduction in the seedling growth at low Mn supply, but this reduction was rescued at high Mn supply. The concentration of Mn and Cd, but not other metals including Cu, Zn, and Fe, was decreased in both the roots and shoots of knockdown lines compared with the wild-type barley. These results indicate that HvNramp5 is a transporter required for uptake of Mn and Cd, but not for Fe, and that barley has a distinct uptake system from rice.Transport of mineral elements from soil to different organs and tissues of plants requires different types of transporters (Hall and Williams, 2003; Nevo and Nelson, 2006; Yokosho et al., 2009; Olsen and Palmgren, 2014; Sasaki et al., 2016), which include the zinc-regulated transporters, iron-regulated transporter-like protein family; the natural resistance-associated macrophage protein (Nramp) family of transporters; the multidrug and toxic compound extrusion protein transporters; the heavy metal ATPase transporters; the oligopeptide transporters family; the ATP-binding cassette family of transporters; and the cation-diffusion facilitator family of transporters. Among them, Nramp represents a transporter family for metal ion in all organisms including bacteria, animals, and plants (Curie et al., 2000; Nevo and Nelson, 2006). Some members of this family in plants have been functionally characterized, especially in model plants such as Arabidopsis (Arabidopsis thaliana) and rice (Oryza sativa). In Arabidopsis, there are six members of Nramp transporter proteins. AtNramp1 is localized to the plasma membrane of root cells and functions as a high-affinity transporter for Mn uptake (Cailliatte et al., 2010). Both AtNramp3 and AtNramp4 are localized to the tonoplast and play redundant roles in Fe, exporting from the vacuole during seed germination and in Mn homeostasis at the adult stage (Thomine et al., 2000; Lanquar et al., 2005, 2010). AtNramp6 is targeted to a vesicular-shaped endomembrane compartment and is implicated in the distribution or availability of Cd within cells (Cailliatte et al., 2009). However, the function of AtNramp2 and AtNramp5 has not been characterized.On the other hand, there are seven members of Nramp transporter family in the rice genome, of which four have been functionally characterized. They all are localized to the plasma membrane but show different roles. OsNramp1 shows transport activity for Fe and Cd in yeast and is proposed to be involved in Cd accumulation (Takahashi et al., 2011). OsNramp3 is localized at the vascular tissues of nodes and plays an important role in distribution of Mn, but not Fe and Cd (Yamaji et al., 2013). On the other hand, OsNrat1 (OsNramp4) transports trivalent Al ion (Xia et al., 2010) and is required for high Al tolerance in rice roots. Finally, OsNramp5 functions as a major transporter responsible for root Mn and Cd uptake (Ishimaru et al., 2012; Ishikawa et al., 2012; Sasaki et al., 2012). However, the function of OsNramp2, OsNramp6, and OsNramp7 is unknown.In addition to Nramp members characterized in rice and Arabidopsis, some members in other plant species have also been characterized. For example, a soybean (Glycine max) Nramp transporter, GmDMT1 is implicated in the ferrous iron transport (Kaiser et al., 2003). Nramp1 isolated from Noccaea caerulescens, a Zn/Cd hyperaccumulator, is involved in the influx of Cd across the endodermal plasma membrane and plays a key role in Cd flux into the stele and root-to-shoot Cd transport (Milner et al., 2014). In Malus baccata, Nramp1 is capable of mediating the distribution of ions as well as transport of Fe, Mn, and Cd (Xiao et al., 2008). Besides, Nramp1 and Nramp3 in tomato (Solanum lycopersicum) have also been suggested to be involved in Mn transport (Bereczky et al., 2003). When NcNramp3 and NcNramp4 from Noccaea caerulescens were expressed in yeast, NcNramp3 transported Fe, Mn, and Cd, while NcNramp4 also transported Zn in addition to Fe, Mn, and Cd (Oomen et al., 2009). However, Nramp4 isolated from Thlaspi japonicum, a Ni hyperaccumulator, showed transport activity for Ni but not for Zn, Cd, or Mn in yeast (Mizuno et al., 2005). These findings indicate that Nramp members have a diverse role in metal transport in plants.Barley (Hordeum vulgare) is the fourth most important cereal crop in the world; however, less progress has been made in understating of molecular mechanisms on mineral element transport in barley due to its large genome size. For example, no Nramp members in barley have been functionally characterized so far. In this study, we first isolated barley Nramp member, HvNramp5, which is a close homolog of rice OsNramp5. Detailed functional analysis revealed that HvNramp5 is involved in the uptake of both Mn and Cd, but not of Fe in barley roots. Furthermore, we found that different from OsNramp5, HvNramp5 showed a distinct pattern in the gene expression, cellular localization, and transport substrate.     

Turnip mosaic virus Moves Systemically through Both Phloem and Xylem as Membrane-Associated Complexes

Plant viruses move systemically in plants through the phloem. They move as virions or as ribonucleic protein complexes, although it is not clear what these complexes are made of. The approximately 10-kb RNA genome of Turnip mosaic virus (TuMV) encodes a membrane protein, known as 6K₂, that induces endomembrane rearrangements for the formation of viral replication factories. These factories take the form of vesicles that contain viral RNA (vRNA) and viral replication proteins. In this study, we report the presence of 6K₂-tagged vesicles containing vRNA and the vRNA-dependent RNA polymerase in phloem sieve elements and in xylem vessels. Transmission electron microscopy observations showed the presence in the xylem vessels of vRNA-containing vesicles that were associated with viral particles. Stem-girdling experiments, which leave xylem vessels intact but destroy the surrounding tissues, confirmed that TuMV could establish a systemic infection of the plant by going through xylem vessels. Phloem sieve elements and xylem vessels from Potato virus X-infected plants also contained lipid-associated nonencapsidated vRNA, indicating that the presence of membrane-associated ribonucleic protein complexes in the phloem and xylem may not be limited to TuMV. Collectively, these studies indicate that viral replication factories could end up in the phloem and the xylem.Plant viruses use the host preexisting transport routes to propagate infection to the whole plant. After replication in the initially infected cells, viruses move cell to cell through plasmodesmata (PD) and start a new round of replication in the newly infected cells. This cycle is repeated until viruses reach vascular tissues, where they enter into the conducting tubes for systemic movement. Several studies have indicated that plant viruses are passively transported along the source-to-sink flow of photoassimilates and thus are believed to move systemically through the phloem (for review, see Hipper et al., 2013).The conducting tube of the phloem is the sieve element. The mature sieve element is enucleated and relies on the associated companion cells for the maintenance of its physiological function (Fisher et al., 1992). The specialized PD connecting one sieve element with one companion cell is called the pore plasmodesmal unit (PPU). Different from the other PDs, PPUs are always branched on the companion cell side but have only one channel on the sieve element side (Oparka and Turgeon, 1999). It is believed that the loading and uploading of viral material during phloem transport are through PPUs. Even though the size exclusion limit of PPUs (Kempers and Bel, 1997) is larger than that of the other PDs (Wolf et al., 1989; Derrick et al., 1990), PPUs should not allow, in their native state, virions or viral ribonucleoprotein (vRNP) complexes to pass through. It is thus believed that specific interactions between virus and host factors are required to allow the viral entity to go through. For instance, the movement protein of Cucumber mosaic virus (CMV) is targeted to PPUs (Blackman et al., 1998), suggesting that this viral protein modifies the size exclusion limit of PPUs and helps viral entry into sieve elements.Most plant viruses are assumed to move systemically through the phloem as virions. This assumption is based on the observation that Coat Protein (CP) deletions debilitating virus assembly prevent systemic infection (Brault et al., 2003; Zhang et al., 2013; Hipper et al., 2014). Some investigations showed the actual presence of virions in sieve elements. This is the case for the icosahedral Tobacco ringspot virus (Halk and McGuire, 1973), Carrot red leaf virus (Murant and Roberts, 1979), Potato leaf roll virus (Shepardson et al., 1980), and Beet western yellows virus (Hoefert, 1984). In addition, virions also were observed in phloem sap, such as the icosahedral CMV (Requena et al., 2006) and the rigid rod-shaped Cucumber green mottle mosaic virus (Simón-Buela and García-Arenal, 1999). Some viruses also are believed to move as ribonucleic protein (RNP) complexes, since systemic movement was observed in CP mutants where virion assembly was hindered. For instance, Tobacco rattle virus, Potato mop-top virus, Brome mosaic virus, and Tomato bushy stunt virus can still move systemically when the CP gene has been deleted from the viral genome (Swanson et al., 2002; Savenkov et al., 2003; Gopinath and Kao, 2007; Manabayeva et al., 2013). For potyviruses, it is still not clear if long-distance transport involves exclusively viral particles or if vRNP complexes also are implicated (Dolja et al., 1994, 1995; Cronin et al., 1995; Schaad et al., 1997; Kasschau and Carrington, 2001; Rajamaki and Valkonen, 2002). But whether virions or vRNP complexes are involved in viral movement, the full nature of the viral entity being implicated has not been defined.Xylem also is used for systemic infection of viruses, but its importance in viral transport generally has been overlooked. Vessel elements are the building blocks of xylem vessels, which constitute the major part of the water-upward-transporting system in a plant. The side walls of mature vessel elements contain pits, which are areas lacking a secondary cell wall; the end walls of the mature vessel elements are removed, and the openings are called perforation plates (Roberts and McCann, 2000). CP or virions of some plant viruses of all different shapes have been detected in the xylem vessels and/or guttation fluid, suggesting that these viruses may move systemically through xylem vessels. For example, the CP of the icosahedral Tomato bushy stunt virus (Manabayeva et al., 2013) and Rice yellow mottle virus (Opalka et al., 1998), the CP of the rigid rod-shaped Soilborne wheat mosaic virus (Verchot et al., 2001) and Cucumber green mottle mosaic virus (Moreno et al., 2004), and the flexuous rod-shaped Potato virus X (PVX; Betti et al., 2012) were detected in xylem vessels. Colocalization of anti-Rice yellow mottle virus antibodies and a cell wall marker for cellulosic β-(1-4)-d-glucans over vessel pit membranes suggests that the pit membranes might be a pathway for virus migration between vessels (Opalka et al., 1998). Moreover, flexuous rod-shaped virions of Zucchini yellow mosaic virus were found in both xylem vessels of root tissue and the guttation fluid (French and Elder, 1999). Finally, icosahedral Brome mosaic virus (Ding et al., 2001) and rigid rod-shaped Tomato mosaic virus and Pepper mild mottle virus (French et al., 1993) virions were found in guttation fluid. Guttation fluid originates from xylem exudate, indicating that these plant viruses can move through xylem within the infected plant. The above studies, however, mainly relied on electron microscopy and infection assays and may have missed the presence of other viral components that might be involved in transport.Turnip mosaic virus (TuMV) is a positive-strand RNA virus belonging to the family Potyviridae, genus Potyvirus, which contains around 30% of the currently known plant viruses and causes serious diseases in numerous crops (Shukla et al., 1994). Potyviruses are nonenveloped, flexuous rod-shaped particles of 680 to 900 nm in length and 11 to 13 nm in diameter. The genomic approximately 10-kb RNA encodes a polyprotein, which is processed into at least 11 mature proteins. TuMV remodels cellular membranes into viral factories, which are intracellular compartments involved in viral replication and movement. These compartments take the form of vesicles of approximately 100 nm in diameter originating from the endoplasmic reticulum (Grangeon et al., 2012). These vesicles contain viral RNA (vRNA) and viral and host proteins involved in vRNA replication (Beauchemin et al., 2007; Beauchemin and Laliberté, 2007; Dufresne et al., 2008; Huang et al., 2010; Grangeon et al., 2012). The viral membrane 6K₂ protein is involved in the membrane alterations and vesicle production (Beauchemin et al., 2007). The membrane-bound replication complexes can move intracellularly and cell to cell (Grangeon et al., 2013) at a rate of one cell being infected every 3 h (Agbeci et al., 2013). Intercellular trafficking of the replication complex is likely mediated by the PD-localized potyviral proteins Cytoplasmic Inclusion (CI) and P3N-PIPO (for N-terminal Half of P3 fused to the Pretty Interesting Potyviridae ORF; Carrington et al., 1998; Wei et al., 2010; Vijayapalani et al., 2012) as well as CP (Dolja et al., 1994, 1995), Viral Protein genome-linked (VPg; Nicolas et al., 1997; Rajamaki and Valkonen, 1999, 2002), and Helper Component-Proteinase (HC-Pro; Cronin et al., 1995; Kasschau et al., 1997; Rojas et al., 1997; Kasschau and Carrington, 2001), which are involved in both cell-to-cell and vascular movement.It is expected that, ultimately, TuMV reaches the vascular tissues of the plant, but how and under what form it is released into the conducting tubes are not known. To further understand viral spread and systemic movement, we investigated the distribution of 6K₂-tagged TuMV factories in all of the leaf and stem tissues other than the epidermal cells. We found TuMV factories in all tissues. Interestingly, we observed 6K₂-tagged vesicles, containing vRNA and viral replication proteins, in both phloem sieve elements and xylem vessels. We confirmed that TuMV could move systemically through xylem by a so-called stem-girdling assay, which induces cell death of the phloem without affecting xylem integrity. Hence, our study indicates that membrane-associated TuMV replication complexes are involved in the systemic movement of the virus.     

Multispectral Phloem-Mobile Probes: Properties and Applications

Using Arabidopsis (Arabidopsis thaliana) seedlings, we identified a range of small fluorescent probes that entered the translocation stream and were unloaded at the root tip. These probes had absorbance/emission maxima ranging from 367/454 to 546/576 nm and represent a versatile toolbox for studying phloem transport. Of the probes that we tested, naturally occurring fluorescent coumarin glucosides (esculin and fraxin) were phloem loaded and transported in oocytes by the sucrose transporter, AtSUC2. Arabidopsis plants in which AtSUC2 was replaced with barley (Hordeum vulgare) sucrose transporter (HvSUT1), which does not transport esculin in oocytes, failed to load esculin into the phloem. In wild-type plants, the fluorescence of esculin decayed to background levels about 2 h after phloem unloading, making it a suitable tracer for pulse-labeling studies of phloem transport. We identified additional probes, such as carboxytetraethylrhodamine, a red fluorescent probe that, unlike esculin, was stable for several hours after phloem unloading and could be used to study phloem transport in Arabidopsis lines expressing green fluorescent protein.The phloem of higher plants consists of a series of longitudinally arranged sieve elements (SEs), companion cells (CCs), and associated parenchyma elements (Heo et al., 2014). The SEs translocate a diverse range of solutes, proteins, and RNAs from source to sink organs and perform key roles in solute delivery and signaling (Turgeon and Wolf, 2009; Ham and Lucas, 2014). The phloem is a delicate tissue, and examining its structure and function has proven to be a difficult task (Knoblauch and Oparka, 2012; Truernit, 2014). Arguably, the most reliable way to assess the rate of phloem transport in different organs is by using radiolabeled solutes derived photosynthetically from ¹⁴CO₂ (Kölling et al., 2013). In parallel, autoradiography provides a valuable means of imaging the distribution of radiolabeled solutes in different tissues (Housley and Fisher, 1975; Kölling et al., 2013). However, both of these methods are time consuming and limited by resolution. In the last decade, the use of fluorescent tracers has become prominent, allowing phloem transport to be imaged in living SEs with significantly improved resolution above autoradiography (Knoblauch and Oparka, 2012). Importantly, these probes cannot be used as substrates for Suc loading, which in many species, occurs by active, carrier-mediated transport (Turgeon and Wolf, 2009).Schumacher (1933) was the first plant biologist, to our knowledge, to study phloem transport using the fluorescent molecule fluorescein. Since then, however, only a few additional phloem-mobile probes have been discovered. Two such probes are carboxyfluorescein (CF; Grignon et al., 1989; Oparka et al., 1994) and 8-hydroxypyrene-1,3,6-trisulphonic acid (HPTS; Wright and Oparka, 1996). When applied in the ester (acetate) form, these probes are phloem mobile, although the exact mechanism by which they enter the phloem is unknown. In the case of CF, it is possible that this probe diffuses into the phloem and is retained in SEs by ion trapping (Wright and Oparka, 1996), a characteristic that it may share with many phloem-mobile herbicides (Hsu and Kleier, 1996). In contrast, HPTS is a highly charged molecule that should not cross membranes (Wright and Oparka, 1996), but it enters the phloem readily. Despite a lack of understanding of how these probes are loaded into the phloem, they have been used extensively in monitoring phloem transport (Knoblauch and van Bel, 1998). They have also found use in imaging symplastic pathways after unloading (Oparka et al., 1994; Roberts et al., 1997; Savage et al., 2013) and identifying symplastic domains in developing tissues (Gisel et al., 1999; Stadler et al., 2005). However, both CF and HPTS emit in the green spectrum, restricting their use for imaging movement in cells that express GFP as a reporter.The limited number of existing probes for phloem transport prompted us to explore unique small molecules differing in excitation and emission spectra as potential tracers. Using an Arabidopsis (Arabidopsis thaliana) seedling screen, we tested the phloem mobility of several small-molecule probes. In addition, we explored the use of esculin as a phloem-mobile tracer. Esculin is a fluorescent coumarin glucoside that is transported in oocytes by AtSUC2 (Sivitz et al., 2007), the major Suc transporter that loads the phloem in Arabidopsis (Gottwald et al., 2000). Here, we describe the development and application of a range of probes for monitoring phloem transport. These small probes cover absorbance/emission maxima ranging from 367/454 to 546/576 nm, allowing them to be used on plant material expressing different fluorescent reporter proteins. We describe the properties of these probes and show how they can be used in both pulse- and dual-labeling studies of phloem transport.     

Cavitation Resistance in Seedless Vascular Plants: The Structure and Function of Interconduit Pit Membranes

Plant water transport occurs through interconnected xylem conduits that are separated by partially digested regions in the cell wall known as pit membranes. These structures have a dual function. Their porous construction facilitates water movement between conduits while limiting the spread of air that may enter the conduits and render them dysfunctional during a drought. Pit membranes have been well studied in woody plants, but very little is known about their function in more ancient lineages such as seedless vascular plants. Here, we examine the relationships between conduit air seeding, pit hydraulic resistance, and pit anatomy in 10 species of ferns (pteridophytes) and two lycophytes. Air seeding pressures ranged from 0.8 ± 0.15 MPa (mean ± sd) in the hydric fern Athyrium filix-femina to 4.9 ± 0.94 MPa in Psilotum nudum, an epiphytic species. Notably, a positive correlation was found between conduit pit area and vulnerability to air seeding, suggesting that the rare-pit hypothesis explains air seeding in early-diverging lineages much as it does in many angiosperms. Pit area resistance was variable but averaged 54.6 MPa s m⁻¹ across all surveyed pteridophytes. End walls contributed 52% to the overall transport resistance, similar to the 56% in angiosperm vessels and 64% in conifer tracheids. Taken together, our data imply that, irrespective of phylogenetic placement, selection acted on transport efficiency in seedless vascular plants and woody plants in equal measure by compensating for shorter conduits in tracheid-bearing plants with more permeable pit membranes.Water transport in plants occurs under tension, which renders the xylem susceptible to air entry. This air seeding may lead to the rupture of water columns (cavitation) such that the air expands within conduits to create air-vapor embolisms that block further transport. (Zimmermann and Tyree, 2002). Excessive embolism such as that which occurs during a drought may jeopardize leaf hydration and lead to stomatal closure, overheating, wilting, and possibly death of the plant (Hubbard et al., 2001; Choat et al., 2012; Schymanski et al., 2013). Consequently, strong selection pressure resulted in compartmentalized and redundant plant vascular networks that are adapted to a species habitat water availability by way of life history strategy (i.e. phenology) or resistance to air seeding (Tyree et al., 1994; Mencuccini et al., 2010; Brodersen et al., 2012). The spread of drought-induced embolism is limited primarily by pit membranes, which are permeable, mesh-like regions in the primary cell wall that connect two adjacent conduits. The construction of the pit membrane is such that water easily moves across the membrane between conduits, but because of the small membrane pore size and the presence of a surface coating on the membrane (Pesacreta et al., 2005; Lee et al., 2012), the spread of air and gas bubbles is restricted up to a certain pressure threshold known as the air-seeding pressure (ASP). When xylem sap tension exceeds the air-seeding threshold, air can be aspirated from an air-filled conduit into a functional water-filled conduit through perhaps a large, preexisting pore or one that is created by tension-induced membrane stress (Rockwell et al., 2014). Air seeding leads to cavitation and embolism formation, with emboli potentially propagating throughout the xylem network (Tyree and Sperry, 1988; Brodersen et al., 2013). So, on the one hand, pit membranes are critical to controlling the spread of air throughout the vascular network, while on the other hand, they must facilitate the efficient flow of water between conduits (Choat et al., 2008; Domec et al., 2008; Pittermann et al., 2010; Schulte, 2012). Much is known about such hydraulic tradeoffs in the pit membranes of woody plants, but comparatively little data exist on seedless vascular plants such as ferns and lycophytes. Given that seedless vascular plants may bridge the evolutionary transition from bryophytes to woody plants, the lack of functional data on pit membrane structure in early-derived tracheophytes is a major gap in our understanding of the evolution of plant water transport.In woody plants, pit membranes fall into one of two categories: the torus-margo type found in most gymnosperms and the homogenous pit membrane characteristic of angiosperms (Choat et al., 2008; Choat and Pittermann, 2009). In conifers, water moves from one tracheid to another through the margo region of the membrane, with the torus sealing the pit aperture should one conduit become embolized. Air seeding occurs when water potential in the functional conduit drops low enough to dislodge the torus from its sealing position, letting air pass through the pit aperture into the water-filled tracheid (Domec et al., 2006; Delzon et al., 2010; Pittermann et al., 2010; Schulte, 2012; but see Jansen et al., 2012). Across north-temperate conifer species, larger pit apertures correlate with lower pit resistance to water flow (r_pit; MPa s m⁻¹), but it is the ratio of torus-aperture overlap that sets a species cavitation resistance (Pittermann et al., 2006, 2010; Domec et al., 2008; Hacke and Jansen, 2009). A similar though mechanistically different tradeoff exists in angiosperm pit membranes. Here, air seeding reflects a probabilistic relationship between membrane porosity and the total area of pit membranes present in the vessel walls. Specifically, the likelihood of air aspirating into a functional conduit is determined by the combination of xylem water potential and the diameter of the largest pore and/or the weakest zone in the cellulose matrix in the vessel’s array of pit membranes (Wheeler et al., 2005; Hacke et al., 2006; Christman et al., 2009; Rockwell et al., 2014). As it has come to be known, the rare-pit hypothesis suggests that the infrequent, large-diameter leaky pore giving rise to that rare pit reflects some combination of pit membrane traits such as variation in conduit membrane area (large or small), membrane properties (tight or porous), and hydrogel membrane chemistry (Hargrave et al., 1994; Choat et al., 2003; Wheeler et al., 2005; Hacke et al., 2006; Christman et al., 2009; Lee et al., 2012; Plavcová et al., 2013; Rockwell et al., 2014). The maximum pore size is critical because, per the Young-Laplace law, the larger the radius of curvature, the lower the air-water pressure difference under which the contained meniscus will fail (Jarbeau et al., 1995; Choat et al., 2003; Jansen et al., 2009). Consequently, angiosperms adapted to drier habitats may exhibit thicker, denser, smaller, and less abundant pit membranes than plants occupying regions with higher water availability (Wheeler et al., 2005; Hacke et al., 2007; Jansen et al., 2009; Lens et al., 2011; Scholz et al., 2013). However, despite these qualitative observations, there is no evidence that increased cavitation resistance arrives at the cost of higher r_pit. Indeed, the bulk of the data suggest that prevailing pit membrane porosity is decoupled from the presence of the single largest pore that allows air seeding to occur (Choat et al., 2003; Wheeler et al., 2005 Hacke et al., 2006, 2007).As water moves from one conduit to another, pit membranes offer considerable hydraulic resistance throughout the xylem network. On average, r_pit contributes 64% and 56% to transport resistance in conifers and angiosperms, respectively (Wheeler et al., 2005; Pittermann et al., 2006; Sperry et al., 2006). In conifers, the average r_pit is estimated at 6 ± 1 MPa s m⁻¹, almost 60 times lower than the 336 ± 81 MPa s m⁻¹ computed for angiosperms (Wheeler et al., 2005; Hacke et al., 2006; Sperry et al., 2006). Presumably, the high porosity of conifer pits compensates for the higher transport resistance offered by a vascular system composed of narrow, short, single-celled conduits (Pittermann et al., 2005; Sperry et al., 2006).Transport in seedless vascular plants presents an interesting conundrum because, with the exception of a handful of species, their primary xylem is composed of tracheids, the walls of which are occupied by homogenous pit membranes (Gibson et al., 1985; Carlquist and Schneider, 2001, 2007; but see Morrow and Dute, 1998, for torus-margo membranes in Botrychium spp.). At first pass, this combination of traits appears hydraulically maladaptive, but several studies have shown that ferns can exhibit transport capacities that are on par with more recently evolved plants (Wheeler et al., 2005; Watkins et al., 2010; Pittermann et al., 2011, 2013; Brodersen et al., 2012). Certainly, several taxa possess large-diameter, highly overlapping conduits, some even have vessels such as Pteridium aquilinum and many species have high conduit density, all of which could contribute to increased hydraulic efficiency (Wheeler et al., 2005; Pittermann et al., 2011, 2013). But how do the pit membranes of seedless vascular plants compare? Scanning electron micrographs of fern and lycopod xylem conduits suggest that they are thin, diaphanous, and susceptible to damage during specimen preparation (Carlquist and Schneider 2001, 2007). Consistent with such observations, two estimates of r_pit imply that r_pit in ferns may be significantly lower than in angiosperms; Wheeler et al. (2005) calculated r_pit in the fern Pteridium aquilinum at 31 MPa s m⁻¹, while Schulte et al. (1987) estimated r_pit at 1.99 MPa s m⁻¹ in the basal fern Psilotum nudum. The closest structural analogy to seedless vascular plant tracheids can be found in the secondary xylem of the early-derived vesselless angiosperms, in which tracheids possess homogenous pit membranes with r_pit values that at 16 MPa s m⁻¹ are marginally higher than those of conifers (Hacke et al., 2007). Given that xylem in seedless vascular plants is functionally similar to that in vesselless angiosperms, we expected convergent r_pit values in these two groups despite their phylogenetic distance. We tested this hypothesis, as well as the intrinsic cavitation resistance of conduits in seedless vascular plants, by scrutinizing the pit membranes of ferns and fern allies using the anatomical and experimental approaches applied previously to woody taxa. In particular, we focused on the relationship between pit membrane traits and cavitation resistance at the level of the individual conduit.     

Enhanced Sucrose Loading Improves Rice Yield by Increasing Grain Size

Yield in cereals is a function of grain number and size. Sucrose (Suc), the main carbohydrate product of photosynthesis in higher plants, is transported long distances from source leaves to sink organs such as seeds and roots. Here, we report that transgenic rice plants (Oryza sativa) expressing the Arabidopsis (Arabidopsis thaliana) phloem-specific Suc transporter (AtSUC2), which loads Suc into the phloem under control of the phloem protein2 promoter (pPP2), showed an increase in grain yield of up to 16% relative to wild-type plants in field trials. Compared with wild-type plants, pPP2::AtSUC2 plants had larger spikelet hulls and larger and heavier grains. Grain filling was accelerated in the transgenic plants, and more photoassimilate was transported from the leaves to the grain. In addition, microarray analyses revealed that carbohydrate, amino acid, and lipid metabolism was enhanced in the leaves and grain of pPP2::AtSUC2 plants. Thus, enhancing Suc loading represents a promising strategy to improve rice yield to feed the global population.Rice (Oryza sativa) is a staple food for nearly one-half of the global population. Given the rapid growth of the world’s population, there is an urgent need to increase rice yield. Rice yield is a complex trait that is directly associated with grain size, panicle number, and the number of grains per panicle (Xing and Zhang, 2010). Increasing grain size is a prime breeding target, and several genes known to control rice grain size, such as GRAIN SIZE3 (GS3), GS5, GW2 QTL for rice grain width and weight (GW2), GW8, and rice seed width5, have been identified (Fan et al., 2006; Song et al., 2007; Shomura et al., 2008; Li et al., 2011a; Wang et al., 2012). However, our knowledge of the mechanisms that control rice yield is limited. Thus, further improving rice yield remains a challenge for breeders (Sakamoto and Matsuoka, 2008). Identifying and characterizing unique genes or targets that regulate yield traits would improve our understanding of the molecular mechanisms that regulate yield traits and facilitate the breeding of new rice varieties with higher yields.The carbohydrates in rice grains originate from photosynthesis that is carried out predominantly in leaves (sources). Therefore, grain filling and rice yield depend on the efficient transport of carbohydrates from the leaves to seeds (sinks). In most plants, Suc is the main carbohydrate transported long distance in the veins to support the growth and development of roots, flowers, fruits, and seeds (Baker et al., 2012; Braun, 2012). Recently, the entire pathway for the export of Suc from leaves has been elucidated (Baker et al., 2012; Braun, 2012). Suc is synthesized in leaf mesophyll cells and diffuses from cell to cell through plasmodesmata until it reaches the phloem parenchyma cells (Slewinski and Braun, 2010). The SWEET transporters mediate Suc efflux from the phloem parenchyma cells into the apoplast, where Suc is subsequently loaded into the phloem sieve element-companion cell (SE/CC) complexes by Suc transporters (SUTs; Braun and Slewinski, 2009; Ayre, 2011; Chen et al., 2012). The resultant accumulation of Suc in sieve elements produces a hydrostatic pressure gradient that results in the bulk flow of Suc through a conduit of contiguous sieve elements, leading to its arrival and unloading in sink tissues (Lalonde et al., 2004; Baker et al., 2012).Genetic evidence has demonstrated that apoplastic Suc phloem loading is critical for growth, development, and reproduction in Arabidopsis (Arabidopsis thaliana). AtSWEET11 and AtSWEET12 are localized to the plasma membrane of the phloem and are expressed in a subset of phloem parenchyma cells in minor veins. These transporters mediate Suc efflux from phloem parenchyma cells into the apoplast prior to Suc uptake by SE/CC (Chen et al., 2012). The atsweet11 or atsweet12 single mutants exhibit no aberrant phenotypes, possibly due to genetic redundancy. However, atsweet11;12 double mutants are mildly chlorotic and display slower growth and higher levels of starch and sugar accumulation in the leaves than do wild-type plants (Chen et al., 2012). Arabidopsis phloem-specific sucrose transporter (AtSUC2) is a phloem-specific SUT that is expressed specifically in companion cells (Stadler and Sauer, 1996). AtSUC2 plays an essential role in phloem Suc loading and is necessary for efficient Suc transport from source to sink tissues in Arabidopsis (Stadler and Sauer, 1996; Gottwald et al., 2000; Srivastava et al., 2008). The atsuc2 mutants show stunted growth, retarded development, and sterility. Furthermore, these mutants accumulate excess starch in the leaves and fail to transport sugar efficiently to the roots and inflorescences (Gottwald et al., 2000).The proper control of carbohydrate partitioning is fundamental to crop yield (Braun, 2012). It has been reported that increasing sink grain strength by improving assimilate uptake capacity could be a promising approach toward obtaining higher yield. For example, seed-specific overexpression of a potato (Solanum tuberosum) SUT increased Suc uptake and growth rates of developing pea (Pisum sativum) cotyledons (Rosche et al., 2002). In addition, the Suc uptake capacity of grains and storage protein biosynthesis was increased in transgenic wheat (Triticum aestivum) plants expressing the barley (Hordeum vulgare) SUT HvSUT1 under the control of an endosperm-specific promoter (Weichert et al., 2010). Moreover, it was recently found that these transgenic wheat plants had a higher thousand grain weight and grain width and length, as well as a 28% increase in grain yield (Saalbach et al., 2014).Since the carbohydrates in rice grains originate from photosynthesis in source leaves, and carbohydrate partitioning from source leaves to heterotrophic sinks (e.g. seeds) is mediated by Suc transport in plants (Lalonde et al., 2004; Ayre, 2011), enhancing the capacity for Suc transport from leaves to seeds theoretically could increase crop yield. However, until now, enhancing Suc transport from leaves to seeds has not been shown to improve yield (Ainsworth and Bush, 2011).Here, we tested the hypothesis that enhancing Suc transport from leaves to seeds would increase rice yield. We expressed Arabidopsis SUC2 under control of the phloem protein2 promoter (pPP2) in rice and found that enhancing Suc loading did indeed increase rice yield. The pPP2::AtSUC2 plants produced larger grain than the wild type and showed grain yield increases of up to 16% in field trials. Our results suggest that manipulating phloem Suc transport is a useful strategy for increasing grain yield in rice and other cereal crops.     

Direct X-Ray Microtomography Observation Confirms the Induction of Embolism upon Xylem Cutting under Tension

Direct visualization shows enhanced embolism of xylem samples when they are collected under tension.Embolism resistance is a critically important trait for evaluating the ability of plants to survive and recover from drought periods and predicting future drought-induced forest decline (Choat et al., 2012). However, recent publications have provided evidence that some measurement techniques used to evaluate the hydraulic function and vulnerability to cavitation of plant organs may be prone to artifacts (Sperry et al., 2012; Cochard et al., 2013; Torres-Ruiz et al., 2014; Trifilò et al., 2014). The discovery of these artifacts has raised questions regarding the reliability of some previously published plant hydraulics data, in particular data relating to the refilling of embolized xylem conduits while the xylem is under tension. In this context, Wheeler et al. (2013) reported that sampling plant organs by cutting while the xylem is under tension can induce artificial increases in the degree of embolism at the moment of sample excision, even when cuts are made under water. The methodology applied by Wheeler et al. (2013), however, did not allow the visualization of embolized or functional vessels, and native embolism levels could not be determined in intact plants before any cutting was done.Whereas Scoffoni and Sack (2014) showed that the artifact described by Wheeler et al. (2013) has no impact on leaf xylem hydraulic conductance, there is some uncertainty about its importance in stems or shoots (Trifilò et al., 2014; Venturas et al., 2014). The results of Wheeler et al. (2013) indicate that more embolism could be induced by cutting samples that are under midrange xylem tension (e.g. at midday or under conditions of water stress). Potential overestimation of embolism due to changes in the xylem tension during the day has important implications for our understanding of plant water relations, since they could erroneously suggest that daily patterns of embolism formation and repair are routine in many woody plant species. Debate continues regarding the implications of a cutting artifact for the existence of a mechanism that allows plants to repair embolism while the xylem is under tension, so-called novel refilling (Salleo et al., 1996; Cochard and Delzon, 2013; Sperry, 2013; Delzon and Cochard, 2014). To avoid the excision artifact, Wheeler et al. (2013) recommended the relaxation of the xylem tension prior to excision by rehydrating plant tissue for anywhere between 2 min and 2 h. However, recent results from Trifilò et al. (2014) indicated that the rehydration procedures used by Wheeler et al. (2013) for relaxing the samples might favor xylem refilling and embolism repair (rehydration artifact), suggesting that the artifact resides in the relaxing procedure rather than in the cutting procedure. In light of these data, the assessment of the artifact described by Wheeler et al. (2013) using noninvasive techniques on intact plants, such as direct observation using x-ray microtomography (micro-CT; McElrone et al., 2013; Cochard et al., 2014) or magnetic resonance imaging (Choat et al., 2010; Zwieniecki et al., 2013), is useful to visually assess changes in embolism after cutting stems.     

Complexation of Arsenite with Phytochelatins Reduces Arsenite Efflux and Translocation from Roots to Shoots in Arabidopsis

Complexation of arsenite [As(III)] with phytochelatins (PCs) is an important mechanism employed by plants to detoxify As; how this complexation affects As mobility was little known. We used high-resolution inductively coupled plasma-mass spectrometry and accurate mass electrospray ionization-mass spectrometry coupled to HPLC to identify and quantify As(III)-thiol complexes and free thiol compounds in Arabidopsis (Arabidopsis thaliana) exposed to arsenate [As(V)]. As(V) was efficiently reduced to As(III) in roots. In wild-type roots, 69% of As was complexed as As(III)-PC₄, As(III)-PC₃, and As(III)-(PC₂)₂. Both the glutathione (GSH)-deficient mutant cad2-1 and the PC-deficient mutant cad1-3 were approximately 20 times more sensitive to As(V) than the wild type. In cad1-3 roots, only 8% of As was complexed with GSH as As(III)-(GS)₃ and no As(III)-PCs were detected, while in cad2-1 roots, As(III)-PCs accounted for only 25% of the total As. The two mutants had a greater As mobility, with a significantly higher accumulation of As(III) in shoots and 4.5 to 12 times higher shoot-to-root As concentration ratio than the wild type. Roots also effluxed a substantial proportion of the As(V) taken up as As(III) to the external medium, and this efflux was larger in the two mutants. Furthermore, when wild-type plants were exposed to l-buthionine sulfoximine or deprived of sulfur, both As(III) efflux and root-to-shoot translocation were enhanced. The results indicate that complexation of As(III) with PCs in Arabidopsis roots decreases its mobility for both efflux to the external medium and for root-to-shoot translocation. Enhancing PC synthesis in roots may be an effective strategy to reduce As translocation to the edible organs of food crops.Arsenic (As) contamination in the environment is caused by both geogenically and/or anthropogenically derived activities. This problem is the most serious in South and Southeast Asia where As-contaminated groundwater has been extracted for drinking and for irrigating rice (Oryza sativa) crops (Brammer and Ravenscroft, 2009). As contamination in soil can cause phytotoxicity and consequently yield losses (Panaullah et al., 2009) and elevated levels of As in rice grain that may pose a significant risk to human health (Meharg and Rahman, 2003; Zhu et al., 2008; Meharg et al., 2009). To develop mitigation strategies to reduce the transfer of As to the food chain requires a better understanding of the mechanisms of As uptake, translocation, and detoxification. It is known that As accumulation varies greatly among different plant species (e.g. Raab et al., 2007) and also among different genotypes within a species (e.g. Norton et al., 2009). Since root-to-shoot translocation is often the bottleneck for the accumulation of metal(loid)s in the shoots (Zhao and McGrath, 2009), understanding what controls As translocation within plants is important for designing strategies to decrease As concentrations in the edible parts of food crops.With the exception of As hyperaccumulating plants, translocation of As from roots to shoots is generally restricted in most plant species (for review, see Zhao et al., 2009). An explanation for this limited translocation is that arsenate [As(V)] is rapidly reduced to arsenite [As(III)] in roots, followed by complexation of As(III) with phytochelatins (PCs) and subsequent sequestration in root vacuoles (Dhankher et al., 2006; Raab et al., 2007; Zhao et al., 2009). The extent of As(III) complexation may therefore determine its mobility in roots. To test this hypothesis, we used the model plant Arabidopsis (Arabidopsis thaliana) mutants defective in glutathione (GSH) or PC synthesis, as well as manipulation of thiol synthesis in wild-type plants by the use of the specific inhibitor l-buthionine sulfoximine (BSO) and sulfur (S) deprivation. Both the PC-deficient mutant cad1-3 and the GSH-deficient mutant cad2-1 were isolated by their phenotype of cadmium (Cd) sensitivity (Howden et al., 1995a, 1995b). cad1-3 is a recessive loss-of-function mutant with a mutation in the PC synthase gene (AtPCS1; Ha et al., 1999) and is unable to synthesize PCs in response to Cd exposure (Howden et al., 1995b). cad2-1 has a deletion in the gene encoding the γ-glutamylcysteine synthetase, resulting in 60% to 85% lower levels of GSH compared with the wild type and little production of PCs in response to Cd exposure (Howden et al., 1995a; Cobbett et al., 1998).As(III) has a high affinity to thiol groups, and there is strong evidence that PCs play a constitutive role in the detoxification of As through complexation of As(III) in As nonhyperaccumulator plants. As strongly induces PC synthesis (Grill et al., 1987; Sneller et al., 1999; Schmöger et al., 2000). Both cad1-3 and cad2-1 are hypersensitive to As(V) (Ha et al., 1999; Li et al., 2006). Inhibition of GSH and PC synthesis by BSO results in As hypersensitivity in a number of plant species (Schmöger et al., 2000; Hartley-Whitaker et al., 2002; Schat et al., 2002). It has been shown that overexpression of PCS enhances As tolerance in transgenic plants, but interestingly not As accumulation (Li et al., 2004; Gasic and Korban, 2007). Furthermore, a range of intact As(III)-PC complexes has been identified in sunflower (Helianthus annuus) and Thunbergia alata plants after exposure to As(V) or As(III) (Raab et al., 2005; Bluemlein et al., 2008). In contrast, As hyperaccumulators, such as Pteris vittata, appear not to rely mainly on PC-dependent strategies for As detoxification, as very small proportions of As in roots and fronds are complexed with thiols (Webb et al., 2003; Zhao et al., 2003; Raab et al., 2004; Pickering et al., 2006). Lack of As(III)-PC complexation in P. vittata may be one of the important reasons for the highly efficient translocation of As from roots to fronds (Su et al., 2008; Zhao et al., 2009).While the role of PCs in As sensitivity is well established, how they influence As mobility in plants is not clear. Gong et al. (2003) showed that PCs may be transported from roots to shoots in a study involving root-specific expression of the wheat (Triticum aestivum) PCS gene (TaPCS1) in the Arabidopsis cad3-1 mutant. Furthermore, both root-specific and ectopic expression of TaPCS1 was found to enhance long-distance transport of Cd from roots to stems and rosette leaves, suggesting that PCs may be carriers of Cd in xylem transport. However, direct measurements of the xylem sap collected from As-exposed sunflower showed only traces of nonreactive oxidized PC₂ and oxidized glutathione (GSSG) with no evidence of As-PC complexation (Raab et al., 2005). Similarly, only trace levels of PCs were detected in the xylem sap from Cd-exposed Brassica napus (Mendoza-Cózatl et al., 2008). The role of PCs in the xylem mobility of As has not been investigated in detail. Interestingly, recent studies have shown that GSH, PCs, and other thiol peptides can be transported from shoots to roots via phloem (Chen et al., 2006; Li et al., 2006). High levels of PCs, GSH, and Cd were found in the phloem sap of B. napus, suggesting that thiol peptides may be carriers of Cd in the long-distance phloem transport (Mendoza-Cózatl et al., 2008).Here, we present evidence that decreasing As(III)-PC complexation in Arabidopsis roots led to greater As mobility, manifested by enhanced As(III) efflux to the external medium and enhanced As translocation from roots to shoots.     

SWEET17, a Facilitative Transporter,Mediates Fructose Transport across the Tonoplast of Arabidopsis Roots and Leaves

Fructose (Fru) is a major storage form of sugars found in vacuoles, yet the molecular regulation of vacuolar Fru transport is poorly studied. Although SWEET17 (for SUGARS WILL EVENTUALLY BE EXPORTED TRANSPORTERS17) has been characterized as a vacuolar Fru exporter in leaves, its expression in leaves is low. Here, RNA analysis and SWEET17-β-glucuronidase/-GREEN FLUORESCENT PROTEIN fusions expressed in Arabidopsis (Arabidopsis thaliana) reveal that SWEET17 is highly expressed in the cortex of roots and localizes to the tonoplast of root cells. Expression of SWEET17 in roots was inducible by Fru and darkness, treatments that activate accumulation and release of vacuolar Fru, respectively. Mutation and ectopic expression of SWEET17 led to increased and decreased root growth in the presence of Fru, respectively. Overexpression of SWEET17 specifically reduced the Fru content in leaves by 80% during cold stress. These results intimate that SWEET17 functions as a Fru-specific uniporter on the root tonoplast. Vacuoles overexpressing SWEET17 showed increased [¹⁴C]Fru uptake compared with the wild type. SWEET17-mediated Fru uptake was insensitive to ATP or treatment with NH₄Cl or carbonyl cyanide m-chlorophenyl hydrazone, indicating that SWEET17 functions as an energy-independent facilitative carrier. The Arabidopsis genome contains a close paralog of SWEET17 in clade IV, SWEET16. The predominant expression of SWEET16 in root vacuoles and reduced root growth of mutants under Fru excess indicate that SWEET16 also functions as a vacuolar transporter in roots. We propose that in addition to a role in leaves, SWEET17 plays a key role in facilitating bidirectional Fru transport across the tonoplast of roots in response to metabolic demand to maintain cytosolic Fru homeostasis.Sugars are main energy sources for generating ATP, major precursors to various storage carbohydrates as well as key signaling molecules important for normal growth in higher plants (Rolland et al., 2006). Depending on the metabolic demand, sugars are translocated over long distances or stored locally. SWEET (for SUGARS WILL EVENTUALLY BE EXPORTED TRANSPORTERS) and SUC/SUT (for Sucrose transporter/Sugar transporter)-type transporters are responsible for transfer of Suc from the phloem parenchyma into the sieve element companion cell complex for long-distance translocation (Riesmeier et al., 1992; Sauer, 2007; Kühn and Grof, 2010; Chen et al., 2012). Suc or hexoses derived from Suc hydrolysis in the cell wall are then taken up into sink cells by SUT (Braun and Slewinski, 2009) or monosaccharide transporters, such as sugar transporter1 (Sauer et al., 1990; Pego and Smeekens, 2000; Sherson et al., 2003). Alternatively, sugars are thought to move between cells via plasmodesmata (Voitsekhovskaja et al., 2006; Ayre, 2011). Major sugar storage pools within plant cells are soluble sugars stored in the vacuole, starch in plastids, and lipids in oil bodies.Vacuoles, which can account for approximately 90% of the cell volume (Winter et al., 1993), play central roles in temporary and long-term storage of soluble sugars (Martinoia et al., 2007; Etxeberria et al., 2012). Some agriculturally important crops such as sugar beet (Beta vulgaris; Leigh, 1984; Getz and Klein, 1995), citrus (Citrus spp.; Echeverria and Valich, 1988), sugarcane (Saccharum officinarum; Thom et al., 1982), and carrot (Daucus carota; Keller, 1988) can store considerable amounts (>10% of plant dry weight) of Suc, Glc, or Fru in vacuoles of the storage parenchyma. Due to a high capacity of vacuoles for storing sugars, vacuolar sugars can serve as an important carbohydrate source during energy starvation, e.g. after starch has been exhausted (Echeverria and Valich, 1988), as well as for the production of other compounds (e.g. osmoprotectants). Sugars are known to regulate photosynthesis; therefore, the release of sugars from vacuoles could be important for modulating photosynthesis (Kaiser and Heber, 1984). Moreover, vacuole-derived sugars are commercially used to produce biofuels, such as ethanol, from sugarcane. Knowledge of the key transporters involved in sugar exchange between the vacuole and cytoplasm is thus relevant in the context of bioenergy (Grennan and Gragg, 2009).To facilitate the exchange of sugars across the tonoplast, plant vacuoles are equipped with a multitude of transporters (Neuhaus, 2007; Etxeberria et al., 2012; Martinoia et al., 2012) comprising both facilitated diffusion and active transport systems of vacuolar sugars (Martinoia et al., 2000). Typically, Suc is actively imported into vacuoles by tonoplast monosaccharide transporter (AtTMT1/AtTMT2; Schulz et al., 2011) and exported by the SUT4 family (AtSUC4, OsSUT2; Eom et al., 2011; Payyavula et al., 2011; Schulz et al., 2011). Two H⁺-dependent sugar antiporters, the vacuolar Glc transporter (AtVGT1; Aluri and Büttner, 2007) and AtTMT1 (Wormit et al., 2006), mediate Glc uptake across the tonoplast to promote carbohydrate accumulation in Arabidopsis (Arabidopsis thaliana). The Early Responsive to Dehydration-Like6 protein has been shown to export vacuolar Glc into the cytosol (Poschet et al., 2011), likely via an energy-independent diffusion mechanism (Yamada et al., 2010). Defects in these vacuolar sugar transporters alter carbohydrate partitioning and allocation and inhibit plant growth and seed yield (Aluri and Büttner, 2007; Wingenter et al., 2010; Eom et al., 2011; Poschet et al., 2011).In contrast to numerous studies on vacuolar transport of Suc and Glc, limited efforts have been devoted to the molecular mechanism of vacuolar Fru transport even though Fru is predominantly located in vacuoles (Martinoia et al., 1987; Voitsekhovskaja et al., 2006; Tohge et al., 2011). Vacuolar Fru is important for the regulation of turgor pressure (Pontis, 1989), antioxidative defense (Bogdanović et al., 2008), and signal transduction during early seedling development (Cho and Yoo, 2011; Li et al., 2011). Thus, control of Fru transport across the tonoplast is thought to be important for plant growth and development. One vacuolar Glc transporter from the Arabidopsis monosaccharide transporter family, VGT1, has been reported to mediate low-affinity Fru uptake when expressed in yeast (Saccharomyces cerevisiae) vacuoles (Aluri and Büttner, 2007). Yet, the high vacuolar uptake activity to Fru intimates the existence of additional high-capacity Fru-specific vacuolar transporters (Thom et al., 1982). Recently, quantitative mapping of a quantitative trait locus for Fru content of leaves led to the identification of the Fru-specific vacuolar transporter SWEET17 (Chardon et al., 2013).SWEET17 belongs to the recently identified SWEET (PFAM:PF03083) super family, which contains 17 members in Arabidopsis and 21 in rice (Oryza sativa; Chen et al., 2010; Frommer et al., 2013; Xuan et al., 2013). Based on homology with 27% to 80% amino acid identity, plant SWEET proteins were grouped into four subclades (Chen et al., 2010). Analysis of GFP fusions indicated that most SWEET transporters are plasma membrane localized. Transport assays using radiotracers in Xenopus laevis oocytes and sugar nanosensors in mammalian cells showed that they function as largely pH-independent low-affinity uniporters with both uptake and efflux activity (Chen et al., 2010, 2012). In particular, clade I and II SWEETs transport monosaccharides and clade III SWEETs transport disaccharides, mainly Suc (Chen et al., 2010, 2012). Mutant phenotypes and developmental expression of several SWEET transporters support important roles in sugar translocation between organs. The clade III SWEETs, in particular SWEET11 and 12, mediate the key step of Suc efflux from phloem parenchyma cells for phloem translocation (Chen et al., 2012). Moreover, SWEETs are coopted by pathogens, likely to provide energy resources and carbon at the site of infection (Chen et al., 2010). Mutations of SWEET8/Ruptured pollen grain1 in Arabidopsis, and RNA inhibition of OsSWEET11 (also called Os8N3 or Xa13) in rice, and petunia (Petunia hybrida) NEC1 resulted in male sterility (Ge et al., 2001; Yang et al., 2006; Guan et al., 2008), possibly caused by inhibiting the Glc supply to developing pollen (Guan et al., 2008). Interestingly, two members, SWEET16 and SWEET17, of the family localize to the tonoplast (Chardon et al., 2013; Klemens et al., 2013). Allelic variation or mutations that affect SWEET17 expression caused Fru accumulation in Arabidopsis leaves, indicating that it plays a key role in exporting Fru from leaf vacuoles (Chardon et al., 2013). A more recent study demonstrated that SWEET16 also functions as a vacuolar sugar transporter (Klemens et al., 2013). Surprisingly, however, SWEET17 expression in mature leaves was comparatively low (Chardon et al., 2013), which leads us to ask whether SWEET17 could mainly function in other tissues under specific developmental or environmental conditions. Although Arabidopsis SWEET17 has been shown to transport Fru in a heterologous system where it accumulated in part at the plasma membrane (Chardon et al., 2013), the biochemical properties of SWEET17 were still elusive. SWEET16 and SWEET17 from Arabidopsis belong to the clade IV SWEETs. Whether clade IV proteins both transport vacuolar sugars in planta deserves further studies.Here, we used GUS/GFP fusions to reveal the root-dominant expression and vacuolar localization of the SWEET17 protein in vivo and its regulation by Fru levels. Phenotypes of mutants and overexpressors were consistent with a role of SWEET17 in bidirectional Fru transport across root vacuoles. The uniport feature of SWEET17 transport was further confirmed using isolated mesophyll vacuoles. Similarly, SWEET16 is also shown to function in vacuolar sugar transport in roots. Our work, performed in parallel to the two other studies (Chardon et al., 2013; Klemens et al., 2013), provides direct evidence for Fru uniport by SWEET17 and presents functional analyses to uncover important roles of these vacuolar transporters in maintaining intracellular Fru homeostasis in roots.