- © 2018 American Society of Plant Biologists. All rights reserved.
Abstract
Plants secrete a plethora of metabolites into the rhizosphere that allow them to obtain nutrients necessary for growth and modify microbial communities around the roots. Plants release considerable amounts of photosynthetically fixed carbon into the rhizosphere; hence, it is important to understand how carbon moves from the roots into the rhizosphere. Approaches used previously to address this question involved radioactive tracers, fluorescent probes, and biosensors to study sugar movement in the roots and into the rhizosphere. Although quite effective for studying sugar movement, it has been challenging to obtain data on spatial and temporal variability in sugar exudation using these techniques. In this study, we developed a gel-based enzyme-coupled colorimetric and fluorometric assay to image glucose (Glc) in vivo and used this assay to show that there is spatial variability in Glc release from plant roots. We found that the primary roots of maize (Zea mays) released more Glc from the base of the root than from the root tip and that the Glc release rate is reduced in response to water stress. These findings were confirmed independently by quantifying Glc release in well-watered and water-stressed maize primary roots using high-performance anion-exchange chromatography. Additionally, we demonstrated differential patterns of Glc exudation in different monocot and eudicot plant species. These findings and their implications on root-rhizosphere interactions are discussed.
The partitioning of carbohydrates from leaves to developing tissues is critical for plant growth and development. Therefore, it is important to understand how assimilated carbon, in the form of sugars, moves in plants and how this process is differentially regulated during plant development (Julius et al., 2017). Marschner (1986) reports that 30% to 60% of photosynthetic product is transported to roots, and a significant portion of this carbon (1%–30%) is deposited in the rhizosphere by mucilage, exudation, or decomposition of dead roots (Marschner, 1986; Jones et al., 2009). Lynch and Whipps (1990) report a rhizodeposition rate of 4% to 70% of photosynthetic products in various species, and Boyer et al. (2010) report a rhizodeposition rate of carbon at 1% to 11% in wheat (Tritium aestivum) and barley (Hordeum vulgare). These rhizodeposits, including root exudates, are the major source of carbon for the microbial populations in the soil and have been shown to impact the diversity and abundance of microbial populations in the rhizosphere (Broeckling et al., 2008; Badri and Vivanco, 2009; Kawasaki et al., 2016). To improve the understanding of root-microbe interactions, several studies have examined the composition of exudates in diverse plant species, including Arabidopsis (Arabidopsis thaliana; Chaparro et al., 2013), rice (Oryza sativa; Bacilio-Jimenez et al., 2003), maize (Zea mays; Carvalhais et al., 2011), wheat (Vancura, 1964; Warren, 2015), and Brachypodium distachyon (Kawasaki et al., 2016).
The predominant research approach used to examine the composition of root exudates is to grow the plants in a hydroponic solution (or sterile medium), collect the exudates released into the growth medium, and analyze the constituents using chromatography and/or mass spectrometry techniques (Vancura, 1964; Curl and Truelove, 1986; Gregory, 2006; Chaparro et al., 2013; Kawasaki et al., 2016). Analysis of root exudates from several plants revealed that amino acids, sugars, and organic ions are the major constituents of root exudates; among the sugars exuded are pentoses, hexoses, disaccharides, and trisaccharides. While these methods have been instrumental in providing information about the composition of root exudates, they cannot provide information about the spatial differences in root exudation processes. The differential release of simple sugars, such as Suc and Glc, as labile sources of energy can greatly influence the microbial abundance and diversity of microbes in the rhizosphere (Jaeger et al., 1999; Chaparro et al., 2013; Kawasaki et al., 2016). Hence, it is critical to understand the spatial and temporal exudation patterns of simple sugars.
Another approach used to study carbon movement into the rhizosphere is to monitor the distribution of radioactive tracers (Minchin and Thorpe, 2003; Babst et al., 2005; Pausch and Kuzyakov, 2011; Tran et al., 2017). Radioactive sugars (or sugar analogs) are fed through a cut surface of the plant, or alternatively, radioactive carbon dioxide is fed to be incorporated into the plant by photosynthesis, and the movement of the tracer in the plant and into the rhizosphere is tracked by radiation detectors (Babst et al., 2005; Keutgen et al., 2005; Kiser et al., 2008; Tran et al., 2017). This enables monitoring of the spatial and temporal movement of sugars within the plant and into the rhizosphere. However, these radioactivity techniques cannot provide information about the composition of the labeled exudates being released and cannot identify the metabolites being released (Jaeger et al., 1999; Pausch and Kuzyakov, 2011).
To address spatiotemporal variability in root exudation of specific metabolites, bacterial strains containing genetically encoded biosensors have been combined with specialized imaging technology to detect metabolite exudation into the rhizosphere (Jaeger et al., 1999; Darwent et al., 2003; Pini et al., 2017). Using this technique, Suc exudation was detected along the growing tip of Avena barbata (Jaeger et al., 1999) and pea (Pisum sativum) roots (Pini et al., 2017). Similar bacterial biosensors could be developed to detect Glc in the rhizosphere, especially since the genetic resources necessary for their development are available (Chaudhuri et al., 2008; Jones et al., 2013). Nevertheless, these transgenically expressed biosensors have a relatively narrow range of detection, and the transgenic bacterial colonies have to be maintained and cultivated uniformly in the growth medium (Jones et al., 2013; Pini et al., 2017) to effectively assess Glc exudation processes, which limits their routine use.
To circumvent the limitations of these methods and to detect in vivo Glc exudation, we developed a gel-based enzyme-coupled colorimetric and fluorometric assay and utilized it to characterize the Glc exudation patterns in well-watered and water-stressed maize primary roots. Glc exudation also was quantified using high-performance anion-exchange (HPAE) chromatography, confirming our findings. Additionally, we demonstrate the differential patterns in Glc exudation in different monocot and eudicot plant species. The method is straightforward and can be modified easily to image tissue-specific Glc localization in vivo with fresh, frozen, or lyophilized plant tissues. Owing to the simplicity of the assay, it can be adapted for use with any plant species or plant tissues and can be combined with the approaches described above to understand the movement of sugar in vivo and into the rhizosphere.
RESULTS
Development of a Gel-Based Glc Assay
The method for in vivo Glc localization was developed from the fluorometric assay described by McLaughlin and Boyer (2004). The assay utilizes the enzymes glucose oxidase (GO) and horseradish peroxidase (HRP) as well as the compound 10-acetyl 3,7-dihydroxyphenoxazine (Ampliflu Red). If Glc is encountered by the assay constituents, it is converted to gluconolactone and hydrogen peroxide by GO. The HRP then catalyzes the reaction between the released hydrogen peroxide and the colorless (nonfluorescent) Ampliflu Red, in a 1:1 stoichiometry, to form magenta-colored (and fluorescent) resorufin. In their original protocol, McLaughlin and Boyer (2004) incorporated the assay constituents in a gel made of 10% Difco gelatin and 5% polyethelene glycol to localize Glc in maize ovary tissues. However, in preliminary experiments in this study, it was found that the current composition of Difco gelatin and polyethelene glycol (from several commercial sources) contained substances that interfered with the assay. We tested alternative gelling materials, identified a low-gelling agarose that did not interfere with the assay, and utilized it to develop a similar gel-based enzyme-coupled colorimetric and fluorometric assay (for details, see “Materials and Methods”) to detect Glc released from plant roots.
Spatial Variability in Glc Exudation from Well-Watered Maize Primary Roots
Glc released from maize roots was assayed by placing the roots on a gel containing GO, HRP, and Ampliflu Red. Figure 1A shows magenta color around maize roots, indicating the Glc-dependent conversion of colorless Ampliflu Red into magenta-colored resorufin and demonstrating that Glc is released from maize roots. After 26 min of exudation, the primary roots showed a distinct spatial pattern of Glc release, with less Glc being released from the root tip and more Glc released from the rest of the root (Fig. 1A). No signal was detected from the roots placed on a control gel without GO (C-GO). Previous studies have shown that maize primary roots have endogenous peroxidase activity (Liszkay et al., 2004; Zhu et al., 2007); hence, it was not surprising that there was color development from the roots placed on a gel lacking HRP (C-HRP), although it was not as intense compared with the complete (C) medium. When the experiment was allowed to continue for longer than 30 min, the magenta color around the roots became diffuse over time and the spatial variability in Glc release was not as demarcated as in earlier time points (Supplemental Fig. S1; Supplemental Table S1). This is likely due to the diffusion of Glc (at a rate of 200 μm min−1, as shown in Supplemental Fig. S2) from areas of higher to lower concentration along the length of the root and into the gel around the roots. As positive and negative controls, air-dried filter paper strips containing 120 mm d-Glc were placed at the bottom of the C, C-HRP, and C-GO gels. As expected, color development indicating the presence of Glc was seen only in the strips placed on the C gel, demonstrating that the control (C-GO and C-HRP) gels were prepared appropriately.
Spatial variability in Glc exudation from maize primary roots. A, Primary roots of 2-d-old maize seedlings grown in well-watered vermiculite were imaged 26 min after being placed on gel containing Ampliflu Red reagent with GO and HRP (complete medium, denoted by C) or C medium without HRP (C-HRP) or GO (C-GO). The blue arrow indicates filter paper strips containing air-dried 120 mm d-Glc that were placed on the gel containing the C, C-HRP, or C-GO assay constituents as positive and negative controls. The development of magenta color indicates the production of resorufin from Ampliflu Red in a Glc-dependent manner. The experiment was repeated at least four times with similar results. B, Epifluorescence imaging of the differential Glc exudation from the maize primary root. Intact seedlings were placed on gel containing Glc assay constituents (GO, HRP, and Ampliflu Red reagent), and the time course of Glc exudation from the primary root tip and root base was monitored by epifluorescence microscopy. The fluorescence per image was calculated using ImageJ software, and the pixels per unit of area are shown. A detailed analysis of the quantification is shown in Supplemental Figure S3. Data are means ± se; n = 3 to 4 roots. The relative fluorescence scale is shown at the bottom of B. The experiment was performed twice with similar results, and representative images are shown.
To assess the spatial variability in Glc exudation more closely, primary roots of seedlings were placed on the C gel, and the root tip region and the root base region were imaged at various times to capture the Glc-dependent conversion of nonfluorescent Ampliflu Red to fluorescent resorufin. The dimensions of the root tip and root base regions were selected based on the data from Figure 1A. The epifluorescence images of the root tip and base regions clearly show incremental increases in fluorescence over time and that there is more fluorescence in the root base compared with the root tip region (Fig. 1B; Supplemental Fig. S3), indicating that more Glc was being exuded from the root base compared with the root tip region. The images of roots placed on control media (C-GO and C-HRP) showed quantifiably less fluorescence (Supplemental Figs. S3 and S4) than those of roots on C medium, validating the assay conditions. Glc also was released from roots grown on germination paper, indicating that the observations in Figure 1 were not due to the artifactual release of Glc from damaged roots (Supplemental Fig. S5).
The exudates from the root tip and base regions were analyzed by HPAE chromatography to quantify Glc levels and to independently confirm the differential Glc exudation from the maize primary roots. Glc was extracted and quantified from pooled samples of filter paper strips containing exudates from the root tip or base regions of nine seedlings (Fig. 2A). The absolute Glc concentrations in the root tip and base were quantified based on a standard curve (Supplemental Fig. S6) and were normalized to reflect Glc released from the control filter paper strips and for the Glc extraction efficiency (Fig. 2A). The root base region released an average of 36 ng Glc min−1 root−1, which is 5 times more Glc compared with the root tip region, which released an average of 7 ng Glc min−1 root−1 (Fig. 2B). Taken together, the results from the in vivo Glc visualization experiments (Fig. 1) and the Glc quantification experiments (Fig. 2) demonstrate that there is spatial variability in Glc exudation from different regions of maize primary roots grown under well-watered conditions.
Quantification of Glc exudation from the tip and base of maize primary roots. A, Two-day-old maize seedlings were placed on filter paper strips for 30 min to collect Glc exuded from the root tip and base. Glc from triplicate samples (with exudates pooled from nine seedlings each) was extracted and quantified by HPAE chromatography. The efficiency of Glc extraction was assessed by including filter paper strips with 450 ng of d-Glc (FP+G). Filter paper (FP) strips without added Glc or exudates were used as controls. Data are means ± se; n = 3 (from triplicate samples). Letters denote significant differences between samples as determined by one-way ANOVA and the Student-Newman-Keuls test (P ≤ 0.05). B, Average Glc exudation rate (ng min−1 during the first 30 min of incubation on the gel) from the tip and base of the well-watered maize primary root. Data are means ± se; n = 3 (from triplicate samples). The asterisk denotes a significant difference at P < 0.01 using Student’s t test. The experiment was performed twice with similar results.
Glc Exudation from the Water-Stressed Maize Primary Roots
In water deficit conditions, maize roots accumulate high levels of hexoses (Sharp et al., 1990) as well as enzymes and proteins that could be involved in Glc accumulation (Zhu et al., 2007; Spollen et al., 2008; Voothuluru et al., 2016). The increased concentration of hexoses in water-stressed roots (400 mm compared with 100 mm in well-watered root tissues) plays an important role in osmotic adjustment of the tissues and in withstanding the conditions of low water availability (Meyer and Boyer, 1981; Sharp et al., 1990). To assess if the increased Glc accumulation in water-stressed roots has an impact on Glc exudation, primary roots of maize seedlings grown under water deficit conditions (water potential of −1.4 MPa) were placed on a C gel for in vivo Glc localization. The results showed that Glc was exuded from water-stressed maize primary roots; however, there was no spatial variability in the pattern of Glc exudation in roots imaged after 26 min (Fig. 3A). Earlier examination revealed spatial variability after 10 min, with reduced Glc exuded from the tip compared with the base (Supplemental Fig. S7, A and B); however, this pattern was no longer discernible after 22 min (Supplemental Fig. S7, C–E), potentially due to Glc diffusion (Supplemental Fig. S2). A similar temporal and spatial pattern of Glc exudation was observed when the water-stressed primary roots were placed on a C gel containing mannitol that was isoosmotic to the water stress medium (Supplemental Fig. S2E). Quantification of Glc exudation from water-stressed roots confirmed the findings that the root tip and base regions released similar amounts of Glc (Fig. 3B). Overall, water-stressed roots exuded less Glc than well-watered roots; the basal region released 3 times less Glc compared with the basal region of well-watered roots. However, the root tip region released at least twice as much Glc compared with well-watered controls (Figs. 2 and 3B; Supplemental Fig. S2F).
Glc exudation from water-stressed maize primary roots. A, Primary roots of 2-d-old water-stressed maize seedlings grown in vermiculite at a water potential of −1.4 MPa were imaged 26 min after being placed on complete (C) or control (C-HRP or C-GO) gels. The experiment was performed three times with similar results. B, Average Glc exuded from the root tip and base of the water-stressed maize primary root per min (Glc exuded from the roots was collected, extracted, and quantified as described in “Materials and Methods”). Data are means ± se; n = 3 (from triplicate samples). The experiment was performed twice with similar results. No statistical differences were observed.
Glc Exudation Patterns Differ in Primary Roots of Different Plant Species
To test if the differential Glc exudation seen in maize primary roots is conserved across different plant species, the Glc exudation patterns from the primary roots of three eudicot and three monocot crop species were examined. After 2 d of growth in well-watered conditions, primary roots were placed on the C and control (C-HRP and C-GO) gels to detect spatial patterns in Glc exudation (Fig. 4). The soybean (Glycine max; Fig. 4A), common bean (Phaseolus vulgaris; Fig. 4B), and rice (Fig. 4F) Glc exudation patterns show that Glc is exuded uniformly along the length of the primary roots. Additionally, there was no difference in signal between roots on C and C-HRP (Supplemental Table S2), suggesting that Glc exudation is slow and/or little Glc is released but the activity of peroxidase is relatively strong in these species. In cotton (Gossypium hirsutum; Fig. 4C), more Glc was exuded from the root tip region, with negligible Glc being exuded from the base of the primary roots. The exudation patterns in sorghum (Sorghum bicolor; Fig. 4D) and wheat (Fig. 4E) were similar to that in maize (Fig. 1A), with less Glc being exuded from the root tip than the rest of the primary root. Although the Glc exudation pattern is similar in wheat and sorghum, when incubated for similar lengths of time, there was less intense color development in wheat (Supplemental Fig. S8; Supplemental Table S2), indicating that less Glc is exuded by wheat roots.
Glc exudation from the primary root of soybean (A), common bean (B), cotton (C), sorghum (D), wheat (E), and rice (F) seedlings. Roots of 2-d-old seedlings grown in well-watered vermiculite were imaged at various times after being placed on complete (C) or control (C-HRP or C-GO) gels. The blue arrow indicates filter paper strips containing air-dried 120 mm d-Glc that were placed on the gel containing the C, C-HRP, or C-GO assay constituents as positive and negative controls. The experiments were repeated two or more times with similar results.
DISCUSSION
Plant roots exude significant amounts of carbon in different forms, including soluble carbohydrates, such as Suc, Fru, and Glc (Marschner, 1986; Farrar et al., 2003). However, in most of the work performed to date, exudates were collected from plants grown either in hydroponic solutions or in sand flushed with nutrient solutions, and the constituent compounds, including Glc, were quantified using chromatography techniques, which obscured any spatial differences in exudation along the roots (Curl and Truelove, 1986; Gregory, 2006; Kawasaki et al., 2016). Based on the release of the Glc analog 3-O-methyl-d-glucopyranose, Jones and Darrah (1996) inferred that Glc could be released into the rhizosphere by passive diffusion across a concentration gradient between the root and the rhizosphere. Although these analyses imply Glc exudation, to date there is no direct evidence showing Glc release from plant roots. In this study, we developed a gel-based enzyme-coupled colorimetric assay to detect Glc in vivo and demonstrate that Glc is released from plant roots (Figs. 1 and 4). Furthermore, this work revealed that there is differential spatial variability in Glc exudation in different plant species. In maize, quantifiably less Glc was exuded by the root tip compared with the base (Fig. 1), even though the root tip has a higher concentration of Glc (Jones and Darrah, 1996). It can be hypothesized that the root tip, containing dividing and elongating cells, utilizes the Glc for various metabolic activities at a much higher rate than the basal region. Therefore, the spatial variability in well-watered roots of maize, sorghum, and wheat could simply be due to the differential utilization of Glc in the root tip compared with the base. However, this does not explain the spatial pattern seen in roots of other plants, particularly cotton, which shows increased activity in the apical region of the root (Fig. 4C; Supplemental Fig. S8C). Furthermore, water-stressed maize roots cumulatively released less Glc compared with well-watered roots, despite accumulating a higher concentration of Glc, which is necessary for osmotic adjustment (Sharp et al., 1990). Taken together, these results strongly indicate that Glc exudation from root tissues is a highly regulated process.
Although there is evidence that Glc is released into the rhizosphere (Jones and Darrah, 1996; Figs. 1 and 4), the path of Glc movement from the root into the rhizosphere is not known. The likely source of Glc is from the metabolism of phloem-delivered Suc, which is thought to be transported symplasmically from the phloem into the cortical tissues (radial outward movement) or into the meristematic and elongating tissues that are located apical to the terminus of the conducting phloem (apical downward movement; Giaquinta et al., 1983; Warmbrodt, 1987; Baker et al., 2016; Ross-Elliott et al., 2017). Suc metabolism could occur within the cytoplasm, with the Glc subsequently exported into the rhizosphere, or Suc could be first exported into the apoplast, where it could be hydrolyzed by cell wall invertases, leading to the release of Glc into the rhizosphere. Recently, Lunkova et al. (2017) showed that maize roots have substantial vacuolar invertase activity in the region of cell elongation (∼2–7 mm from the root tip), with the highest activity in the cortical and epidermal cells, and that the activity was reduced in the meristem and more basal region of the roots. These results suggest that Suc is hydrolyzed within the symplasm and that Glc is released into the rhizosphere. It is also possible that the primary roots exude larger carbohydrates into the rhizosphere, which are then cleaved by endogenous hydrolyzing enzymes with differential spatial activity to produce the patterns of Glc release observed in this study. However, even in this case, it is clear that the primary roots regulate the production of Glc in a spatially differential pattern in situ, with intriguing implications for root-rhizosphere interactions. Further work is necessary to resolve these possibilities and to further characterize the mechanisms of Glc release into the rhizosphere.
The release of Glc and other soluble sugars provides a major source of carbon for the microorganisms growing in the rhizosphere (Curl and Truelove, 1986; Walker et al., 2003). Several studies have identified variation in the microbial population and diversity along the length of plant roots, and it has been hypothesized that this is due to differential spatial and temporal exudation from roots (Marschner et al., 2001; Bais et al., 2006; Watt et al., 2006). The differential Glc-release patterns observed in this study (Figs. 1 and 4) provide evidence in support of this hypothesis. Watt et al. (2006) found that the apical region of wheat roots had less bacteria compared with the rest of the roots, and it is tempting to speculate that this is due in part to the reduced Glc release from this region (Fig. 4D). There may be some significance in the finding that the species that release more Glc overall (maize, sorghum, and wheat) had less Glc released from the root tip region (Figs. 1 and 4). Further research is needed to test whether the Glc-release patterns in different plants impact the rhizosphere microbial population. Research also is warranted to understand the impacts of stress-induced changes in Glc release on root-rhizosphere interactions. It has been shown that roots with reduced growth rates accumulate more pathogenic bacteria in the apical region, which may be detrimental to root growth and plant health (Watt et al., 2003). Given that water-stressed maize roots release relatively more Glc in the root tip region compared with well-watered roots (Figs. 2 and 3), it would be important to know if the increased Glc release coupled with the reduced growth rate of water-stressed roots impacts the root association with pathogenic bacteria.
The simple and robust method for Glc visualization described herein could be used to assess in vivo tissue-specific Glc localization in other plant tissues. This method is flexible and highly customizable, and Glc could be detected in fresh, frozen, or lyophilized tissues (McLaughlin and Boyer, 2004). Additionally, it could be modified further to image Suc (by adding invertase to the assay constituents) or potentially other polysaccharides. However, users need to take several precautions while performing the assay protocols and interpreting the results. Ampliflu Red can be photooxidized by light, even in darkroom conditions and more easily under intense lighting used in conjunction with epifluorescence microscopy (Zhao et al., 2012). This photooxidation is enhanced by the presence of peroxidases, present either in the assay constituents or in the experimental tissues; hence, the use of C and control gels for all experiments is necessary to account for any background color/fluorescence development. It is also important to consider the rate of Glc release from the experimental tissues. The conversion of colorless/nonfluorescent Ampliflu Red to pink-colored/fluorescent resorufin by hydrogen peroxide occurs at a 1:1 stoichiometry only when the initial concentration of Ampliflu Red is higher than that of hydrogen peroxide (Zhou et al., 1997). Excess hydrogen peroxide can further oxidize the pink/fluorescent resorufin to a brown-colorless/nonfluorescent compound, although the rate is 30-fold slower than the Ampliflu Red conversion to resorufin (Zhou et al., 1997). Given this information, if tissues release large amounts of Glc, colorimetric or fluorescence measurements need to be conducted for shorter durations. On the other hand, if the rate of Glc release from tissues is slower, the background color/fluorescence development on the control gels will become a more significant factor in interpreting the results. For example, soybean and bean primary roots showed pink color development on C and control gels (particularly C-HRP). The low level of Glc being released from soybean and bean roots, coupled with the activity of endogenous root peroxidase, may be sufficient to convert Ampliflu Red to resorufin on the C-HRP gel. The color development of the roots placed on C-GO gels may indicate the production of hydrogen peroxide or other oxidizing compounds from the roots due to prolonged incubation of the roots on the C-GO gels. Nevertheless, we show that the assay can be optimized and can be utilized to observe biologically informative spatial distribution patterns of Glc release from plant roots. In the long term, integrating this new Glc imaging technology with plant physiology and developmental biology will enable us to better understand carbohydrate partitioning during plant development under favorable and unfavorable conditions.
MATERIALS AND METHODS
Plant Materials
Plant materials were bean (Phaseolus vulgaris) ‘Blue Lake 274’, cotton (Gossypium hirsutum) ‘AU90810’, maize (Zea mays) ‘B73’, rice (Oryza sativa) var Swarna, sorghum (Sorghum bicolor) ‘Macia’, soybean (Glycine max) ‘Williams 82’, and wheat (Triticum aestivum) var Coker 9553.
Plant Growth Conditions
Germinated seeds were transplanted into well-watered (0.03 MPa) or low-water-potential (−1.4 MPa) vermiculite hydrated with 1 mm CaSO4 as described previously (Spollen et al., 2000) and grown at 29°C and near-saturation humidity in the dark until harvest (2 d after transplanting). The low water potentials were measured at the beginning of each experiment by isopiestic thermocouple psychrometry (Boyer and Knipling, 1965).
Visualization of Glc Exudation
Glc exudation from primary roots was visualized by a gel-based enzyme-coupled colorimetric and fluorometric assay modified from McLaughlin and Boyer (2004). Two glass plates (20 cm × 20 cm × 0.2 cm) separated by an adhesive foam sheet (Fiber-Craft) with cutouts to create molds of different dimensions (12 cm × 2 cm × 0.2 cm for well-watered plants and 9 cm × 2 cm × 0.2 cm for water-stressed plants) were held together with binder clips. A liquid gel mixture was prepared with 0.05 m phosphate buffer (Sigma P7994) at pH 7.4, 1.5% (w/v) low-gelling agarose (Sigma A9414), and 1 mm CaSO4 (Sigma 255548), which was heated to 65°C or above to dissolve the agarose. The solution was cooled to 37°C (by placing in a water bath) and the assay constituents GO (Sigma GO543) at a final concentration of 1.8 units mL−1, HRP (Sigma P8125) at a final concentration of 1.5 units mL−1, and Ampliflu Red (Sigma 90101) at a final concentration of 200 µm were added. Immediately, the gel mixture was poured into the custom-made molds and cooled to form a gel at room temperature. After the gel was set (approximately 20 min), one of the glass plates was removed gently and the gel (along with the casting mold on the other glass plate) was placed in a large petri dish (25 cm × 25 cm × 2.5 cm) and stored at 4°C until just before use. Since Ampliflu Red can oxidize in light, the gel was prepared using a green safelight (Saab et al., 1990) and stored protected from light.
Two-day-old seedlings were gently removed from the vermiculite, and the intact primary roots were placed on the gel and covered with a glass plate (0.5 mm thick) to ensure contact between the roots and the gel. Care was taken to remove any vermiculite loosely bound to the roots without damaging them. The gel was allowed to warm to room temperature, and the reaction progress was captured on a light box using a digital camera (Supplemental Fig. S9; Olympus C5050). In maize and sorghum, the experiment was terminated 30 min after the roots were placed on the gel, since longer incubation resulted in further oxidation of resorufin to a colorless (nonfluorescent) compound, decreasing the signal (McLaughlin and Boyer, 2004). For other species, the experiment was carried out for longer durations (40–60 min). At the end of the experiment, air-dried filter paper strips containing 120 mm d-Glc were placed on the gel as controls to validate the assay conditions. In some experiments, if the Glc strips were placed on the C or C-HRP gel for longer than 5 min, the strips turned brown due to further oxidation of resorufin (Supplemental Figs. S2C and S8G). The experiments were technically and biologically repeated at least two times with similar results.
The images obtained from the digital camera were analyzed further using ImageJ software (Abramoff et al., 2004), and the magenta color indicating Glc exudation was quantified as follows. The hue of the images was adjusted to filter through only the magenta pixels, and the saturation was adjusted to remove background color in all the images. Subsequently, the area around the root (within each gel lane) was selected and the number of magenta pixels per unit of area was quantified. Data are means ± se; n = 3 to 6 roots. For statistical significance, data were analyzed by ANOVA using SigmaPlot (Systat Software); means, when significantly different, were separated using the Student-Newman-Keuls test (at P ≤ 0.05).
In well-watered maize primary roots, the progress of the assay and the development of fluorescent resorufin (excitation at 563 nm and emission at 587 nm; McLaughlin and Boyer, 2004) was captured by an epifluorescence microscope (Leica M205FA) with an attached color digital camera (Leica DFC7000T). The epifluorescence images were converted to grayscale, and the fluorescence intensity was pseudocolored using the Royal lookup table from ImageJ (Abramoff et al., 2004). The fluorescence intensity per image (pixels per unit of area) was quantified on a scale of 0 to 255 using the analyze tool from ImageJ. The background fluorescence due to the photooxidation of Ampliflu Red (Zhao et al., 2012), particularly in C and control (C-GO) gels, interfered with the quantification. Therefore, to account for the background fluorescence, the pixel intensities from 0 to 5 of the grayscale (the lowest detection levels of fluorescence) were not included in the quantification of the images (the quantification, when conducted without this correction, still showed significant differences in fluorescence between roots placed on C and control [C-HRP and C-GO] gels but erroneously did not show a difference among the roots on C-HRP and C-GO gels). Data are means ± se; n = 3 to 4 roots. For statistical significance, data were analyzed by ANOVA using SigmaPlot (Systat Software); means, when significantly different, were separated using the Student-Newman-Keuls test (P ≤ 0.05).
Quantification of Glc Exudation
The intact primary roots of three maize seedlings were placed on Whatman filter paper (No. 1) strips (2 cm × 2 cm) moistened with 0.05 m phosphate buffer at pH 7.4 to collect exudates from the root tip and base. To ensure proper contact between the primary roots and the filter paper, the roots were covered with a glass plate (0.5 mm thickness). After 30 min, filter paper strips were collected and stored in a −80°C freezer until further use.
The dimensions of the root tip and base of well-watered and water-stressed seedlings were determined by the visualization of Glc exudation (Figs. 1 and 3). In well-watered seedlings, the root tip was contained in the 0- to 1.5-cm region from the root apex and the root base was in the 5- to 7-cm region from the root apex. In water-stressed seedlings, the root tip was in the 0- to 1-cm region from the root apex and the root base was in the 2- to 4-cm region from the root apex. We considered expressing the Glc exudation on a per gram root dry weight basis. However, since water-stressed roots have thicker walls (Wu et al., 1994), they are heavier than well-watered tissues and would erroneously show lower levels of Glc exudation (than the levels reported in this work). Therefore, we calculated Glc exudation based on the length of the roots (or root regions) to show differential Glc exudation from well-watered and water-stressed roots.
Exudates from root tips and bases were collected onto filter paper strips. Glc from the triplicate samples (filter paper strips containing exudates from nine seedlings each) was extracted with 2 mL of methanol:chloroform:water (12:15:3) solution (Leach and Braun, 2016). After extraction, the aqueous solution (∼2 mL) was concentrated to 500 µL using a Savant SpeedVac (Thermo-Fisher Scientific SC110) and Glc in the samples was quantified by HPAE chromatography (Thermo-Fisher Scientific ICS-5000; Leach et al., 2017). The concentration of Glc in the samples was determined using a standard curve obtained by measuring known Glc standards. Control filter paper strips (without exudates or added Glc) released measurable amounts of Glc, which was quantified and subtracted from the absolute Glc quantification. The efficacy of Glc extraction was 65% to 87% in different replicate experiments, and it was assessed by including samples with 450 ng of d-Glc (from three filter paper strips with 150 ng each of d-Glc). The controls (filter paper strips without or with added d-Glc) and samples with exudates from root tips and bases were analyzed in triplicate. Data are means ± se of triplicate samples. Letters denote significant differences between samples as determined by one-way ANOVA and the Student-Newman-Keuls test (P ≤ 0.05). The amounts of Glc exuded from the root tip and base were normalized for Glc recovery. Statistical significance was determined at P < 0.05 using Student’s t test in Excel (Microsoft).
Supplemental Data
The following supplemental materials are available.
Supplemental Figure S1. Time course of Glc exudation patterns from the primary root of maize seedlings grown in well-watered conditions.
Supplemental Figure S2. Glc diffusion rate on the complete gel.
Supplemental Figure S3. Quantification of the fluorescence from images of maize primary roots after 20 min of being placed on complete or negative control medium.
Supplemental Figure S4. Time course of epifluorescence imaging of the differential Glc exudation from maize primary roots on negative control medium.
Supplemental Figure S5. Epifluorescence imaging of the differential Glc exudation from the 2-d-old maize primary roots grown on germination paper.
Supplemental Figure S6. Representative standard curve used for Glc quantification.
Supplemental Figure S7. Time course of Glc exudation from water-stressed maize primary roots.
Supplemental Figure S8. Images of Glc exudation patterns from the primary root of soybean, bean, cotton, wheat, and rice seedlings.
Supplemental Figure S9. Image of the experimental setup.
Supplemental Table S1. Quantification of the Glc exudation from maize roots.
Supplemental Table S2. Quantification of the Glc exudation from roots of different species.
Acknowledgments
We thank two anonymous reviewers for comments that significantly improved the article, Ben Julius and Dr. Kristen Leach for their technical assistance with HPAE chromatography measurements, and Drs. Gary Stacey and Debbie Finke for providing the soybean and wheat seeds.
Footnotes
↵1 This research was supported in part by the National Science Foundation Plant Genome Research Program (grant nos. IOS-1025976 and IOS-1444448 to D.M.B.).
↵2 Current address: Department of Ecology and Evolutionary Biology, University of Tennessee, Knoxville, TN 37996.
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- Received May 17, 2018.
- Accepted September 11, 2018.
- Published September 20, 2018.
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