Research ArticlePlant biology

Two glutamate- and pH-regulated Ca2+ channels are required for systemic wound signaling in Arabidopsis

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Science Signaling  14 Jul 2020:
Vol. 13, Issue 640, eaba1453
DOI: 10.1126/scisignal.aba1453

Glutamate-dependent systemic signaling in plants

Upon wounding, plants generate systemic Ca2+ waves and electrical signals that propagate from the wound site to distal tissues (see the Focus by Fichman et al.). The glutamate receptor–like proteins GLR3.3 and GLR3.6 are required for leaf-to-leaf systemic wound signals in Arabidopsis thaliana. Shao et al. found that wounding or the application of glutamate induced root-to-shoot Ca2+ and electrical signaling in Arabidopsis, which required GLR3.3, GLR3.6, and inhibition of the proton pump AHA1. In cultured mammalian cells, GLR3.3 and GLR3.6 functioned as pH-sensitive, glutamate-gated Ca2+ channels. These findings suggest that wounding induces both the leakage of glutamate from the phloem into the apoplastic space and an increase in the apoplastic pH, leading to the activation of GLRs and the generation of systemic Ca2+ waves and electrical signals.

Abstract

Plants defend against herbivores and nematodes by rapidly sending signals from the wounded sites to the whole plant. We investigated how plants generate and transduce these rapidly moving, long-distance signals referred to as systemic wound signals. We developed a system for measuring systemic responses to root wounding in Arabidopsis thaliana. We found that root wounding or the application of glutamate to wounded roots was sufficient to trigger root-to-shoot Ca2+ waves and slow wave potentials (SWPs). Both of these systemic signals were inhibited by either disruption of both GLR3.3 and GLR3.6, which encode glutamate receptor–like proteins (GLRs), or constitutive activation of the P-type H+-ATPase AHA1. We further showed that GLR3.3 and GLR3.6 displayed Ca2+-permeable channel activities gated by both glutamate and extracellular pH. Together, these results support the hypothesis that wounding inhibits P-type H+-ATPase activity, leading to apoplastic alkalization. This, together with glutamate released from damaged phloem, activates GLRs, resulting in depolarization of membranes in the form of SWPs and the generation of cytosolic Ca2+ increases to propagate systemic wound signaling.

INTRODUCTION

Plants are constantly under mechanical stress caused by weather conditions such as wind and by biotic factors, including pathogens and herbivorous animals. Such factors can lead to tissue damage or wounding in both underground and aboveground organs. Particularly impactful to crop production is pest infestation by leaf-chewing insects and root nematodes. In response to wounding, plants initiate sophisticated mechanisms to repair the wounded tissues and defend against pests. The plant hormone jasmonic acid (JA) is a critical regulator of wound responses and is synthesized upon tissue damage (16). Studies have revealed molecular pathways for JA signal transduction that begins with the intracellular receptor COI1 (710), followed by the degradation of JAZ family repressors to activate JA-responsive genes (1114). JA-based responses occur immediately not only at the wound site but also in unwounded distal organs linked to the wound site through the vasculature (15). JA biosynthesis takes place within 1 min in leaves distal to the wounded leaf (1618), implicating a long-distance rapid wound signal moving from the wounded leaf to distal leaves. Several candidates for this rapid signal have been identified, including slow wave potentials (SWPs), glutamate, Ca2+, and reactive oxygen species (ROS) (1922).

SWPs are electrical signals generated upon wounding by the depolarization and repolarization of the cell membrane in some cell populations in plants. Although the precise origin of SWPs is still under debate, SWPs remain a viable candidate for the systemic wound signal (1923). In particular, several studies have linked SWPs to systemically induced JA biosynthesis and wound responses through genetic analysis of SWP-defective mutants and artificial manipulation of the leaf potential by current injection (24). The Arabidopsis thaliana genetic mutants defective in SWP include those with disrupted glutamate-receptor-like (GLR) genes, such as GLR3.3 and GLR3.6. The glr3.3glr3.6 double mutant is specifically compromised in the capacity to propagate the wound SWP signal to the distal leaves, which is accompanied by impaired JA biosynthesis in undamaged leaves, supporting the hypothesis that electrical signals can be a rapid systemic wound signal (24, 25). SWP signals in most studies are recorded by tissue surface electrodes, making it difficult to interpret the contribution of different cell types to these SWP signals. To overcome this problem, the electrical penetration graph (EPG) has been used to detect changes in the membrane potential of Arabidopsis sieve elements during plant-herbivore interactions (26). EPG recordings have shown that the glr3.3glr3.6 double mutant fails to propagate wound-induced SWPs, consistent with the results from conventional surface electrode recording and confirming the importance of GLR3.3 and GLR3.6 in SWP propagation (27).

Work by Kumari et al. (28) suggests that the P-type adenosine triphosphatase (ATPase), Arabidopsis H+-ATPase 1 (AHA1), plays a role in modulating SWPs in response to wounding, suggesting that SWPs may be a proxy of plasma membrane potential. Because depolarization is critical for systemic wound signals, AHA1 activity should inhibit depolarization through hyperpolarization or repolarization of the membrane. As a result, the repolarization phase is extended in the aha1 mutant and shortened in the gain-of-function AHA1 allele ost2-2D (28). These changes have altered the SWPs with a consequence in systemic wound response. Thus, the exact pattern of the SWPs is functionally relevant to wound signal propagation.

The systemic cytosolic Ca2+ wave has been shown to be another early wound response event (2934). Plant GLRs are putative Ca2+-permeable channels and receptors for glutamate, and direct testing of the involvement of glutamate in wound responses has identified a connection between glutamate and wound-responsive Ca2+ waves (34). The application of 50 to 100 mM glutamate at a wound site triggers a long-distance Ca2+ wave typical of that induced by mechanical or insect-inflicted wounding events, indicating that glutamate may serve as a trigger of the long distance rapid wound signal. The plant phloem contains 10 to 100 mM glutamate, and the concentration of glutamate in the apoplast, the space between the cell membrane and cell wall, may rise to 50 mM at damage sites (34). Thus, it is proposed that the glutamate released from the phloem may serve as a damage-associated molecular pattern (DAMP), and GLR3.3 and GLR3.6 could be activated in response, leading to the increase of cytosolic Ca2+ concentration ([Ca2+]cyt) (34). Furthermore, wound-induced cytosolic Ca2+ increase, together with calmodulin, initiates disassociation of the JAV1-JAZ8-WRKY51 complex, leading to the activation of JA biosynthetic genes (35). Although these studies support the importance of local and systemic Ca2+ signals in the wound response, the puzzling question concerns the underlying mechanism for the propagation of this [Ca2+]cyt increase from wound sites to distal leaves. Specifically, despite evidence supporting the essential function of GLR3.3 and GLR3.6 proteins for the propagation of the systemic wound signal, it remains unknown whether they are indeed glutamate-sensitive Ca2+-permeable channels.

ROS have also been proposed to act as a long-distance rapid wound signal (36). Using a transgenic plant expressing luciferase driven by the ROS-responsive AtZAT12 promoter (37, 38), researchers have identified a wound-induced systemic ROS signal traveling at a rate of 8.4 cm/min, which depends on the respiratory burst oxidase homolog D (RbohD) in Arabidopsis (36). A newly developed method (39) can directly measure ROS produced upon wounding in planta using the oxidation of fluorescence probes such as 2′,7′-dichlorofluorescin diacetate. In addition, heat or high light stress can also generate systemic electrical signals that may be linked to ROS, because the amplitude of SWPs is reduced in the rbohD mutant (40).

Although parasitic nematodes do not wound plants the same way as chewing insects, nematode attacks also induce a systemic defense response that involves the production of JA (41). In tomato, the root-knot nematode induces the systemic transmission of electrical and ROS signals from attacked roots to the leaves, leading to an increased accumulation of JAs in the leaves (42). Tomato mutants lacking GLR3.5 or RBOH1 are compromised in nematode-induced electrical and ROS signals in the leaves, suggesting that ROS and SWPs may serve as nematode-induced root-to-shoot systemic signals in tomato plants (42).

Although several candidates for systemic wound signals have been identified, little is known about the relationship of these candidates, especially concerning SWPs and systemic Ca2+ waves. In particular, GLR3.3, GLR3.6, and AHA1 are involved in both the SWPs and Ca2+ waves, but little is known about the mechanism underlying the function of GLRs and AHA1 in the wound response. Here, we established a root-to-shoot long distance signaling system in Arabidopsis and showed that glutamate activated both SWPs and systemic Ca2+ waves that depended on the actions of GLR3.3, GLR3.6, and AHA1. The GLRs were functionally connected to AHA1 in systemic signaling because they were glutamate-activated and proton-inhibited Ca2+-permeable channels required for generating both the membrane depolarization and Ca2+ increases responsible for electrical and Ca2+ waves in plants.

RESULTS

Wounding and glutamate trigger systemic root-to-shoot [Ca2+]cyt waves

As previously reported (34), systemic [Ca2+]cyt increase is a hallmark of the leaf-to-leaf long distance signaling in response to wounding. To establish a model for studying the root-to-shoot systemic wound response, we used transgenic plants carrying a single copy of the Ca2+-responsive fluorescent protein GCaMP6s (43) to monitor [Ca2+]cyt changes in live plants while wounding the roots. When mechanical wounding was applied to roots by severing the main root at a site 2 cm away from the root-shoot junction, a strong vasculature-based [Ca2+]cyt increase was observed in the rosette leaves (Fig. 1, A and B, and movie S1). Also, the [Ca2+]cyt increase was seen in all leaves, unlike the leaf-to-leaf system in which the [Ca2+]cyt changes preferentially propagated to leaves at positions n ± 3 and n ± 5 relative to the wounded leaf (34). Such differences likely resulted from the vascular connection of main roots to all the above-ground organs. Video recording of the time course of the [Ca2+]cyt rise indicated a rapid and uniform change in Ca2+ in all leaves within 1 min of wounding (movie S1), similar to the time course of the leaf-to-leaf systemic [Ca2+]cyt wave (34). We then tested whether glutamate at the wound site in the root also elicited systemic [Ca2+]cyt changes, as shown in the leaf-to-leaf signaling system (34). Fifteen minutes after wounding, when the wound-responsive Ca2+ waves returned to the resting level, application of a 10-μl drop of 100 mM glutamate to the wound site induced another strong and rapid systemic [Ca2+]cyt increase in all the leaves of the rosette, with similar kinetics as that caused by wounding (Fig. 1, C and D, and movie S2).

Fig. 1 Wounding and glutamate trigger root-to-shoot transmission of [Ca2+]cyt waves.

(A) Imaging of WT (Col-0) plants expressing the genetically encoded intracellular Ca2+ indicator GCaMP6s at the indicated time points after cutting the main root. (B) Kinetics of [Ca2+]cyt increases in leaves 4, 5, and 6 quantified along the time course after wounding. n = 10 plants. Boxplot shows max [Ca2+]cyt in the time frame. (C) Ca2+ imaging of WT plants expressing GCaMP6s at the indicated time points after adding 100 mM glutamate at the wound site. Scale bar, 1 cm. (D) Kinetics of [Ca2+]cyt increases in leaves 4, 5, and 6 quantified along the time course after application of glutamate. n = 10 plants. Boxplot shows max [Ca2+]cyt in the time frame. (E) Comparison of the time courses of [Ca2+]cyt increase in leaf 6 when roots were cut at either 2 or 5 cm away from the root-shoot junction. n = 10 plants (2 cm) and 13 plants (5 cm). Boxplot shows max [Ca2+]cyt in the time frame. (F and G) [Ca2+]cyt increases recorded in leaf 6 of three groups of plants after wounding (F) and in the same leaves again after application of 100 mM (group A), 50 mM (group B), or 20 mM (group C) glutamate to the wound sites (G). n = 8 plants (group A), 10 plants (group B), and 10 plants (group C). Error bars, means ± SEM. Boxplots show the minimum, 25th percentile, median, 75th percentile, and maximum of the max [Ca2+]cyt data points in the time frame. Asterisks indicate statistically significant differences compared with the WT by two-tailed Student’s t test. **P < 0.01.

We explored the effect of different wound sites along the roots on the systemic response and found that the response weakened with distance from the root-shoot junction. The maximum ΔF/F0 induced by a cut 2 cm away from the junction was significantly larger than that caused by a cut at 5 cm (Fig. 1E). We also found that wounding a lateral root triggered a similar but weaker systemic [Ca2+]cyt increase as compared to wounding the main root (fig. S1A). To test the effect of different glutamate concentrations on systemic Ca2+ waves, we divided plants into three groups (A, B, and C) and then wounded the plant roots 2 cm away from the root-shoot junction. The three groups showed similar [Ca2+]cyt increases in response to wounding (Fig. 1F). After 15 min, we added 100, 50, and 20 mM glutamate to the wound site of the group A, B, and C plants, respectively, and found that 50 and 100 mM glutamate triggered a significantly larger [Ca2+]cyt increase than did 20 mM glutamate (Fig. 1G). These results demonstrated that wound-induced root-to-shoot systemic [Ca2+]cyt rise was dependent on both the concentration of apoplastic glutamate and the distance of wound sites from the root-shoot junction.

Glutamate, like wounding, induces a root-to-shoot systemic electrical wave

Wounding initiates leaf SWPs that propagate from the wounded leaf to undamaged distal leaves at a speed of ~5 cm/min, which induces systemic JA biosynthesis and qualifies SWPs as a systemic wound signal (24). We performed leaf-to-leaf SWP recordings as described in earlier studies (24) and obtained results similar to those published (fig. S2). We then tested whether SWPs were also produced during the root-to-shoot systemic wound response. When the main root was wounded, the surface potential (SWP) measured at a rosette leaf showed a quick hyperpolarization, followed by prolonged depolarization and repolarization phases (Fig. 2, A to D). Wounding lateral roots also triggered a similar systemic SWP (fig. S1B). The SWPs measured for the root-to-shoot wound response appeared similar to SWPs in the leaf-to-leaf response, consistent with the hypothesis that the root-to-shoot wound response also involves SWPs as a long-distance signal. As one of the plant DAMPs, glutamate triggers a systemic [Ca2+]cyt rise that, like SWPs, also causes JA synthesis in distal leaves (34). To address the relationship between SWPs and glutamate, we used the root-to-shoot wound response system developed here to test the possibility that glutamate released at (or added to) the wound site triggers electrical signals represented by SWPs. Fifteen minutes after wounding, we added glutamate to the wound site and observed an SWP on a rosette leaf (Fig. 2, A to C and E), reminiscent of the one generated by wounding. We also observed glutamate-triggered leaf-to-leaf SWPs that were similar to those induced by wounding (fig. S2, A to C). This finding made a critical link between glutamate and electrical signals (SWPs), indicating that glutamate release at the wound site was sufficient for the generation of both SWPs and systemic Ca2+ waves.

Fig. 2 Wounding and glutamate trigger a root-to-shoot systemic electrical signal.

(A) Typical wound- and glutamate-induced SWPs recorded at leaf 6. The time points at which the root was wounded (W) and 100 mM glutamate was added to the wound site (Glu) are marked with arrowheads. Four parameters, including hyperpolarization amplitude (Amphyp), hyperpolarization duration (Durhyp), depolarization amplitude (Ampdep), and depolarization duration (Durdep) were used to compare SWPs. (B and C) Comparison of amplitude (B) and duration (C) of wound- and glutamate-induced SWPs. n = 78 plants for wound-induced SWPs, and 69 plants for glutamate-induced SWPs. Boxplots show the minimum, 25th percentile, median, 75th percentile, and maximum of the data points. (D) Two typical wound-induced SWPs recorded at leaf 6. (E) Two typical glutamate-induced SWPs recorded at leaf 6.

To quantitatively analyze root-to-shoot SWPs, we divided the SWP into the hyperpolarization phase and depolarization phase (Fig. 2A) and collected data on the amplitude and duration for both phases. We found that both the amplitude and duration for hyperpolarization and depolarization were similar between the wound- and glutamate-induced SWPs (Fig. 2, B and C), indicating that glutamate-induced SWPs mimic the wound-induced SWPs.

To further compare leaf-to-leaf and root-to-shoot SWPs, we analyzed in detail the depolarization phases of these SWPs. In previous studies, the depolarization phase of wound-induced leaf-to-leaf SWPs appears smooth with a single sustained phase (24, 27). We found SWPs with monophasic patterns similar to those previously described and, in addition, more complex leaf-to-leaf SWPs consisting of an initial fast depolarization followed by a slower and sustained depolarization (fig. S2). We found that 56% (44 of 78) of wound-induced root-to-shoot SWPs consisted of an initial fast depolarization, followed by a slower depolarization, and 44% (34 of 78) were simpler, showing a single sustained phase (Fig. 2D). For glutamate-induced root-to-shoot SWPs, the depolarization phase of 61% (42 of 69) of plants was monophasic, whereas 39% (27 of 69) of plants showed an initial fast and then slower depolarization phase (Fig. 2E). We concluded that root-to-shoot SWPs and leaf-to-leaf SWPs in our study and those described in previous studies were comparable during the depolarization phase (24, 27). Together, these results suggest that glutamate released at the wound site may be a trigger of SWPs in the systemic wound response in both root-to-shoot and leaf-to-leaf models.

GLR3.3 and GLR3.6 are required for root-to-shoot systemic wound signaling

In the leaf-to-leaf wound response model, GLR3.3 and GLR3.6 are necessary for the propagation of wound-induced SWPs and systemic [Ca2+]cyt rise (24, 25, 34). Because GLRs are putative glutamate receptors in plants, the involvement of GLR3.3 and GLR3.6 in the systemic wound response further supports the importance of glutamate as the initial signal at the wound site. Our results showed that glutamate at the wound site induced both [Ca2+]cyt increases and SWPs, consistent with the idea of glutamate being a trigger for systemic wound signaling. We thus tested the possibility that GLR3.3 and GLR3.6 may also be essential for the root-to-shoot wound response.

When roots were wounded, the amplitude and duration of the depolarization phase of the systemic SWPs were significantly reduced in the glr3.3glr3.6 double mutant (Fig. 3, A to F), whereas the duration of the hyperpolarization phase was slightly longer than wild type (Col-0) (Fig. 3D). We then added glutamate to the wound sites of plant roots and found that, compared with Col-0, the SWPs in a glr3.3glr3.6 mutant rosette leaf were also significantly impaired in both the hyperpolarization and depolarization phases (Fig. 3, A, B, G to J). These results showed that GLR3.3 and GLR3.6 played essential roles in the wound- and glutamate-induced systemic SWPs in root-to-shoot long-distance signaling.

Fig. 3 GLR3.3 and GLR3.6 participate in root-to-shoot wound signaling.

(A and B) Typical recordings of wound-induced (W) and glutamate-induced (Glu) SWPs in leaf 6 in Col-0 (A) and glr3.3glr3.6 (B) plants. The time points at which the root was wounded and glutamate was added to the wound site are marked with arrowheads. (C to F) Wound-induced SWPs in leaf 6 of Col-0 and glr3.3glr3.6 plants are described by amplitude (C) and duration (D) of hyperpolarization and by amplitude (E) and duration (F) of depolarization. (G to J) Glutamate-induced SWPs in leaf 6 of Col-0 and glr3.3glr3.6 plants are described by amplitude (G) and duration (H) of hyperpolarization and amplitude (I) and duration (J) of depolarization. The WT (Col-0) data from leaf 6 are the same as those shown in Fig. 2A; n = 78 plants for wound-induced SWPs and n = 69 plants for glutamate-induced SWPs. For experiments in glr3.3glr3.6, n = 71 plants. Boxplots show the minimum, 25th percentile, median, 75th percentile, and maximum of the data points. Asterisks indicate statistically significant differences compared with the WT by two-tailed Student’s t test. ****P < 0.0001.

In parallel assays, we found that wound- and glutamate-induced root-to-shoot systemic [Ca2+]cyt increases were nearly abolished in the glr3.3glr3.6 double mutant (fig. S3, A to D, and movies S3 and S4). Furthermore, we measured the systemic JA response by checking the mRNA abundance for marker genes JAZ7, JAZ10, and OPR3 in rosette leaves and found that wound-induced, JA-dependent transcript abundances were much lower in the glr3.3glr3.6 double mutant compared to Col-0 (fig. S3, E to G). These results confirmed the essential roles of GLR3.3 and GLR3.6 during the systemic wound response not only in the leaf-to-leaf system but also in the root-to-shoot system.

P-type H+-ATPase activity plays a role in the root-to-shoot systemic wound response

SWPs in the root-to-shoot model involved both hyperpolarization and depolarization phases (Fig. 2). Although it is not clear exactly how SWPs correspond to plasma membrane potentials, previous studies have suggested that P-type H+-ATPases may be involved in producing the SWPs (4449). We examined the importance of P-type H+-ATPase in root-to-shoot systemic signaling by two approaches. First, we studied the effect of the fungus-derived, H+-ATPase–activating compound fusicoccin (FC) on SWPs induced by wounding or glutamate application in the root-to-shoot systemic response (fig. S4). The rosette leaves of plants divided into three groups (A, B, and C) and wounded on the roots 2 cm away from the root-shoot junction showed similar SWPs in response to wounding (fig. S4, A to C). Fifteen minutes after wounding, we added 50 mM glutamate, 2.5 μM FC, or 2.5 μM FC combined with 50 mM glutamate to the wound sites of the group A, B, and C plants, respectively. Whereas glutamate treatment elicited a typical SWP and FC treatment alone had no effect (fig. S4, B and C, right panel), the combination of glutamate plus FC slightly increased the amplitude of the hyperpolarization phase but, notably, eliminated both the amplitude and duration of the depolarization phase (fig. S4, B and C, right panel), demonstrating that the activation of H+-ATPases by FC blocked the depolarization phase of glutamate-induced SWPs.

Second, we used a genetic approach to test the effect of ost2-2D, a gain-of-function allele of AHA1, on the wound- and glutamate-induced SWPs (Fig. 4, A and B). When the roots of ost2-2D were wounded, the SWPs of the rosette leaf showed higher hyperpolarization amplitude and normal duration as compared with Col-0 (Fig. 4, C and D). In the depolarization phase, ost2-2D showed lower amplitude and shorter duration than did Col-0 (Fig. 4, E and F). We then added glutamate to the wound sites and found that ost2-2D showed higher amplitude and normal duration in the hyperpolarization phase as compared with Col-0 (Fig. 4, G and H). In the depolarization phase, the duration of SWPs on the ost2-2D rosette leaf was significantly reduced as compared to those of Col-0 (Fig. 4, I and J), showing again that constitutively active AHA1 inhibited the depolarization phase of both wound- and glutamate-induced SWPs. Together, the results from both pharmacological and genetic approaches indicated that the inhibition of P-type H+-ATPase was required for plants to generate and/or propagate SWPs in the root-to-shoot model, which is consistent with published findings on the leaf-to-leaf wound response (28).

Fig. 4 Constitutive activation of P-type H+-ATPase inhibits wound- and glutamate-induced SWPs.

(A and B) Typical recording of wound- and glutamate-induced SWPs in leaf 6 in Col-0 (A) and ost2-2D (B) plants. The time points when the root was wounded (W) and glutamate was added (Glu) to the wound site are marked with arrowheads. (C to F) Wound-induced SWPs in leaf 6 of Col-0 and ost2-2D plants as described by amplitude (C) and duration (D) of hyperpolarization and amplitude (E) and duration (F) of depolarization. (G to J) Glutamate-induced SWPs in leaf 6 of Col-0 and ost2-2D plants as presented by amplitude (G) and duration (H) of hyperpolarization and amplitude (I) and duration (J) of depolarization. n = 27 plants for Col-0 and n = 39 plants for ost2-2D. Boxplots show the minimum, 25th percentile, median, 75th percentile, and maximum of the data points. Asterisks indicate statistically significant differences compared with the WT by two-tailed Student’s t test. **P < 0.01, ***P < 0.001, and ****P < 0.0001.

The Ca2+-permeable channel activities of GLR3.3 and GLR3.6 are gated by extracellular glutamate and pH

Data presented here and in earlier studies support the essential role of GLR3.3 and GLR3.6 in the systemic wound response for both leaf-to-leaf and root-to-shoot models (24, 25, 34). These GLRs appear to couple the SWPs and Ca2+ waves in some way so that these systemic signals become interdependent. One hypothesis is that GLR3.3 and GLR3.6 may function as glutamate-activated Ca2+-permeable channels, stimulating Ca2+ influx that alters plasma membrane potentials. However, it remains to be tested whether GLR3.3 and GLR3.6 are indeed glutamate-gated Ca2+ channels. Our data herein (Fig. 4 and fig. S4) and a previous report (28) both identified P-type H+-ATPase as a critical component in the systemic wound response, possibly as a result of its contribution to the plasma membrane potential and thus to SWPs. We explored the interplay of the P-type H+-ATPase and GLR3.3 and GLR3.6 by examining the channel properties of Arabidopsis GLR3.3 and GLR3.6, which is key to further understanding systemic wound signaling.

The Arabidopsis GLR family consists of 20 members that are implicated in various functions [reviewed in (50)]. The channel properties, however, are largely unexplored because only a few of the 20 members have been studied using patch-clamp approaches. For example, AtGLR3.4 expressed in human embryonic kidney (HEK) cells shows Ca2+-permeable channel activity that is enhanced by the presence of 1 to 2 mM asparagine, serine, or glycine (51). AtGLR1.4 appears to function as a nonselective cation channel when expressed in frog oocytes and is sensitive to 1 mM of a broad range of amino acids (52). AtGLR3.2 and AtGLR3.3 can conduct Ca2+-permeable current in the absence of any ligand when coexpressed in monkey kidney COS-7 cells with AtCORNICHON1 and AtCORNICHON4, Arabidopsis homologs of sorting proteins that control the trafficking of ionotropic glutamate receptors (iGluRs) in animals (53). Also, the biochemically reconstituted GLR3.3 ligand-binding domain (LBD) appears to show a preference for l-glutamate and the sulfur-containing amino acids l-cysteine and l-methionine (54). Together, the studies so far indicate that some plant GLRs can be gated by amino acids at low millimolar or submillimolar concentrations, and some are constitutively open without any ligands. However, Toyota et al. (34) suggest that glutamate, but not other amino acids, is specifically effective in eliciting Ca2+ waves in systemic wound response that requires putative glutamate receptors GLR3.3 and GLR3.6. Furthermore, the concentration of glutamate must be 50 mM or higher to have a significant effect (34). These findings are not consistent with the known channel properties of GLRs reported so far, whether GLR3.3 and GLR3.6 indeed serve as the glutamate receptors responsible for the systemic wound response. We decided to bridge this critical gap between channel activities and in planta function of the GLRs in the context of systemic wound response by two approaches to delineating GLR3.3 and GLR3.6 functional properties.

First, we examined the activity of GLR3.3 and GLR3.6 in mediating Ca2+ influx using single-cell Ca2+ imaging experiments in HEK293T cells coexpressing the Ca2+ indicator GCaMP6s and plant GLRs. However, we initially failed to observe any activity in plant GLR3.3 and GLR3.6 using this assay. Further investigation of the literature on animal iGluRs indicated that a subset of these receptors, specifically the N-methyl-d-aspartate receptors (NMDARs), are not only gated by their ligands but are also highly sensitive to protons. The open probability of NMDARs is reduced with increasing proton concentration (or lowering the pH) (55), and a structural study described the mechanism of this pH sensitivity (56). If Arabidopsis GLR3.3 and GLR3.6 are also inhibited by low pH, then that would explain why we did not observe any activity when we used typical acidic pH conditions to apply glutamate and other amino acids to the HEK cells in the attempt to activate GLR3.3 and GLR3.6. We found that, under higher pH (7.5 and 8.5), the external application of 100 mM glutamate induced a large [Ca2+]cyt increase in the HEK293T cells transiently coexpressing GLR3.3 or GLR3.6 and GCaMP6s (Fig. 5A and movies S5 and S6). Under lower pH (5.5 and 6.5), however, glutamate-induced [Ca2+]cyt increases were negligible (Fig. 5A). These data indicated that GLR3.3 and GLR3.6, like animal NMDARs, were proton-sensitive.

Fig. 5 Ca2+ imaging of HEK293T cells expressing GLR3.3 or GLR3.6.

(A) Single-cell imaging of [Ca2+]cyt increases in HEK293T cells expressing GLR3.3 or GLR3.6, as induced by 100 mM glutamate at the indicated pH values. (B) [Ca2+]cyt increases induced by the indicated concentrations of glutamate at pH 8.5 in HEK293T cells expressing GLR3.3 or GLR3.6. (C) [Ca2+]cyt changes induced by 100 μM glutamate in cells expressing GLR3.3, GLR3.6, or Grik2. (D) [Ca2+]cyt in HEK293T cells expressing GLR3.6 and stimulated with the indicated amino acids at pH 8.5. Amino acids were added at 100 mM, except for tyrosine and tryptophan, which were used at lower concentrations because of lower solubility. For relative max (FF0)/F0, the mock control was set to 1. For all experiments, n = 3 independent experiments, with ~60 cells imaged in each experiment. Error bars depict means ± SEM. Student’s t test. *P < 0.05, **P < 0.01.

We then applied different concentrations of glutamate (10, 25, 50, and 100 mM) to the HEK293T cells coexpressing GCaMP6s plus either GLR3.3 or GLR3.6 under pH 8.5 conditions, and found that only 50 and 100 mM glutamate induced significant [Ca2+]cyt increases (Fig. 5B). This was a surprising finding given that animal iGluRs can be activated by micromolar concentrations of glutamate (57). To rule out the possibility that the marked difference in ligand sensitivity between plant and animal GLRs resulted from problems in our Ca2+ imaging system, we used a rat iGLuR, glutamate receptor ionotropic, kainate 2 (Grik2), as a control. We found that 100 μM glutamate triggered a large [Ca2+]cyt increase in HEK293T cells expressing Grik2 and GCaMP6s under pH 7.5, whereas no response was observed for Arabidopsis GLR3.3 or GLR3.6 under the same conditions (Fig. 5C).

Animal iGluRs are gated by multiple amino acids, including glutamate, glycine, serine, and alanine (57), and it has been shown that several plant GLRs are also gated by multiple amino acids (51, 52, 58). We thus tested the effect of each of the 20 amino acids on the activity of GLR3.6 and found that only glutamate (at 100 mM), but not other amino acids, triggered significant [Ca2+]cyt increases in HEK293T cells coexpressing GLR3.6 and GCaMP6s (Fig. 5D and movies S5, S6, and S7). This result is consistent with the report showing that glutamate is the only amino acid that specifically elicits a systemic Ca2+ wave in plants (34).

To further investigate the channel properties of GLR3.3 and GLR3.6, we conducted patch-clamp experiments on HEK292T cells expressing the individual GLRs. Using the standard bath solution containing 10 mM Ca2+ and 100 mM glutamate, we observed cation-carried inward currents in HEK293T cells expressing GLR3.3 or GLR3.6 under pH 7.5 (Fig. 6, A to D). Because several previous reports showed that some plant GLRs can be activated by lower amounts (<1 mM) of amino acids (51, 52, 58), we included rat Grik2 as a control. We consistently recorded large inward currents in HEK293T cells expressing Grik2 in the standard bath solution containing 10 mM Ca2+ and 100 μM glutamate, whereas no inward current was recorded for GLR3.3 or GLR3.6 beyond that observed in the mock control (Fig. 6, E and F). We thus concluded that GLR3.3 and GLR3.6 were not activated by low millimolar or submillimolar concentrations of glutamate, consistent with the earlier results using single-cell Ca2+ imaging (Fig. 5, A to C). This conclusion supports the finding that only high millimolar concentrations of glutamate (above 50 mM) can elicit a systemic Ca2+ wave in plants (34). Moreover, we found that the inward currents mediated by GLR3.3 or GLR3.6 were significantly larger when bath pH was adjusted to 7.5 and 8.5, as compared to lower pH (5.5 and 6.5) (Fig. 6, A to D), confirming that GLR3.3 and GLR3.6 were inhibited by the extracellular protons.

Fig. 6 Patch-clamp recordings of the channel activities of GLR3.3 and GLR3.6.

(A) Typical whole-cell recordings of inward currents in HEK293T cells expressing GLR3.3, as induced by 100 mM glutamate at the indicated pH values. (B) Average current–voltage curves for GLR3.3 at different pH values, n = 4 to 10 cells per condition. (C) Typical whole-cell recordings of inward currents in HEK293T cells expressing GLR3.6, as induced by 100 mM glutamate at the indicated pH values. (D) Average current–voltage curves for GLR3.6 at different pH values. n = 4 to 10 cells per condition. (E) Typical whole-cell recordings of inward currents in cells expressing GLR3.3, GLR3.6, or Grik2 as induced by 100 μM glutamate at pH 7.5. (F) Average current–voltage curves for GLR3.3, GLR3.6, and Grik2. n = 4 to 6 cells per condition. Error bars depict means ± SEM. Student’s t test. *P < 0.05, **P < 0.01.

Because the standard bath solution contained not only Ca2+ but also Na+ and K+, we characterized the permeability of GLR3.3 and GLR3.6 to these monovalent and divalent cations using patch-clamping. We added 140 mM Na+, K+, or Cs+, and 112 mM Ca2+ or Ba2+, respectively, to the bath solution containing 100 mM glutamate with pH adjusted to pH 8.5 and performed patch-clamping in the whole-cell configuration (figs. S5, A to J, and S6, A to J). We found that GLR3.3 or GLR3.6 conducted inward currents with both monovalent and divalent cations as carriers with a preference for Ca2+ and Ba2+. The permeability sequence of GLR3.3 was Ba2+ > Ca2+ > Na+ = K+ > Cs+, and for GLR3.6, the sequence was Ba2+ > Ca2+ > K+ > Na+ > Cs+, showing that GLR3.3 and GLR3.6 are nonselective cation channels that prefer Ba2+ and Ca2+ (table S1).

Animal iGluRs can assemble as homo- and heterotetrameric channels, and functional channels are formed exclusively by the assembly of subunits of the same receptor class (57). Arabidopsis GLR subunits interact with multiple other GLRs in a yeast mating-based split ubiquitin system (59), indicating that Arabidopsis GLRs might also form heteromeric complexes. However, a fusion of GLR3.3 to the reporter β-glucuronidase (GLR3.3-GUS) is expressed in the phloem region, whereas GLR3.6-GUS is expressed in the xylem contact cells (25), suggesting that GLR3.3 and GLR3.6 are unlikely to form heterotetrameric channels in plants because they are present in distinct cell populations.

DISCUSSION

The root-to-shoot systemic wound response is a robust model for studying plant long-distance signaling

Most studies on systemic wound signaling have focused on the leaf-to-leaf model in Arabidopsis (24, 25, 28, 34). Although it is established that the amounts of 12-oxo-phytodienoic acid, a JA precursor, increase and JA-responsive genes are induced in the shoot when the root is wounded (60, 61), root-to-shoot systemic wound signaling remains largely unexplored. During the preparation of this study, it was reported that electrical signals and ROS are involved in the systemic root-to-shoot response when tomato plants are damaged by the root-knot nematode, leading to an increased accumulation of JAs in the leaves (42). The authors describe the importance of tomato GLR3.5 (a homolog of Arabidopsis GLR3.3 and GLR3.6) and RBOH1 in the nematode-induced root-to-shoot systemic signaling process (42). Although it is difficult to compare this study using nematode in tomato with earlier studies using mechanical wounding in Arabidopsis, the results suggest that root-to-shoot signaling and leaf-to-leaf signaling may share some common components.

Here, we established a root-to-shoot systemic wound signaling system in Arabidopsis and compared it with the leaf-to-leaf model in the same species. In addition to showing the participation of the same signals, including glutamate, SWPs, and Ca2+ waves, in the root-to-shoot systemic wound response, we also made progress in understanding the functional relationship of these signals in the context of the channel properties of plant GLRs.

The glutamate-dependent Ca2+ channel activity of GLR3.3 and GLR3.6 couples systemic Ca2+ and electrical waves in the wound response

Although systemic signaling through nerve action potentials has been well established in animals, electrical signaling mechanisms in plants had been controversial until a pioneering study (24) identified the genes encoding GLR3.3 and GLR3.6 as essential genetic determinants for propagation of electrical signals from a wounded leaf to distal leaves, leading to a systemic response to wounding. An important advance has linked wound- and glutamate-induced Ca2+ waves to these same GLRs (25, 34), further emphasizing the central role of GLRs in long-distance signaling during the plant wound response. What are plant GLRs, and why are they required for systemic wound signaling? At the center of this question lie the functional properties of plant GLRs.

Animal iGluRs are nonselective cation channels, and some of them, especially those of the NMDAR family, are Ca2+-permeable (57). Although plant GLRs contain similar domains to those present in the NMDARs, including the N-terminal domain, the receptor or LBD, the pore domain, and the cytosolic C-terminal domain, it is difficult to predict channel activity and ionic selectivity of plant GLRs by comparing with animal NMDARs because of the low conservation of the pore sequence, and particularly the residues dictating the Ca2+ permeability of NMDAR channels (50, 62). Previous studies support the hypothesis that plant GLRs, like animal NMDARs, function as Ca2+-permeable channels that may be activated by amino acids (5153, 63). Therefore, it makes sense that GLRs are important in wound- and glutamate-induced long-distance Ca2+ signaling. However, there exists a major gap between studies using electrophysiological approaches and those measuring systemic wound responses in plants. The data collected by patch-clamping animal cells expressing plant GLRs indicate that at least two plant GLRs, including AtGLR3.4 and AtGLR1.4, are Ca2+-permeable channels gated by 1 mM amino acids including Asn, Ser, Gly, Met, Trp, Phe, Leu, Tyr, and Thr (51, 52, 58). In contrast, 50 mM or higher concentrations of glutamate, but not other amino acids, specifically induce GLR3.3- and GLR3.6-dependent Ca2+ waves in the leaf-to-leaf wound response (34). The concentrations of glutamate required for systemic signaling are a couple of magnitudes higher than those shown to induce channel activity in the electrophysiological studies, although different members of the GLR family have been analyzed in different studies. To connect the channel activity and the role of these GLRs in the systemic wound response, our work here has addressed both the electrophysiological properties of GLR3.3 and GLR3.6 and their function in systemic wound responses.

We confirmed that high concentrations of glutamate (50 mM or higher) were essential to elicit a robust systemic Ca2+ wave and electrical signal in the root-to-shoot system. Further, we showed that GLR3.3 and GLR3.6 expressed in HEK293T cells are activated by the same concentration of glutamate (50 to 100 mM), bridging the gap between channel properties of these GLRs and their function in systemic wound response. Our finding that Arabidopsis GLR3.3 and GLR3.6 were activated by 50 to 100 mM glutamate (Figs. 5 and 6, figs. S5 and S6, and table S1) is in stark contrast to the ligand gating of animal iGluRs that are activated by a micromolar range of external glutamate (57). This finding, however, is consistent with the special physiology of plants in that apoplastic concentrations of glutamate are normally in the low millimolar range, whereas 20 to 100 mM glutamate is found in the phloem sap (34, 64, 65), making it feasible for plants to use a glutamate gradient as a DAMP. Under resting conditions, apoplastic glutamate should not be sufficient to activate GLRs, consistent with our finding that GLR3.3 and GLR3.6 were not active in the presence of low millimolar glutamate. However, upon wounding, the damaged phloem cells would release glutamate to the apoplastic space, where it reaches 50 to 100 mM and activates GLRs in the plasma membranes of adjacent cells required for systemic wound signaling. This low ligand sensitivity may be unique to plant GLRs, and the structural basis for such a property deserves investigation.

The pH-sensitive activity of GLR3.3 and GLR3.6 links the function of P-type H+-ATPase to the systemic wound response

Protons profoundly inhibit animal NMDAR activity, and alkali pH enhances the open probability of the channels (55). Here, we found that Arabidopsis GLR3.3 and GLR3.6 channels are also highly sensitive to apoplastic pH, with alkali pH as an essential condition for channel opening. This finding provides further insight into the function of P-type H+-ATPase in the systemic root-to-shoot wound response documented herein and in the leaf-to-leaf long-distance signaling previously described (28). We propose that upon wounding, inhibition of P-type H+-ATPase activity causes apoplastic alkalinization, which cooperates with glutamate released into the apoplastic space to activate GLRs, leading to the influx of Ca2+ and other cations through GLRs, thereby depolarizing the membrane. As a result, constant activation of P-type H+-ATPase through a gain-of-function mutation in ost2-2D mutant or by FC treatment might block the alkalization of apoplastic space (Fig. 4 and fig. S4) and therefore the activation of GLRs (Figs. 5 and 6), inhibiting membrane depolarization and Ca2+ influx. This does not conflict with the AHA1 function in the report in which AHA1 may be activated after the Ca2+ influx to repolarize the membrane (28). In other words, inhibition and activation of AHA1 may act as a critical cycle for changes in membrane potential and apoplastic pH that hold the key for Ca2+ entry and signal propagation through GLRs.

To support the hypothesis that AHA1 is involved in the wound systemic response, the measurement of apoplastic pH changes upon wounding will be important. However, because of technical limitations, we did not have any success in the rapid and accurate recording of apoplastic pH changes in response to wounding. One previous report (66) shows that wounding a leaf induces cytosolic acidification, suggesting that proton pumps are inhibited upon wounding, which supports our hypothesis.

The finding that GLR3.3 and GLR3.6 were only active when pH was higher than 6.5 raises another key question: Could plant apoplastic pH reach the threshold high enough to activate GLR3.3 and GLR3.6? In one study (67), the authors fused pHluorin, a well-established pH sensor, to a transmembrane domain so that the fusion protein would be anchored in the plasma membrane (PM) with the pH-sensing moiety facing the apoplast. Using this PM-anchored pHluorin, membrane-associated apoplastic pH (around 6.5) is found to be much more alkaline than that of the overall cell wall (67). Thus, inhibition of P-type H+-ATPase would increase the pH further to a range higher than the resting level of 6.5, which would be sufficient for the activation of GLR3.3 and GLR3.6. Future studies should directly analyze the apoplastic pH changes in response to wounding and glutamate application in the root-to-shoot and leaf-to-leaf models.

In addition, the data presented herein and results from other studies (28) may also provide a possible mechanism for how plants terminate the SWPs and Ca2+ waves: When the [Ca2+]cyt waves pass by, the cells would re-activate the P-type H+-ATPase that would pump protons out, repolarize the cells, and acidify the apoplast, which would inhibit GLR channel activity and terminate the systemic SWP and Ca2+ signal. In summary, our results reported here have uncovered critical regulatory properties of GLR3.3 and GLR3.6 that connect glutamate, H+-ATPase, SWP, and Ca2+ waves in plant systemic wound signaling. Some key questions remain to be answered regarding the mechanisms underlying both the initiation and propagation of the signals. What triggers the inhibition of H+-ATPase immediately after wounding to increase apoplastic pH to enable GLR activation by glutamate? How does a signal propagate along the vascular tissues? Future studies will be directed toward answering these questions on plant long-distance signaling that, to some extent, mimics nerve transmission of signals in animals in that glutamate, electrical, and Ca2+ signals are intertwined into a complex network.

MATERIALS AND METHODS

Plant material and growth conditions

Seeds were sterilized with 10% (v/v) bleach and sown on agar plates containing 1/2 Murashige and Skoog (MS) medium [1/2 MS, 0.8% (w/v) Phyto agar, and 1% (w/v) sucrose, pH adjusted to 5.8 with KOH]. Plates were incubated at 4°C for 3 days for stratification and then transferred to 22°C growth room with a 12-hour light/12-hour dark cycle (100 μmol m−2 s−1) for 6 days. Seedlings were then transferred onto new plates (2 plants per plate) and grown for additional 15 days before electrical recording or calcium imaging upon wounding. The ost2-2D seeds were kindly provided by Jian-Min Zhou (68). The glr3.3 (SALK_099757C) and glr3.6 (SALK_091801C) were ordered from Arabidopsis Biological Resource Center (ABRC) and crossed to obtain the glr3.3glr3.6 double mutant. The glr3.3glr3.6 mutant was crossed with a transgenic line containing UBQ10 promoter-driven GCaMP6s and further brought to homozygosity with both GCaMP6s and glr3.3glr3.6 background.

Surface potential recordings and current injection

The leaf surface potential recording method was modified from the published method (24). An Axopatch 200B patch-clamp amplifier (Axon Instruments) with a Digidata 1550 digitizer (Axon Instruments), was applied to record the surface potential. The pClamp 10.7 software (Axon Instruments) was used for data acquisition and analysis. The amplifier was turned to I-Clamp Normal mode, and the sampling rate is 1000 Hz. The recording silver electrode was connected to the leaf surface using 10 μl of 0.5% (w/v) agar containing 10 mM KCl, and the reference silver electrode was inserted into the agar medium.

Vector construction, cell culture, and transfection

The cDNA encoding GCaMP6s was amplified from HBT-GCaMP6-HA43 (43), and the cDNAs encoding GLR3.3 and GLR3.6 were kind gifts from José A. Feijó (53) and Edward E. Farmer (24). The GLR3.3 and GLR3.6 and GCaMP6s were cloned into a single vector, pBudCE4.1 (Invitrogen), for coexpression in HEK293T cells.

HEK293T cells were cultured in Dulbecco’s modified Eagle’s medium supplemented with 10% fetal bovine serum in a 5% CO2 incubator at 37°C in a moist atmosphere. HEK293T cells were transfected using the Lipofectamine 3000 Transfection Reagent Kit (Invitrogen). Plasmids for transfection were extracted from Escherichia coli (DH5α) using QIAGEN Plasmid Mini Kit (Qiagen), and 2 μg of plasmids in total was added into each well of six-well plates (Nunc). HEK293T cells showing bright green fluorescence were used for patch-clamp experiments and Ca2+ imaging experiments 48 hours after transfection.

Whole-cell patch-clamp recording

The whole-cell patch-clamp experiments were performed using an Axopatch 200B patch-clamp setup (Axon Instruments) with a Digidata 1550 digitizer (Axon Instruments) as previously reported (69, 70). pClamp10.7 software (Axon Instruments) was used for data acquisition and analysis.

The standard pipette solution for all experiments contained 140 mM CsCl, 5 mM EGTA, and 10 mM Hepes (pH 7.5; adjusted with CsOH) as previously described (71). The standard bath solution contained 100 mM Na-glutamate, 40 mM NaCl, 5 mM KCl, 2 mM MgCl2, 10 mM CaCl2, 10 mM Hepes, and 10 mM glucose (pH 7.5) (adjusted with NaOH). Different pH values were indicated in the figures.

For monovalent-cation substitution experiments, the bath solution was changed as follows: (i) 100 mM Na-glutamate, 40 mM NaCl, 10 mM glucose, and 10 mM Hepes (adjusted to pH 8.5 with NaOH); (ii) 100 mM K-glutamate, 40 mM KCl, 10 mM glucose, and 10 mM Hepes (adjusted to pH 8.5 with KOH); and (iii) 100 mM Cs-glutamate, 40 mM CsCl, 10 mM glucose, and 10 mM Hepes (adjusted to pH 8.5 with CsOH).

For divalent-cation substitution experiments, the bath solution was changed as follows: (i) 50 mM Ca-(glutamate)2, 62 mM CaCl2, 10 mM glucose, and 10 mM Hepes [adjusted to pH 8.5 with Ca(OH)2]; and (ii) 50 mM Ba-(glutamate)2, 62 mM BaCl2, 10 mM glucose, and 10 mM Hepes [adjusted to pH 8.5 with Ba(OH)2]. A ramp voltage protocol of 2-s duration from −160 to +30 mV (holding potential 0 mV) was applied 1 min after accessing a whole-cell configuration, and currents were recorded every 20 s for 40 times in total for each cell. The 40 current traces were used for statistical analysis for average current-voltage curves. Permeability ratios for monovalent and divalent cations against Cs+ were calculated as previously described (71).

Single-cell calcium imaging in HEK293T cells

HEK293T cells expressing GCaMP6s were monitored by a Zeiss Axio Observer Z1 Inverted Microscope using a 20× objective as previously reported (72). The interval of data acquisition was 5 s. The software iVision 4.5 (BioVision Technologies) was used for data acquisition and analysis. The standard solution for Ca2+ imaging contained 120 mM NaCl, 3 mM KCl, 1 mM MgCl2, 1.2 mM NaHCO3, 10 mM glucose, and 10 mM Hepes (pH 7.5). About 60 s after initiation of imaging procedure, the bath was perfused using a peristaltic pump with a solution containing 100 mM Na-glutamate, 20 mM NaCl, 3 mM KCl, 10 mM CaCl2, 1 mM MgCl2, 1.2 mM NaHCO3, 10 mM glucose, and 10 mM Hepes (pH 7.5) to elicit Ca2+ entry through active channels. Different pH values were indicated in the figures.

Whole-plant calcium imaging

Transgenic Arabidopsis plants stably expressing GCaMP6s were imaged with a Zeiss Lumar v12 epifluorescence stereoscope equipped with a 6× objective lens and Retiga 12-bit camera. The green fluorescent protein–based Ca2+ indicator, GCaMP6s, was excited using a mercury lamp (Intensilight Hg Illuminator, Zeiss), and a 440/470-nm excitation filter. The green fluorescent signal passing through a 535/550-nm filter was acquired every 2 s with the Retiga 12-bit camera using iVision software (BioVision Technologies).

To elicit wound responses, Arabidopsis roots were severed with a pair of scissors. For glutamate treatment, after a 15-min recovery period after cutting, 100 mM glutamate (solved in ddH2O and adjusted pH to 5.5 using tris base) was added to the cut site.

Using the ImageJ software, GCaMP6s signals were analyzed overtime at several regions of interest. To calculate the fractional fluorescence changes (ΔF/F), the equation ΔF/F = (FF0)/F0 was used, where F0 denotes the average baseline fluorescence determined by the average of F over the first 10 frames of the recording before the wound/treatment (72).

Total RNA isolation, cDNA synthesis, and qPCR

For quantitative polymerase chain reaction (qPCR) analysis, the main roots of Col-0 and the glr3.3 and glr3.6 mutant were severed with scissors at 2 cm away from the root-shoot junction. After the wound, shoot samples were harvested at 0, 0.5, 1, and 2 hours, and rapidly frozen in liquid nitrogen. Total RNA was extracted from plant samples using TRIzol reagent (Invitrogen), followed by synthesis of Poly (dT) cDNA using the M-MLV Reverse Transcriptase (Promega). Real-time quantitative reverse transcription polymerase chain reaction (qRT-PCR) was performed using the SYBR Green Supermix (Bio-Rad) according to the manufacturer’s instructions on a CFX96 Connect Real-Time System (Bio-Rad). All individual reactions were done in triplicate. The primers used for qRT-PCR are listed in table S2.

SUPPLEMENTARY MATERIALS

stke.sciencemag.org/cgi/content/full/13/640/eaba1453/DC1

Fig. S1. Systemic [Ca2+]cyt waves and SWPs induced by cutting a lateral root.

Fig. S2. Leaf-to-leaf SWPs induced by wounding and glutamate.

Fig. S3. Participation of GLR3.3 and GLR3.6 in wound- and glutamate-triggered root-to-shoot [Ca2+]cyt waves.

Fig. S4. Inhibition of glutamate-induced SWPs by FC.

Fig. S5. Conductivity of GLR3.3 to divalent and monovalent cations.

Fig. S6. Conductivity of GLR3.6 to divalent and monovalent cations.

Table S1. Relative ion permeability of GLR3.3 and GLR3.6.

Table S2. Primers.

Movie S1. Systemic [Ca2+]cyt waves in response to cutting the main root.

Movie S2. Root-to-shoot [Ca2+]cyt waves induced by application of glutamate at the wound site.

Movie S3. Loss of root-to-shoot [Ca2+]cyt waves in a glr3.3glr3.6 mutant upon root wounding.

Movie S4. Loss of root-to-shoot [Ca2+]cyt waves in a glr3.3glr3.6 mutant upon application of glutamate at the wound site.

Movie S5. [Ca2+]cyt imaging of control HEK293T cells upon application of glutamate at pH 8.5.

Movie S6. [Ca2+]cyt imaging of GLR3.6-expressing HEK293T cells upon application of glutamate at pH 8.5.

Movie S7. [Ca2+]cyt imaging of GLR3.6-expressing HEK293T cells upon application of histidine at pH 8.5.

REFERENCES AND NOTES

Acknowledgments: We thank J.-M. Zhou for providing ost2-2D seeds, and J. A. Feijó and E. E. Farmer for providing cDNAs encoding GLR3.3 and GLR3.6. Funding: Q.S. was supported by a scholarship from the China Scholarship Council and Q.G. was supported, in part, by the Young Scientists Fund of the National Natural Science Foundation of China (grant no. 31600245). This work was supported by a grant from the NSF (MCB-1714795, to S.L.). Author contributions: S.L. and Q.G. designed the research. Q.G. performed electrophysiology analysis; Q.S. performed whole-plant calcium imaging and q-PCR; Q.S. and D.L. performed SWP recordings; Q.S., Q.G., and H.Z. analyzed the data; and Q.G. and S.L. wrote the paper. Competing interests: The authors declare that they have no competing financial or nonfinancial interests. Data and materials availability: All data needed to evaluate the conclusions in the paper are present in the paper or the Supplementary Materials. Plasmids and Arabidopsis mutants are available upon request.

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