The potassium transporter TaNHX2 interacts with TaGAD1 to promote drought tolerance via modulating stomatal aperture in wheat

Abstract

Drought is a major global challenge in agriculture that decreases crop production. γ-Aminobutyric acid (GABA) interfaces with drought stress in plants; however, a mechanistic understanding of the interaction between GABA accumulation and drought response remains to be established. Here we showed the potassium/proton exchanger TaNHX2 functions as a positive regulator in drought resistance in wheat by mediating cross-talk between the stomatal aperture and GABA accumulation. TaNHX2 interacted with glutamate decarboxylase TaGAD1, a key enzyme that synthesizes GABA from glutamate. Furthermore, TaNHX2 targeted the C-terminal auto-inhibitory domain of TaGAD1, enhanced its activity, and promoted GABA accumulation under drought stress. Consistent with this, the tanhx2 and tagad1 mutants showed reduced drought tolerance, and transgenic wheat with enhanced TaNHX2 expression had a yield advantage under water deficit without growth penalty. These results shed light on the plant stomatal movement mechanism under drought stress and the TaNHX2-TaGAD1 module may be harnessed for amelioration of negative environmental effects in wheat as well as other crops.

TaNHX2 exerts flexible role in stomatal response under drought stress via modulating GABA accumulation in wheat.

INTRODUCTION

Drought stress is the most prevalent environmental factor limiting crop production and productivity. It may become even more grievous due to global climate change (1, 2). Wheat is one of the important staple crops supplying 20% of the total dietary calories globally (3, 4). Wheat is grown in the regions of the world that are most susceptible to drought, and its production is frequently compromised by water scarcity (5). Consequently, understanding the mechanisms and improving wheat tolerance to drought have been a high priority for research activities and wheat breeding programs (6–8).

Throughout evolution, plants have developed adaptive strategies to survive and grow under conditions of low moisture in their environments. Optimization of water uptake and minimization of water loss are two strategies associated with drought acclimation in plants. This is achieved through a variety of adaptive traits, including plastic root system and efficient stomatal function (9, 10). Stomatal closure is the first reaction to drought stress in most plants. Controlling water transpiration through stomata is one of the main strategies for plants to increase drought tolerance and therefore has been the target of molecular breeding strategies for improving drought tolerance in crops (11–14). Extensive evidence indicates that stomata respond directly to changes in air humidity and the water status of distant tissues via the stress hormone abscisic acid (ABA). When guard cells perceive increased ABA levels, their turgor and volume are reduced by the efflux of anions and potassium (K⁺) across the plasma and tonoplast membranes, causing stomatal closure (15). Activation of anion channels at the vacuolar and plasma membranes of guard cells has been regarded as a critical step in stomatal closure. Anion efflux via anion channels causes membrane depolarization, which subsequently drives K⁺ efflux from guard cells through outward-rectifying K⁺ channels (16). Among the solutes released from guard cells, more than 90% originate from vacuoles (17). Further evidence indicates that vacuolar cation/H⁺ antiporter (NHX)–type exchangers also regulate the stomatal aperture by mediating active K⁺ efflux from vacuoles (18, 19). Increasing studies on heterologous and homologous expression in Escherichia coli, yeast, and plant species have been reported for the functional characterization of NHX genes (20–25). Although several NHX exchangers that affect stomatal movement and drought tolerance have previously been identified in plants (26, 27), the NHX-mediated regulatory mechanisms of stomata-regulated drought tolerance in crops remain largely unexplored.

γ-Aminobutyric acid (GABA) is a ubiquitous non-protein amino acid that has been found in uni- and multicellular organisms and is involved in many aspects of plant development (28–30). GABA is mainly produced by the decarboxylation of glutamate, and this process is catalyzed by l-glutamate decarboxylase (GAD). Knockout (KO) of GAD genes markedly reduces the GABA levels (31, 32). An increasing number of studies have suggested that GABA may be a signaling molecule in addition to a metabolite in plants, and this speculation is further strengthened by evidence that GABA application effectively mitigates leaf damage induced by various abiotic stresses in plants (28, 33–38). GABA depletion resulting from GAD1 and GAD2 mutations affects stomata closure and drought tolerance in Arabidopsis. The drought-oversensitive phenotype of the gad1/2 mutant is reversed by functional complementation, which increases GABA accumulation in leaves (39). GABA can bind anion channels and the aluminum-activated malate transporter (ALMT) and regulate their activities in different plant species, suggesting that ALMTs are key transducers of GABA signaling in plants (40, 41). A recent study further revealed that cytosolic GABA signals affect stomatal opening and water use efficiency (WUE) by negatively controlling ALMT9 activity (42). However, the regulators of GABA accumulation on stomatal closure under drought remain largely unknown. As the GABA concentration increases when plants are exposed to diverse abiotic stresses, there are probably regulators with important roles in stress tolerance that affect the GABA synthesis (37, 43–45). However, not much is known about the molecular basis of GABA regulation for its accumulation under drought conditions, which helps plants tolerate and thrive under this stress, and the molecular mechanisms of drought tolerance in crops need to be further explored.

In this study, we showed that the vacuolar-localized potassium/proton exchanger protein TaNHX2 contributes to drought resistance in wheat by modulating the stomatal aperture. Specifically, the results indicate that TaNHX2 affects stomatal behavior partially dependent on GABA accumulation by promoting TaGAD1 activity, in addition to its own K⁺/H⁺ antiporter function. In addition, the fundamental role of the NHX2-GAD1 module in regulating the drought response pathway appears to be conserved across monocot and dicot plants. Our results thus uncover an unanticipated mechanism by which plants enhance drought resilience by targeting a tonoplast-localized cation exchanger to GABA accumulation via glutamate decarboxylase.

RESULTS

TaNHX2 is a putative vacuolar K⁺/H⁺ antiporter in wheat

To identify K⁺/H⁺ antiporter (NHX)–type exchangers in wheat, the amino acid sequence of rice OsNHX1 (46) was used as a query sequence and aligned against sequences in the wheat annotation project database. It is well known that common wheat is an allohexaploid species and contains A, B, and D subgenomes. Therefore, most of the genes present in common wheat as triplicate homoeologs are derived from ancestral species (47, 48). Eighteen putative NHX-type exchanger family members belonging to six complementary sets of homoeologs with collinear order across individual chromosomes of three subgenomes were identified in the wheat genome. On the basis of a phylogenetic tree generated by the NHX family members of wheat, rice, Brachypodium, and Arabidopsis (19, 46), these proteins were divided into three subclasses corresponding to their respective orthologs among these four species (fig. S1A), suggesting that NHX family members are well conserved among different species and likely play fundamental yet distinct roles in plant development and stress responses. In a recent single cell–type transcriptomic analysis, the transcript of TaNHX2 was preferentially enriched in wheat guard cells (49); thus, TaNHX2 was selected for further analysis.

Wheat TaNHX2 clustered with Brachypodium BdNHX2, rice OsNHX2, and Arabidopsis AtNHX2 in class I (fig. S1A). TaNHX2 shared 89.77, 85.53, and 68.6% amino acid identity with BdNHX2, OsNHX2, and AtNHX2, respectively, and exhibited more than 97% identity among the three homoeologs (fig. S1B). Eleven putative transmembrane domains and cation/H⁺ exchanger domains were predicted in TaNHX2 by TMHMM; moreover, protein structure prediction showed that the C terminus of TaNHX2 was a cytosolic domain (fig. S2, A to C). The domain for cation/H⁺ exchanger activity of NHX2 was highly conserved in four species, suggesting that TaNHX2 plays a fundamental role in ion transport, while their multivariate C-terminals implied other functions or molecular interactions in different members. The function of the TaNHX2 protein as a transporter was shown by yeast complementation. In yeast strain Δnhx1, which was deficient in the ScNHX1 gene, growth was strongly inhibited at high Na⁺ and K⁺ concentrations; in contrast, the positive transformant of Δnhx1 containing the TaNHX2 construct notably improved growth compared with the mutants, although it was not fully recovered to the wild-type level (Fig. 1A and fig. S3). In line with this, the growth curves also illustrated the trends in growth rates between wild-type and complementation yeast under normal and stress conditions (fig. S4, A and B).

Fig. 1. TaNHX2 is a putative vacuolar K⁺/H⁺ antiporter in wheat.

(A) Complementation of a yeast nhx1 mutant with TaNHX2. The yeast strain Δnhx1 was transformed with either the empty vector pDR195 or with TaNHX2, respectively, cloned into the yeast expression vector pDR195. Positive transformants and WT (W303-1B) were grown on the Yeast extract Peptone Dextrose medium with Agar (YPDA) medium supplemented with 200 or 300 mM NaCl. (B) Subcellular localization assays of TaNHX2 in wheat protoplast cells. The AtNHX2 was used to construct the tonoplast marker (19). BF, bright field. Scale bar, 10 μm. (C) RT-qPCR analysis of the expression pattern of TaNHX2 in different tissues. (D and E) Expression pattern of TaNHX2 under drought stress conditions in shoots (D) and roots (E) of seedlings. The seedlings were drought treated for different days according to VWC% after water withholding and detached samples for expression assay. TaACTIN was used as the internal control. The values are means (± SE) of three biological replicates. Data are means ± SE. * and *** indicates significant difference at P < 0.05 and P < 0.001 probability, respectively. CK, control.

The subcellular localization of TaNHX2 in wheat was investigated by transiently expressing 35S: TaNHX2-GFP and tonoplast marker AtNHX2 in mesophyll protoplast cells isolated from wheat leaves. The results showed that TaNHX2 was mainly localized in tonoplasts, similar to AtNHX2 in Arabidopsis (Fig. 1B and fig. S5). Furthermore, TaNHX2 expression showed different patterns in various tissues. The highest transcript level of TaNHX2 occurred in glumes, followed by roots, and the expression was moderate and similar among the other tested tissues including leaves, stems, spikes, and seeds (Fig. 1C). In addition, the transcriptional response of TaNHX2 to drought stress was examined. Reverse transcription quantitative polymerase chain reaction (RT-qPCR) analyses showed that drought treatments strongly promoted TaNHX2 expression. In leaves, TaNHX2 transcripts increased at 8 days after water withholding, peaked at 10 days, and slightly declined but were substantially higher than the control until 12 days, while the expression increased to the highest levels in roots at 12 days after water withholding (Fig. 1, D and E). In addition, TaNHX2 showed similar expression patterns within the time indicated, ruling out the potential effect of circadian rhythm (fig. S6). These results suggest that TaNHX2 may act as one of the K⁺/H⁺ transporters in tonoplast vesicles and contribute to the regulation of drought response in wheat.

TaNHX2 functions as a positive regulator of drought avoidance in wheat

To determine the functions of TaNHX2 in the regulation of wheat drought resistance, two TaNHX2 KO mutant lines, N-cr1 and N-cr2, were generated using the CRISPR-Cas9 system. The guide RNA was designed to target a highly conserved region in the fourth exon among the three homoeologs of TaNHX2 and was specifically compared with other members of the NHX family (Fig. 2A). The homozygous mutant with all three TaNHX2 homologs simultaneously knock out was identified and selected for further study. Sequencing analyses showed that N-cr1 conferred a 2–base pair (bp) deletion in TaNHX2-A, TaNHX2-B, and TaNHX2-D and that N-cr2 conferred a 2-bp deletion in TaNHX2-A, a 1-bp deletion in TaNHX2-B, and an 8-bp deletion in TaNHX2-D, causing frameshifting and truncation of TaNHX2-A, TaNHX2-B, and TaNHX2-D homeologs (Fig. 2A). Under normal conditions, no obvious phenotypic changes or developmental abnormalities were observed between the KO lines and wild-type plants (Fig. 2B). To explore drought stress tolerance conferred by the mutation of TaNHX2, the seedlings of KO lines were subjected to drought stress together with wild-type plants. Under drought stress treatment, both the wild-type plants and KO lines displayed a certain degree of growth inhibition. However, both N-cr1 and N-cr2 exhibited more severe inhibition, more wilted leaves (Fig. 2B), and lower survival rates (Fig. 2E) than the wild-type plants. Accordingly, the shoot weight analysis under drought stress conditions showed that the KO plants had a 19.66% reduction in fresh weight and a 17.54% reduction in dry weight compared with the wild-type plants, whereas under normal conditions, there was no difference in dry weight (Fig. 2, F and G). In addition, there were no obvious phenotypic changes in roots between the KO and wild-type plants under either normal or drought stress conditions (Fig. 2, K and L).

Fig. 2. TaNHX2 positively improves the drought tolerance of wheat.

(A) Gene structure and mutation sites of TaNHX2. The symbol “*” indicates the nucleotide deletion, and the base numbers of deletion are shown behind. (B) Drought stress tolerance assay of TaNHX2 KO transgenic and wild-type (WT) plants. Scale bars, 3 cm. (C) Relative transcription levels of TaNHX2 in transgenic overexpression plants were shown. The expression of TaACTIN was used to normalize mRNA levels. The values are means (± SE) of three biological replicates. (D) Drought stress tolerance assay of TaNHX2 overexpression and wild-type plants. Scale bars, 3 cm. (E) Statistical analysis of the survival rates of KO and wild-type plants after drought treatment and recovery. (F and G) Fresh and dry weight of KO and wild-type plants under normal conditions (F) and drought stress conditions (G) (n = 20). (H) Statistical analysis of the survival rates of OE and wild-type plants after drought treatment and recovery. (I and J) Fresh and dry weight of OE and wild-type plants under normal conditions (I) and drought stress conditions (J) (n = 20). (K) Phenotypic analysis in roots of TaNHX2 OE, KO, and wild-type plants under normal conditions or drought stress conditions. Scale bars, 3 cm. (L) Statistical analysis of root length of different transgenic plants (n = 10). Data are means ± SE. *, **, and *** indicates significant difference at P < 0.05, P < 0.01, and P < 0.001 probability, respectively.

To further validate the functions of TaNHX2 in drought tolerance, we generated transgenic wheat plants overexpressing TaNHX2 (TaNHX2-OE). Ten putative TaNHX2-OE lines of wheat cultivar Fielder were obtained, and two TaNHX2-OE lines (N-OE1 and N-OE2) with significantly elevated expression of TaNHX2 were selected for further study (Fig. 2C). The seedlings of the OE lines were subjected to drought stress together with the wild-type plants. In contrast to the KO plants, the OE plants exhibited a higher survival rate (Fig. 2H) and fewer wilted leaves than the wild-type plants under drought conditions, while they were unchanged under normal conditions (Fig. 2, D and I). Phenotypic measurement and statistical analysis revealed that the OE plants increased fresh weight by 15.49% and dry weight by 11.89% compared with the wild-type plants, while under normal conditions, the OE and wild-type plants were comparable in shoot weight (Fig. 2, I and J). Furthermore, consistent with the KO plants, there were no differences in root development between the OE and wild-type plants under normal or drought stress conditions (Fig. 2, K and L). Collectively, these results indicate that TaNHX2 acts as a positive regulator of drought avoidance in wheat.

TaNHX2 modulates stomatal closure under drought stress

To further investigate how TaNHX2 enhances drought tolerance in wheat, the water loss of detached leaves was examined in different transgenic lines. Both N-cr1 and N-cr2 had faster water loss than the wild-type plants, while water loss in the OE lines was significantly slower than in the wild-type plants (Fig. 3A). In addition, electrical conductivity and chlorophyll content were also measured, but there was no substantial difference in any of the transgenic plants (fig. S7, A and B). Considering that water loss mainly occurs through the stomata in plants and because there were significant differences in water loss in detached leaves of the TaNHX2-OE and KO lines, we further investigated the changes in the stomata in different transgenic lines. The stomatal density was then examined, and results showed that the average stomatal density was comparable between the OE, KO, and wild-type plants, suggesting that the stomatal density was not affected by TaNHX2 (fig. S8, A and B). Subsequently, the leaf stomatal aperture in different transgenic plants was observed using an inverted microscope (AE31E, Motic). As shown in Fig. 3B, the stomatal aperture in both the OE and KO plants was comparable to the wild-type plants under normal conditions. As expected, under drought stress conditions, the mean stomatal apertures of the wild-type and transgenic wheat plants decreased correspondingly. However, the OE plants exhibited a higher stomatal closure than the wild-type plants, In contrast, the KO plants exhibited lower stomatal closure than the wild-type plants, indicating that TaNHX2 is involved in modulating stomatal closure under drought stress in wheat (Fig. 3B). In addition, stomatal conductance was decreased in the OE lines and increased in the KO lines compared with the wild-type plants under drought stress conditions (Fig. 3C). Furthermore, transiently expressing Pro^TaNHX2-CDS^TaNHX2-mCherry in tobacco indicated that TaNHX2 was expressed in the guard cells, consistent with its vital role in stomatal movement (Fig. 3D and fig. S9).

Fig. 3. TaNHX2 modulates stomatal closure under drought stress through affects GABA accumulation and GAD enzyme activity.

(A) Water loss of detached leaves of different transgenic plants (n = 3). (B) Stomatal aperture of different transgenic plants in response to drought stress. Epidermal strips were pre-incubated in the stomatal-opening buffer for 1 hour under dark, followed by a 10-min, under constant dark, drought stress (in the air) or normal conditions (in buffer), then incubation dark-to-light transition for 1.5 hours as indicated in the above graphs by the black (dark) or white (light) bars. ns, not significant. (C) Stomatal conductance of different transgenic plants (n = 6). (D) Localization assays of TaNHX2 in N. benthamiana leaves. Scale bar, 20 μm. (E) Infrared thermography of different transgenic plants under normal conditions and drought stress conditions. Scale bars, 3 cm. (F and G) Leaf temperature measurement of CK (F) and drought (G) from (E) using infrared camera software (n = 20, temperature of dots). Means with the same letter did not significantly differ at P < 0.05, according to Tukey’s test. (H) GABA accumulation in the leaves of TaNHX2-OE, KO, and wild-type plants under normal conditions and drought stress conditions (n = 8). (I) GAD enzyme activity in the leaves of OE, KO, and wild-type plants under normal conditions and drought stress conditions (n = 5). The relative GAD activity of the WT under normal conditions was normalized to 1, and the relative GAD activity in each kind of genotype was calculated by comparing wild type. (J and K) Stomatal aperture of WT leaves in response to light or dark. Data are mean ± SE. *, **, and **** indicate significant difference at P < 0.05, P < 0.01, and P < 0.0001 probability, respectively.

Because water loss by transpiration leads to temperature changes on the leaf surface, the leaf surface temperature of the OE, KO, and wild-type plants was measured using an infrared thermal imager. As expected, under drought stress conditions, the leaf surface temperature of the OE plants was higher than that of the wild-type plants, whereas the KO plants exhibited a lower temperature than the wild-type plants, with no substantial difference in plants under normal conditions (Fig. 3, E to G). To determine whether TaNHX2 promotes ABA sensitivity of stomatal closure in wheat, the leaf surface temperatures of the OE, KO, and wild-type plants were measured under ABA treatment. Intriguingly, there was no substantial difference among the OE, KO, and wild-type plants after ABA application, although all plants showed a higher temperature than the corresponding control (fig. S10, A and B). Furthermore, the transcriptional response of TaNHX2 to ABA treatment was examined, which was induced sluggishly by ABA treatment in both leaves and roots, with no substantial difference in the early phase (fig. S10, C and D). These results imply that, in addition to ABA signaling, TaNHX2-mediated stomatal closing may also be involved in other signal pathways.

TaNHX2 affects endogenous GABA accumulation and the GAD activity under drought stress in wheat

The non-protein amino acid GABA has been proposed as an important messenger and plays a signaling role in stomatal regulation under water deficiency (39, 40, 42). To investigate whether TaNHX2 affects endogenous GABA accumulation levels to modulate the stomatal aperture under drought stress, the GABA content was measured in different TaNHX2 transgenic lines. Under normal conditions, there was no substantial difference between the OE, KO, and wild-type plants (Fig. 3H). Consistent with previous studies, GABA accumulation was higher in drought-stressed leaves than in well-watered leaves. Intriguingly, the GABA accumulation of both N-OE1 and N-OE2 was substantially higher than that of the wild-type plants; in contrast, a reduction of GABA accumulation in both KO mutants was observed (Fig. 3H).

GABA is primarily synthesized from the decarboxylation of glutamate by the l-glutamate decarboxylase GAD, especially under stress conditions (32). We further examined GAD enzyme activity in different TaNHX2 transgenic lines. Under normal conditions, the GAD activity of both TaNHX2 overexpression and KO lines was comparable to that of the wild-type control, although N-cr2 showed a decline (Fig. 3I). However, under drought stress conditions, GAD activity was significantly enhanced in independent overexpression lines but decreased in TaNHX2 KO mutants compared with the wild-type (Fig. 3I). These data were consistent with our above observation of GABA accumulation, indicating that TaNHX2 enhances GAD activity and further improves endogenous GABA content under drought stress in wheat.

GABA application can partially rescue the stomatal defect phenotype of the tanhx2 mutant under drought stress

The variance of GABA accumulation in different TaNHX2 transgenic plants revealed above prompted us to further determine whether GABA is a regulator of stomata control in wheat. As expected, consistent with other plant species (42), we found that GABA application suppressed light-induced stomatal opening and dark-induced stomatal closure in wheat. This suggests that GABA also has the potential to be an excellent controller of stomatal movement in wheat, enabling plants to cope with the hazards of drought stress by intelligently regulating stomatal movement (Fig. 3, J and K). To further determine whether TaNHX2 modulated the wheat stomatal closure under drought stress via GABA accumulation, we examined the effect of GABA on leaf surface temperature in the TaNHX2-OE, KO, and wild-type plants under drought stress (fig. S11, A to C). When exogenous GABA with different concentration gradients was applied before drought stress treatment, the leaf surface temperature of both the wild-type and N-cr plants was significantly improved compared with that of those treated with drought stress alone, and it was especially ameliorated in N-cr lines. Notably, pretreatment with 0.2 mM GABA showed a great advance in recovering the leaf surface temperature of N-cr to wild-type levels under drought stress (fig. S11, A to C). An interesting additional observation was that the N-OE plants pretreated with GABA had temperatures comparable to those treated with drought stress alone, indicating that there was sufficient endogenous GABA in the N-OE plants to regulate stomatal movement under drought stress (fig. S11, A to C).

TaNHX2 affected the drought response in wheat, partially dependent on GABA accumulation, implying that other functions of TaNHX2 were also disrupted in N-cr lines. This is understandable because TaNHX2 might encode a tonoplast-localized K⁺/H⁺ antiporter and function in guard cells via its fundamental role in ion transport through the tonoplast. To further clarify the function of TaNHX2 for the K⁺ transporter, the yeast complementation assay was performed using yeast strain R5421, which carries deletions in the K⁺ uptake systems TRK1/TRK2. The mutant yeast strain was unable to grow at low K⁺ concentrations; in contrast, the positive transformant of R5421 containing the TaNHX2 construct grew at low K⁺ concentrations (fig. S12C). Moreover, we performed a K⁺ efflux measurement assay in Xenopus laevis oocytes by pro-injecting 50 nmol of K⁺, incubating the oocytes in a K⁺-free medium, and measuring K⁺ content in the oocytes and medium. TaNHX2-expressed oocytes showed a lower K⁺ content in oocytes and a higher K⁺ efflux in the media than water-injected control oocytes, demonstrating that TaNHX2 mediates K⁺ transport in oocytes (fig. S12, A and B). These results indicate that TaNHX2 functions as a K⁺ transporter, suggesting its role in ion transport in guard cells under drought conditions. Together, we propose that TaNHX2 affects stomatal behavior partially dependent on GABA accumulation and partially dependent on its own tonoplast K⁺ transporter function.

In addition, we detected the expression pattern of TaNHX2 in response to exogenous GABA treatment. The expression of TaNHX2 was induced by exogenous GABA in shoots and peaked at 6 hours, while it was slightly inhibited in the roots (fig. S13, A and B). Consistent with the expression pattern of TaNHX2 upon GABA treatment in the roots, there were no substantial differences in root length or lateral root number in different transgenic lines with or without exogenous GABA (fig. S14, A to C).

TaNHX2 interacts with the TaGAD1 protein

GAD1 and GAD2 are key regulatory points in GABA biosynthesis. To further explore the mechanism of TaNHX2, which modulates stomatal pore aperture via GABA accumulation under drought stress, two GAD homologs in the wheat genome (TaGAD1 and TaGAD2) were identified from the wheat annotation project database. Tissue-specific expression pattern analysis showed that TaGAD1 was expressed in different tissues and highly expressed in roots and spikes, while TaGAD2 was mainly expressed in spikes, glumes, and seeds (fig. S15, A and B). These expression patterns were inconsistent with those in Arabidopsis, in which AtGAD1 was expressed mainly in roots and AtGAD2 was expressed in all organs including roots, leaves, inflorescence stems, and flowers (50). Furthermore, the alignment of the TaGAD1 and TaGAD2 amino acid sequences revealed a conserved N terminus containing the glutamate decarboxylation core domain and a variable C terminus containing a predicted Ca²⁺/CaM binding domain (fig. S15, C and D). We further investigated the subcellular distribution of TaGAD1 and TaGAD2 by expressing TaGAD1–green fluorescent protein (GFP) and TaGAD2-GFP fusion proteins in wheat protoplast cells, and both TaGAD1-GFP and TaGAD2-GFP were localized in the cytoplasm (Fig. 4A and fig. S15E).

Fig. 4. TaNHX2 interacts with TaGAD1.

(A) Subcellular localization assays of TaGAD1 in wheat protoplast cells. Scale bars, 10 μm. (B) LCI assay shows that TaNHX2 interacts with TaGAD1. nLUC-tagged TaNHX2 was cotransformed into tobacco leaves along with cLUC-tagged TaGAD1. TaFO3 and TaGRF were used as negative controls. (C) Verification of protein interaction by split-ubiquitin yeast system. All the transformation colonies were grown on the SD/-Trp-Leu and SD/-Leu-Trp-His-Ade plates, followed by the examination of LacZ reporter gene with X-Gal as a substrate. The combination of APP and Fe65 was used as the positive control. (D) Co-IP assays showing the physical interactions of TaNHX2 and TaGAD1. N. benthamiana leaves were transformed by injection of the Agrobacterium GV3101 cells containing TaNHX2-MYC and TaGAD1-GFP plasmids. Total proteins were immunoprecipitated with GFP-Trap-A, and the immunoblots were probed with the anti-GFP and anti-MYC antibodies, respectively. (E) BiFC assay revealing the interaction of TaNHX2 with TaGAD1. cYFP-tagged TaNHX2 was cotransformed into tobacco leaves along with nYFP-tagged TaGAD1. Scale bars, 50 μm. (F) BiFC assay revealing the interaction of TaNHX2 with TaGAD1 in tobacco protoplast. The OsVPE1 was used to construct the tonoplast marker (51). Scale bar, 10 μm.

The result that TaNHX2 modulated stomatal closure via GABA accumulation under drought stress, together with the potential role of TaGADs in the GABA biosynthesis process, implies that TaNHX2 interacts with TaGADs. We then performed split Luciferase Complementation Imaging Assay (LCI) to determine whether TaNHX2 interacts with these two GAD proteins. Intriguingly, we found that TaGAD1 interacted with TaNHX2, while TaGAD2 did not. Coexpression of cLUC-TaGAD1 with TaNHX2-nLUC generated strong luminescence signals that were not detected in the negative control pairs or when coexpressing TaGAD2 and TaNHX2 (Fig. 4B and fig. S16A). Notably, the signal was further enhanced in the interaction between TaNHX2-nLUC and cLUC-TaGAD1 under drought stress conditions (fig. S17, A and B). We also verified that TaNHX2 interacted with TaGAD1 through a split-ubiquitin yeast system on the quadruple dropout media, followed by the examination of the LacZ reporter gene with X-Gal as a substrate. The yeast cotransformed with pBT3-STE-TaNHX2 and pPR3-N-TaGAD1 grew better than those transformed with the corresponding control, and the chromogenic reaction was also consistent with the positive control (Fig. 4C). Furthermore, co-immunoprecipitation (Co-IP) assays were conducted to confirm the TaNHX2-TaGAD1 interaction using tobacco expressing TaNHX2-MYC, TaGAD1-GFP, and GFP alone by Agrobacterium-mediated transformation; compared with IP using an antibody with GFP, immunoblot assays using an anti-MYC antibody detected a band of TaNHX2-MYC in the TaGAD1-GFP sample, but no band was identified in the GFP sample (Fig. 4D). We then corroborated the TaNHX2-TaGAD1 interaction using bimolecular fluorescence complementation (BiFC) assays, in which an excited yellow fluorescent protein (YFP) reconstitution signal was observed using a laser scanning confocal fluorescence microscope when the fusion constructs TaNHX2-cYFP and nYFP-TaGAD1 were present but not when those transformed with the corresponding control were present (Fig. 4E). Meanwhile, no interaction was detected between TaNHX2 and TaFO3 or TaGRF, two proteins localized in the cytoplasm (fig. S16B), suggesting that the TaNHX2-TaGAD1 interaction in vivo is specific. To further investigate the subcellular distribution of the TaNHX2-TaGAD1 interaction, we co-infiltrated TaNHX2-cYFP, nYFP-TaGAD1, and tonoplast marker OsVPE1 (51) in the mesophyll cells of Nicotiana benthamiana. The tobacco protoplasts collected 48 hours after infiltration yielded merged fluorescence in the tonoplast (Fig. 4F and fig. S16C). Notably, however, we could not detect the luminescence signals when the fusion constructs cLUC-TaNHX2 and TaGAD1-nLUC were present (fig. S16D). In line with this, we also could not detect fluorescence signals when nYFP-TaNHX2 and TaGAD1-cYFP were coexpressed in N. benthamiana leaves (fig. S16E), suggesting that the interaction of TaNHX2 with TaGAD1 is cytosolic domain dependent. These results indicate that TaNHX2 physically interacts with TaGAD1 in vivo. Collectively, we propose that TaNHX2 recruits TaGAD1 localized on the tonoplast, considering that GABA inhibits ALMT9 on the tonoplast and promotes stomatal closure (42) and causes more GABA accumulation surrounding the tonoplast, further weakening vacuolar anion transport.

TaNHX2 targets the C-terminal auto-inhibitory domain of TaGAD1 and up-regulates its activity

Previous reports have indicated that GAD proteins in plants have a region of approximately 30 amino acids at their C terminus, which can bind to CaM to regulate GAD activity (52, 53). Moreover, GADs in Arabidopsis and rice with deletion of this region (auto-inhibitory region) tend to exhibit higher glutamate decarboxylase activity than full-length GAD (39, 54). The findings that TaNHX2 affects the drought response in wheat by affecting endogenous GABA accumulation, together with the physical interaction of TaNHX2 and TaGAD1, prompted us to investigate whether TaNHX2 could mediate TaGAD1 activity. To address this possibility, we further mapped the domains of the TaNHX2-TaGAD1 interaction using LCI assays. Immunoblot analysis and subcellular localization assays were performed to ensure that the localization and accumulation levels of truncated proteins of TaNHX2/TaGAD1 were comparable to those of full-length proteins (fig. S18, A to C). On this basis, we observed that the C terminus of TaGAD1 (TaGAD1-C) interacted with TaNHX2 (Fig. 5, A to D). Furthermore, the C terminus of TaNHX2 (TaNHX2C contains predicated intracellular part) but not the N terminus (TaNHX2N contains transmembrane domains) interacted with TaGAD1-C (Fig. 5, A to D). Moreover, we produced fusion proteins glutathione S-trasferase (GST)–TaGAD1 (full length), GST-TaGAD1ΔC (a form of TaGAD1 that has a C-terminal domain removed), GST–TaGAD1-C (C-terminal domain), and TaNHX2C-HIS (a form of TaNHX2 that has a transmembrane domain removed) in E. coli cells and used them for pull-down experiments. The results confirmed that TaNHX2C interacted with TaGAD1-C (fig. S19). To further investigate the biochemical consequences of the TaNHX2-TaGAD1 interaction, the fusion proteins GST-TaGAD1, GST-TaGAD1ΔC, and TaNHX2C-HIS were expressed in E. coli cells and used for GAD activity experiments. Immunoblot analysis using an antibody against GST showed comparable protein levels between GST-TaGAD1 and GST-TaGAD1ΔC, and consistent with this, co-incubation with TaNHX2C-HIS was also unchanged (Fig. 5E). The content of GABA and GAD enzymatic activity were then determined. We observed lower GABA accumulation and GAD enzymatic activity in GST-TaGAD1 than in GST-TaGAD1ΔC, indicating that the deletion of the C-terminal region of TaGAD1 substantively enhanced its glutamate decarboxylase activity. Notably, we observed that TaGAD1 activity was elevated to GST-TaGAD1ΔC levels by co-incubation with TaNHX2C-HIS (Fig. 5, F and G).

Fig. 5. TaNHX2 targets the C terminus auto-inhibitory domain of TaGAD1 and enhances its activity.

(A) Schematic diagram showing TaGAD1 and its derivatives containing specific protein domains. (B) Schematic diagram showing TaNHX2 and its derivatives with or without the Cation/H⁺ exchanger protein domain. (C) LCI assays show that the C terminus of TaGAD1 associates with TaNHX2. (D) LCI assays show that the TaNHX2-C is associated with the C terminus of TaGAD1 (TaGAD1-C). (E) GAD activity experiments detected evident bands indicative of the predicted GST-TaGAD1 and GST-TaGAD1ΔC size and showed comparable protein levels in all samples. (F and G) GABA accumulation (n = 5) (F) and relative GAD enzyme activity (n = 8) (G) measurements in the E. coli cells system. (H) GAD activity experiments detected evident bands indicative of the predicted TaGAD1-GFP and TaGAD1ΔC-GFP size and showed comparable protein levels in all samples. (I and J) GABA accumulation (n = 8) (I) and relative GAD enzyme activity (n = 6) (J) measurements in N. benthamiana leaves. The tobacco leaves only injected incubation solution were used as the negative control. The relative GAD activity of the TaGAD1 was normalized to 1, and the relative GAD activity in each kind of combination was calculated by comparing TaGAD1. Data are means ± SE. Means with the same letter did not significantly differ at P < 0.05, according to Tukey’s test.

To further confirm this observation, we also conducted GAD activity experiments in N. benthamiana. In line with previous studies (39, 54), the AtGAD1ΔC (a form of AtGAD1 that has a C-terminal domain removed) displayed higher GAD activity and GABA content than AtGAD1 using the tobacco system (fig. S20, A and B). Furthermore, the GABA content and GAD enzymatic activity were measured in the tobacco system expressing TaGAD1-GFP or TaGAD1ΔC-GFP. The protein levels of TaGAD1-GFP and TaGAD1ΔC-GFP were revealed by immunoblotting using an anti-GFP antibody, and the immunoblot data also showed comparable protein levels in all samples (Fig. 5H). Consistent with the results in the E. coli cell system, we observed that simultaneous expression of the fusion proteins TaGAD1-GFP and TaNHX2-MYC significantly increased GABA accumulation and GAD enzymatic activity compared with the coexpression of TaGAD1-GFP and the empty control (Fig. 5, I and J). The same assay was performed on TaNHX2C-MYC in the tobacco system and showed a consistent trend in GABA accumulation and GAD activity with full-length TaNHX2-MYC expression in the tobacco system (Fig. 5, I and J). The results of both in vitro and in vivo experiments supported the hypothesis that TaNHX2 acts on the TaGAD1-C and promotes TaGAD1 activity and that this effect is independent of its ion transporter activity since the TaNHX2C protein without the transmembrane domain of TaNHX2 (Fig. 5, E to J). Moreover, TaNHX2C without Ca²⁺ or CaM enhanced TaGAD1 activity, suggesting that TaNHX2 might be a candidate for a GAD enzyme activator, in addition to Ca²⁺/CaM. Collectively, our results showed that TaNHX2 targets the C-terminal of TaGAD1 and up-regulates TaGAD1 activity, resulting in synthesis of more GABA and contributing to drought resilience.

TaGAD1 positively regulates drought resistance in wheat

To verify the function of TaGAD1, we generated TaGAD1 mutants in Fielder using the CRISPR-Cas9 system. The A subgenome of TaGAD1 in Fielder is missing. Then, a frame-shift null mutant of TaGAD1 (G-cr1, with a 5-bp deletion in TaGAD1-B and a 1-bp insertion in TaGAD1-D) was obtained (fig. S21A). Seedlings of mutants, wild-type plants, and the negative transgenic (NT) control (the plants that underwent a transgenic event but no editing occurred) were exposed to a soil drought treatment in a pot experiment. As expected, compared with the wild-type and NT, G-cr1 exhibited significantly reduced drought resistance (Fig. 6A). The plant survival and biomass of G-cr1 were significantly lower than those of the wild-type and NT plants under drought conditions (Fig. 6B and fig. S21, B and C). Subsequently, we measured the GABA content and GAD enzyme activity in the leaves of G-cr1 and control plants. Under normal conditions, the GABA content of G-cr1 was lower than that of the wild-type and NT plants. Under drought stress treatments, the GABA content of all plants was elevated; however, the GABA content of G-cr1 was significantly lower than that of the wild-type and NT plants (Fig. 6C). Consistent with this, the GAD enzyme activity of different plants also showed similar variations in the G-cr1, wild-type, and NT plants under both control and drought stress conditions (Fig. 6D). In addition, there were no obvious phenotypic changes in the roots in the G-cr1 and wild-type, NT plants under normal and drought stress conditions (fig. S22, A and B), suggesting that TaGAD1 may not affect root development. On the basis of these results, we speculate that TaGAD1 is actively involved in the regulation of drought resistance in wheat.

Fig. 6. KO of TaGAD1 reduces drought resistance in wheat seedlings.

(A) Drought stress tolerance assay of TaGAD1 KO transgenic and control plants. Scale bars, 2 cm. (B) Statistical analysis of the survival rates of G-cr1 and control plants after drought treatment and recovery. (C) GABA accumulation in the leaves of G-cr1 and control plants under normal conditions and drought stress conditions (n = 7). (D) GAD enzyme activity in the leaves of G-cr1 and control plants under normal conditions and drought stress conditions (n = 6). The relative GAD activity of the WT under normal conditions was normalized to 1, and the relative GAD activity in each kind of genotype was calculated by comparing WT. (E) Water loss of detached leaves of G-cr1 and control plants (n = 3). (F) Localization assays of TaGAD1 in N. benthamiana leaves. Scale bar, 20 μm. (G) Stomatal aperture of different transgenic plants in response to drought stress. Epidermal strips were pre-incubated in the stomatal-opening buffer for 1 hour under dark, followed by a 10-min, under constant dark, drought stress (in the air) or normal conditions (in buffer), then incubation dark-to-light transition for 1.5 hours as indicated in the above graphs by the black (dark) or white (light) bars. (H) Infrared thermography of G-cr1 and control plants under normal conditions and drought stress conditions. Scale bars, 3 cm. (I) Leaf temperature measurement from (H) using infrared camera software (n = 20, temperature of dots). Data are means ± SE. *, **, and **** indicates significant difference at P < 0.05, P < 0.01, and P < 0.0001 probability respectively. Means with the same letter did not significantly differ at P < 0.05, according to Tukey’s test.

To further determine whether TaGAD1 enhances drought avoidance in wheat by affecting stomatal movement, the water loss of detached leaves was examined in the G-cr1, wild-type, and NT plants. G-cr1 showed faster water loss than the wild-type and NT plants (Fig. 6E). We also investigated the expression of TaGAD1 by transiently expressing Pro^TaGAD1-CDS^TaGAD1-mCherry in tobacco. Consistent with TaNHX2, the TaGAD1 protein was detected in the guard cells (Fig. 6F and fig. S23). Furthermore, in line with the TaNHX2 transgenic lines, the average stomatal density was similar between the G-cr1 and wild-type or NT plants, suggesting that stomatal density was not affected by TaGAD1 (fig. S24, A and B). The leaf surface temperature and stomatal pore aperture of the G-cr1 plants were measured and compared with those of the control plants. Under drought stress conditions, the G-cr1 plants had lower leaf temperatures than the controls, as expected from the higher transpiration and stomatal aperture, while there was no substantial difference in any of the plants under normal conditions (Fig. 6, G to I). Together, these data indicate that TaGAD1 positively regulates drought resistance by affecting stomatal movement in wheat.

To further verify the function of TaGAD1 in drought resistance, we constructed an overexpressing vector of TaGAD1 and obtained a positive transgenic line (GOE-1). Unfortunately, GOE-1 produced severe negative effects, including stunted panicles with kernel abortion, and kernel development also showed abnormality. Expression analysis showed that the transcript level of TaGAD1 in GOE-1 was nearly 400 times that of the wild type, and we speculated that the excessive expression of TaGAD1 caused the developmental abnormality of GOE-1 (fig. S25, A to F). These phenotypes were similar to those in a previous study in Arabidopsis (55). Thus, GOE-1 was not carried out for further analysis as experimental material in this study.

TaNHX2 improves wheat drought tolerance in reproductive stages

To determine the genetic effects of TaNHX2 in vegetative and reproductive stages under drought stress conditions in the field, the N-OE and N-cr plants were grown and compared with wild-type plants in well-watered and water-limited experiments (fig. S26A). We monitored and calculated the WUE in all of the TaNHX2 transgenic plants at the grain-filling stage. The WUE in both the N-OE and N-cr plants was comparable to the wild-type plants under normal conditions (Fig. 7A). However, under drought stress conditions, the WUE of the N-OE plants was higher than that of the wild-type plants, whereas the N-cr1 plants exhibited a lower WUE than the wild-type plants (Fig. 7B). Moreover, a comparison of yield-related traits under normal conditions revealed that the N-OE and N-cr plants were nearly identical to the wild-type plants in terms of seven important agronomic traits, including plant height, spike length, rachis number of spikes, grain number per spike, grain length, grain width, and thousand kernel weight (TKW). However, under drought stress conditions, the N-OE plants produced longer and wider grains and had a higher TKW than the wild-type plants (Fig. 7, C to J, and fig. S26, B and C). In contrast, the N-cr plants showed smaller grain sizes and produced a lower TKW than the wild-type plants (Fig. 7, C to J). Hence, these results demonstrate that, in wheat, the expression of TaNHX2 confers a yield advantage under water deficit and dose that do not cause a growth and development penalty under well-watered conditions.

Fig. 7. The WUE and yield performance of TaNHX2 wheat lines under field conditions.

(A and B) WUE capacity of WT and transgenic plants subjected to normal (A) and drought stress (B) under field conditions. (C) Phenotypic analysis of TaNHX2 OE, KO, and wild-type in grain width (n = 20) and length (n = 10). Scale bars, 1 cm. (D to J) Plant height (n = 20) (D), spike length (n = 15) (E), rachis number per spike (n = 12) (F), grain number per spike (n = 12) (G), grain length (n = 6) (H), grain width (n = 6) (I), and thousand kernel weight (n = 6) (J). Data are means ± SE. Means with the same letter did not significantly differ at P < 0.05, according to Tukey’s test. * indicates significant difference at P < 0.05 probability.

DISCUSSION

In this study, we provide genetic and biochemical evidence to demonstrate that a potassium/proton transporter TaNHX2 affects the drought response in wheat by controlling the activity of glutamate decarboxylase 1 (TaGAD1), thereby controlling GABA levels. We observed that transgenic wheat plants overexpressing TaNHX2 potentiated drought tolerance, whereas knocking out the TaNHX2 gene using the CRISPR-Cas9 method resulted in significantly drought sensitive phenotypes (Fig. 2), supporting the idea that TaNHX2 is a positive regulator in drought resistance in wheat. We propose that TaNHX2 modulates stomatal closure and drought avoidance transduced by interacting with the C-terminal auto-inhibitory domain of TaGAD1, up-regulates TaGAD1 activity, and promotes GABA accumulation under drought stress. GABA affects stomatal movement and increases WUE mainly by inhibiting ALMT9, causing anion efflux through the tonoplast (42). Thus, GABA accumulation surrounding the vacuole is a good response to drought. TaNHX2 preforms this process in two steps: recruiting more glutamate decarboxylase on the tonoplast and enhancing its activity for more GABA production (Fig. 8).

Fig. 8. A proposed model that TaNHX2 interacts with TaGAD1 to promote drought resistance in wheat.

The tonoplast-localized cation exchanger TaNHX2 contributes to drought resistance transduced by interacting with the C-terminal auto-inhibitory domain of TaGAD1, up-regulates TaGAD1 activity, and promotes GABA accumulation under drought stress. GABA has been found to reduce stomatal opening and increase WUE by negative modulation activity of a guard cell tonoplast-localized anion transporter ALMT9 (42). TaNHX2 promotes stomatal closure by enhancing GABA accumulation and signaling pathway upon drought stress and is essential for drought resistance in wheat.

TaNHX2 encodes a tonoplast-localized K⁺/H⁺ transporter that was grouped in the phylogenetic clade with AtNHX1 and AtNHX2 in our analysis (fig. S1A). However, reciprocal BLAST analysis revealed that TaNHX2 and AtNHX2 shared only a 68% amino acid identity, indicating both sequence conservation and diversification (fig. S1B). In Arabidopsis, both NHX1 and NHX2 are highly expressed in guard cells and functionally redundant; double mutant nhx1nhx2 exhibits a significantly reduced growth phenotype (24). Further experimental evidence indicates that NHX1 and NHX2 mediate active K⁺ uptake into vacuoles to regulate cell turgor and stomatal function. The nhx1nhx2 mutant exhibits pronounced leaf turgor loss and thus delayed stomatal closure compared with the wild type, indicating that the fast response mediated by rapid K⁺ fluxes is impaired in the mutant (19). In line with these observations, we found that TaNHX2 functions as a K⁺ transporter and confers drought avoidance by promoting stomatal closure (Figs. 1 to 3). In the present study, the overexpression of TaNHX2 was driven by the ubiquitin promoter, which might have complicated the phenotypes of our transgenic lines since the stomatal regulation is subtle. It is worthwhile to generate transgenic lines driven by the native cell types or guard cell–specific promoters and perform an in-depth analysis of phenotypes involved in stoma movement in future studies. Plants endure drought stress through a variety of adaptive traits, mainly involving the plastic root system and efficient stomatal function. In this study, there was no obvious phenotypic variation in root traits among the TaNHX2-OE, KO, and wild-type plants both under normal and drought stress conditions, suggesting that TaNHX2 specifically affects stomata to participate in drought resistance in wheat.

The regulation of the stomatal aperture is a critical driver of plant productivity and drought tolerance and has a remarkable influence on climate due to its influence on global carbon and water cycling (56). Upon exposure of plants to drought stress, ABA is the major hormone that initiates the adaptation of plants to drought stress through stomatal closure (14, 57). Notably, in this study, our observations suggest that TaNHX2 plays an unanticipated role in stomatal aperture modulation and that the relationship of this process with the ABA pathway remains to be explored because the expression response of TaNHX2 was sluggish upon ABA treatment in both leaves and roots in the early phase (fig. S10). An increasing amount of evidence has shown that GABA plays a protective role against drought stress in plants by increasing osmolytes and leaf turgor and reducing oxidative damage via antioxidant regulation (45). Recently, GABA was found to reduce stomatal opening and transpirational water loss via negative modulation activity of a stomatal guard cell tonoplast-localized anion transporter ALMT9, which is the major tonoplast-localized channel involved in anion uptake into guard cell vacuoles during stomatal opening (40, 42, 58), thus resulting in increased WUE and drought tolerance. In line with this finding, our study showed that TaNHX2 affected stomatal behavior partially dependent on GABA accumulation under stress conditions. These findings indicate that GABA’s metabolic and signaling pathways are an important layer of regulation for stomatal aperture in drought stress resistance in plants because the effects of GABA on stomata have been observed in different plant species, including dicot and monocot crops (e.g., Arabidopsis, wheat, barley, tobacco, and soybean) (38, 39). However, a central question remains largely unresolved: How are GABA’s metabolic and signaling pathways regulated under drought stress, and what are the mechanisms that underpin this process? In the present study, we revealed that TaNHX2 mediated stomatal movement under drought stress not only via the K⁺/H⁺ transporter but also through a noncanonical function regulating GABA biosynthesis and a signal pathway. We found that GABA accumulation was reduced in the TaNHX2 KO lines but improved in the overexpression lines compared with the wild-type plants under drought stress conditions (Fig. 3). Our observations indicate that TaNHX2 functions to modulate stomatal closure by promoting GABA accumulation in response to drought stress. We therefore uncovered the key regulatory factor that modulates the stomatal aperture by regulating GABA levels.

GABA is primarily synthesized from the decarboxylation of glutamate by GAD, and this pathway is conserved from bacteria to higher organisms. Studies on the function of GAD and its regulation in plants have shown that GAD contains a calmodulin-binding domain in the C-terminal, which enables the activity to be modulated by the Ca²⁺/CaM complex (50, 53). In rice, the C terminus of GAD was shown to have an auto-inhibitory role, as the truncated version resulted in GABA overaccumulation (54). Moreover, we observed that TaGAD1 with deletion of the C-terminal domain exhibited higher glutamate decarboxylase activity than full-length TaGAD1 (Fig. 5, G and J), suggesting that the C terminus of wheat TaGAD1 also has an auto-inhibitory effect. In this work, we further identified a previously unreported interaction module composed of TaNHX2 and TaGAD1 using split firefly luciferase complementation assays, BiFC, split-ubiquitin yeast assays, and Co-IP assays (Fig. 4). Furthermore, we revealed the functions of this module in regulating GABA accumulation by affecting TaGAD1 activity, as simultaneous expression of TaNHX2 and TaGAD1 in both in E. coli and N. benthamiana enhanced TaGAD1 enzymatic activity and GABA accumulation compared with the expression of TaGAD1 alone (Fig. 5, E to J). In addition, we demonstrated that TaNHX2 interacted with the C terminus domain of TaGAD1 but not the N terminus that contained the glutamate decarboxylation domain, suggesting that the interaction of TaNHX2 with the C-terminal domain of TaGAD1 could prevent its auto-inhibitory effect and that this interaction is necessary for the full activation of TaGAD1 activity (Figs. 4 and 5, E to J). In addition, drought stress enhanced the interaction between TaNHX2 and TaGAD1 and was essential for the full activation of TaGAD1 activity (fig. S17). Consistent with this, the GAD2Δ-overexpressing Arabidopsis plants maintained a higher relative leaf water content and survival rate than the wild-type plants after drought treatment (42). Thus, we have revealed a molecular mechanism in which the tonoplast-localized cation exchanger TaNHX2 interacts with glutamate decarboxylase TaGAD1 to control stomatal closure and drought avoidance by regulating GABA accumulation.

Studies have shown that a low cytosolic pH stimulates GABA production by promoting the activity of the GAD protein (53, 59). In the present study, TaGAD1 exhibited higher enzymatic activity at pH 5.7 than at pH 7.0 (fig. S27). As classical cation/H⁺ exchangers, a potential hypothesis is that changes in cytoplasmic pH resulting from TaNHX2 transport activity indirectly affect TaGAD1 activity. The pH condition in the cytoplasm or around the tonoplast membrane may be changed by TaNHX2, causing stronger TaGAD1 activity, more GABA accumulating in a small area, and more precise regulation of ALMT9. However, it is difficult for the present technique using fluorescence probes and cellular imaging to obtain these detailed data. It has also been found that increases in plant GAD activity are dependent upon Ca²⁺/CaM (52, 53, 60). A previous study showed that Arabidopsis calmodulin CaM15 interacted with the NHX protein within the vacuole in a Ca²⁺-dependent manner (61). The putative calmodulin binding site within the C-terminal of TaNHX2 has also been identified, but it remains to be determined whether TaNHX2 interacts with calmodulin proteins. The possible interplay between TaNHX2 and calmodulin and whether the interaction depends on the calmodulin-binding domain of TaGAD1 deserve future research attention.

Crop plants selected for their economic yield need to survive drought stress through mechanisms that maintain crop yield. ABA is known to regulate drought adaptation in plants through stomatal closure and reduced plant growth (57). However, modulating the ABA-induced drought adaptation of plants for a better yield remains a greater challenge because of the potential inadvertent reduction in carbon gain upon stomatal closure and ABA-induced senescence, especially when drought occurs at the reproductive stage (9, 62). In C3 plants, photosynthetic carbon assimilation is generally determined by stomatal control of the ratio of instantaneous rates of photosynthesis and transpiration, among other factors (63, 64). Stomatal closure not only diminishes water loss through transpiration but also reduces CO₂ and nutrient uptake, thus altering metabolic pathways, such as photosynthesis, under drought (9). Compared with phytohormone ABA, GABA’s role appears to be fine-tuning the stomatal aperture (42). The TaNHX2-TaGAD1 hub that modulated stomatal closure under drought stress is largely unresponsive to ABA treatment (fig. S10); therefore, it has a flexible stomatal response to drought stress with less adverse effect on stomatal conductance. Correspondingly, we found that the TaNHX2-OE lines did not affect several important agronomic traits under well-watered conditions but significantly increased TKW and crop yields compared with the wild-type plants under drought stress conditions (Fig. 7). Since the water dissipation of the TaNHX2-OE lines decreased, although the root system architecture and water uptake remained the same as the wild-type plants, the enhanced WUE determined the improved productivity of the TaNHX2-OE lines (Figs. 2 and 7). We speculate that these yield traits were associated with increased grain length and TKW during reproductive development, which may be correlated with the increased WUE conferred by the overexpression of TaNHX2. This finding indicates that TaNHX2 may well be a stomatal modulator of economic significance in response to drought stress. Therefore, TaNHX2 can serve as a target for both genetic engineering and selection for the improvement of wheat drought tolerance. Similar effects of WUE on increased yield have been reported in genetically engineered drought-tolerant rice, maize, and wheat crops (63, 65, 66).

The mechanism of the TaNHX2-TaGAD1 module that affects on stomatal closure under drought stress appears to be conserved across a range of monocot and dicot plants, such as wheat, rice, and Arabidopsis, since we found that the NHX2-GAD1 interaction also occurs in these species (fig. S28). Therefore, insight regarding the NHX2-involved drought resistance pathway is likely to have broad significance. As we found that the genetic manipulation of TaNHX2 gene expression reduced water loss, leading to improved drought performance, our work provides alternative methods for manipulating crop drought stress resilience. The regulatory factors or genes involved in drought resistance identified in this study may also be of great value for genetic improvement in wheat and in other crops.

MATERIALS AND METHODS

Plant materials

All wheat plants were grown in the experimental field of China Agricultural University in Beijing (39°57′N, 116°17′E) and a greenhouse at a relative humidity of 75% and 26/20°C day/night temperatures, with a light intensity of 3000 lux (Master GreenPower CG T 400 W E40, Philips). The surface-sterilized seeds were incubated at 4°C for 3 days in the dark and then exposed to white light at room temperature. Germinated seeds were transplanted into pots. The materials were subjected to different treatments including water deficit treatment, ABA treatment, and GABA treatment. For the mock treatment, the same volume of ddH₂O was sprayed and treated for the same time. For the drought stress phenotype, materials are planted in Pindstrup turf mixed with vermiculite (2:1).

Sequence alignment, homology, and conserved domain analysis

The NHX and GAD homologs were searched against the Ensembl Plants database (http://plants.ensembl.org/) using the BLAST program. Amino acid alignments and the homology analyses were performed using the DNAMAN version of 5.0 (Lynnon Biosoft, Canada) and MEGA 7 software based on observed divergence. Protein domains were searched against National Center for Biotechnology Information (NCBI) Conserved Domain Search (www.ncbi.nlm.nih.gov/) and SMART programs (http://smart.embl-heidelberg.de/).

Production of TaNHX2 overexpression and KO mutants

The seedling leaves cDNA of wheat cultivar Chinese spring was used as the template to amplify the ORF of TaNHX2-D and then inserted into the pMWB122 vector to achieve the Ubi:TaNHX2 construct. For genome editing via CRISPR-Cas9, a single guide RNA (sgRNA) was designed on the basis of the fourth exon of TaNHX2 using the ECRISP Design website (http://crispr.hzau.edu.cn/CRISPR-Cereal/index.php). In essence, a reverse complementary sgRNA sequences with Bsa I cohesive ends were synthesized, then oligonucleotides were annealed and inserted into the terminal vector pBUE411. All binary vectors harboring the desired constructs were transferred into strain EHA105 and transformed into the wheat cultivar Fielder using Agrobacterium-mediated transformation.

Water-deficit drought assays in seedling stage

For phenotypic analysis in seedling stage, the seeds of different genotypes with wild-type plants were transplanted side by side in a cultivation box (32 cm by 24 cm by 9 cm, length × width × depth) that was filled with 1.5 kg of uniformly mixed soil (Pindstrup turf to vermiculite in a ratio of 2:1), respectively, and grown with a 12-hour photoperiod at 20°C in the greenhouse. To make sure the different genotypes being compared were all exposed to same severity of soil drying, the soil water content was recorded by FieldScout TDR 150 Soil Moisture Meter (Spectrum Technologies Inc.) every day after the initiation of water withholding to determine the period of rewatering (fig. S8A). All drought treatment was achieved by stopping irrigation to reach ~3.5% Volumetric Water Content (VWC) for 5 days, and then watering was resumed to allow plants to recover and the number of surviving plants was recorded 5 days later. At least 20 plants of each line were compared in each test, and statistical analyses were based on data obtained from three independent experiments.

For the roots phenotyping, germinated seeds were transplanted into pots (7 cm by 7 cm by 10 cm), which were fully filled with 3-mm vermiculite granules. The control group and treatment group were set up with three replicates, respectively. Drought treatment was achieved by stopping irrigation to reach ~3.5% Relative Soil Water Content (RSWC) for 3 days, and then, the roots were statistically analyzed and recorded phenotypes.

Water-deficit drought assays in the field

For reproductive stages, the plots were designed for the well-watered planting condition and drought treatment conditions according to methods described previously (65). The drought treatment plot was separated by a 1-m deep waterproof layer, and a rain-off shelter was built to prevent plants from receiving irrigation from rainfall. Well-watered plots were irrigated throughout the growth period, while the drought plot was subjected to drought treatment after the first irrigation. The plants that were subjected to drought treatment received approximately 40% of the water that the plants in the sufficiently irrigated plots received.

The photosynthetic parameters were analyzed on the flag leaves at a predetermined date between 0930 and 1100 using a LICOR-6400 CO₂ gas exchange analyzer (LICOR-6400, Lincoln, NE). Then, the photosynthesis (PS) rates and transpiration rates (TRs) in all of the tested genotypes under different conditions were obtained. For the estimation of WUE, calculation was conducted by PSs in relation to TRs (65). Agronomic traits related to yield were analyzed following the maturation of the grains and the harvest.

Subcellular localization

The full-length open reading frame (ORF) of TaNHX2-D, TaGAD1-A, and TaGAD2-A were amplified from Chinese spring genome using specific primers and 2× Hieff Canace Gold PCR Master Mix (catalog no. 10149; Yeasen, Shanghai, China) and inserted into the pCAMBIA1300-GFP vector, respectively. The resulting construct vector and the tonoplast marker vector were co-introduced into wheat protoplasts. Wheat protoplasts transformed with plasmid were cultured overnight, and all the cells were then used to observe the GFP fluorescence accordingly (Carl Zeiss, LSM880). Primer sequences are listed in table S1.

Yeast complementation

The coding sequence of TaNHX2-D was amplified and cloned into the pDR195 and pDR195-GFP vectors, respectively. The constructs were transformed into the Saccharomyces cerevisiae strain Δnhx1 or R5421 (TRK1/TRK2 mutant) by the lithium acetate method, and the cells were grown on minimal mediums (–Ura). Positive transformants of Δnhx1 were diluted and plated onto a Yeast extract Peptone Dextrose medium with Agar (YPDA) medium supplemented with 0, 200, and 300 mM Na⁺ and plated onto an AP (K⁺ deficient) Medium supplemented with 0, 5, 100, or 800 mM KCl, respectively. Positive transformants of R5421 were diluted and plated onto a medium (–Ura) supplemented with 5 and 100 mM KCl, respectively. Primer sequences are listed in table S1.

Oocyte K⁺ measurement assay

The transformed TaNHX2-D or control oocytes, injected with 50 nM K⁺, were prepared for K⁺ measurement assays. For K⁺ release assays, oocytes were washed five times with K⁺-free MBS solution and incubated in K⁺-free MBS for 2 hours. One oocyte incubated in each petri dish represented one sample. The K⁺ content of the bath solution in each petri dish was analyzed using the inductively coupled plasma optical emission spectrometry system (PE-7300DV). K⁺ efflux activities of the oocytes were then calculated.

For the K⁺ measurement of oocytes, the oocytes before and after the K⁺ release assay were collected and washed five times with K⁺-free MBS solution. Oocytes were dried overnight at 80°C. Intracellular cations were extracted from dried oocytes by 4-hour incubation in 4 ml of ddH₂O and followed by filtering using Millipore filters (0.22 mm). Then, the K⁺ concentration was measured as described above.

Stomatal density measurement

Leaves of 2-weeks-old TaNHX2 and TaGAD1 transgenetic lines and wild-type plants were detached and directly fixed by 2.5% glutaraldehyde. The stomatal pictures were obtained using scanning electron microscopy (TM4000, HITACHI, Japan), and then, the stomatal density was calculated.

Stomatal aperture measurement

Leaves of 2-week-old TaNHX2 and TaGAD1 transgenic lines and wild-type plants were collected and incubated in stomatal incubation solution [0.05 M KNO₃/10 mM, MES/50 μM CaCl₂ (pH = 6.15)] under light conditions (300 to 500 μmol·m⁻²·s⁻¹, 25°C) for 1.5 hours or darkness for 1 hour. The leaves were then exposed to air conditions for drought treatment for 10 min or pretreated with GABA. Subsequently, the samples were incubated in the continuous dark, light, light-to-dark, or dark-to-light transition for 30 min or 1 hour. Then, the adaxial sides of the leaf epidermis were peeled off using a cutter blade and leaves were then mounted on slides and observed with an inverted microscope (AE31E, Motic, China), and the stomatal aperture (width/length) was analyzed using the Motic Images Plus 3.0(×64). Two images for each leaf, two leaves for each plant, and five plants for each genotype were used for analysis.

Stomatal conductance measurement

The first fully expanded leaves of 2-week-old TaNHX2 overexpression, KO, and wild-type plants, the widest part of the leaf, were applied to measure stomatal conductance with a portable gas analysis system (SC-1, Decagon Devices Inc.). Three leaves for each plant and three plants for each genotype were used for analysis.

ABA and GABA treatment assays

For ABA treatment assays, the 2.6432 mg of ABA was dissolved in 100 μl of anhydrous ethanol to make 100 mM mother solution, then dissolved in 50 ml of ddH₂O to make 200 μM working solution, and sprayed evenly on the leaf surface of 14-day seedlings 2 hours before infrared imaging. For the mock treatment, the 14-day-old seedlings were sprayed evenly with the same volume of solvent (ddH₂O with a trace amount of anhydrous ethanol) for the same time period.

For GABA treatment assays, the 103.12 mg of GABA was dissolved in 10 ml of ddH₂O to make a 100 mM mother solution, and then 20 and 100 μl of mother solution were dissolved in 10 ml of ddH₂O to make a 0.2 and 1 mM working solution and sprayed evenly on the leaf surface of seedlings 2 hours before infrared imaging. The mock treatments sprayed the seedlings evenly with the same volume of ddH₂O for the same period.

Infrared thermography imaging

The leaf temperature of different TaNHX2 transgenic lines that were subjected to various treatments (in normal conditions, drought treatment, ABA, or GABA treatment) was measured with a portable infrared imager (T40, Teledyne FLIR) and analyzed, accordingly.

Expression localization assays

For expression localization assays, the 35S promoter of pCAMBIA1300-mCherry was replaced by the promotor of TaNHX2-D and TaGAD1-A, respectively, then the full-length ORF of TaNHX2-D and TaGAD1-A were inserted into the vector, respectively. The resulting construct vectors were introduced into N. benthamiana leaves. N. benthamiana leaves transformed with plasmid were cultured for 36 hours and were then used to observe the mCherry fluorescence accordingly (Carl Zeiss, LSM880). Primer sequences are listed in table S1.

RNA extraction and RT-qPCR

Total RNA was extracted from samples using TRIzol (Invitrogen), and first-strand cDNAs (20 μl) were synthesized from 1 μg of starting total RNA using a reverse transcription kit (Vazyme Biotech, R223-01) according to the manufacturer’s instructions. For RT-qPCR, the reaction mixture was composed of the 0.7 μl of cDNA template, 0.2 mM primers, and 5 μl of SYBR Green Mix (Vazyme Biotech, Q121-02/03) in a final volume of 10 μl. Amplification was performed using a CFX96 real-time system (Applied Biosystems). Differences in relative transcript levels were calculated using the 2–ΔCT method relative to wheat TaACTIN (TraesCS5B02G124100). Primer sequences are listed in table S1.

Split-luciferase complementation (LCI) assays

For the split-luciferase assays, the indicated genes were separately fused with the N- or C-terminal of the reporter LUC. The coding sequences of TaNHX2-D and TaGAD1-A were used in LCI assays. The constructs were transformed into the Agrobacterium strain GV3101 and then infiltrated into N. benthamiana leaves. LUC signal was collected after 2 days by using a cooled charge-coupled device camera (NightSHADE LB 985, Berthold Technologies, Bad Wildbad, Germany) after spraying 1 mM d-luciferin on the leaves (Coolaber, CL6928). Primer sequences are listed in table S1.

BiFC assay

For the BiFC assays, the coding sequence of TaNHX2-D was amplified and cloned into the nGFP vector and the coding sequence of TaGAD1-A was amplified and cloned into the cGFP vector. The constructs were transformed into the Agrobacterium strain GV3101 and then infiltrated into N. benthamiana leaves. Forty-eight hours after infiltration, the GFP fluorescence signals for tobacco leaves were imaged with a confocal microscope (LSM880, Carl Zeiss, Heidenheim, Germany). For tobacco protoplasts, TaNHX2-cYFP, nYFP-TaGAD1, and tonoplast marker were co-infiltration in the pavement cells of N. benthamiana, and the tobacco protoplasts were extracted 48 hours after infiltration and then collected merge fluorescence signals with a confocal microscope (LSM880, Carl Zeiss, Heidenheim, Germany). Primer sequences are listed in table S1.

Co-IP assay

For the Co-IP assay, the TaNHX2-D-Myc and TaGAD1-A-GFP fusion constructs were prepared. Then different pairs of specific constructs were cotransformed into the Agrobacterium strain GV3101 and then infiltrated into N. benthamiana leaves. Total proteins were isolated from different positive transformants and incubated with anti-GFP beads at 4°C for 4 to 6 hours with gentle shaking. Then, the corresponding protein was obtained after a series of washing steps and elution processes. Input and eluted protein were then analyzed by Western blot using anti-GFP (mouse, 1:2000; Abclonal, AE012) or anti–c-Myc (mouse, 1:2000; TransGen Biotech, HT101-01). Primer sequences are listed in table S1.

The split-ubiquitin yeast system (DUALmembrane)

The DUALmembrane system was used in accordance with the manufacturer’s protocols. The prey vector pPR3-N or pPR3-N-TaGAD1 and the bait vector pBT3-STE or pBT3-STE-TaNHX2 were cotransformed into the yeast strain NMY51. Yeast cells were spread on SD/-Trp-Leu agar plates and incubated at 29°C for 3 to 5 days after transformation. Colonies grown on SD/-Trp-Leu plates were suspended in SD/-Trp-Leu liquid medium to an optical density of 600 (OD₆₀₀) of 0.6. A 5× dilution series of 10-μl aliquots of cotransformed NMY51 were spotted onto SD/-Trp-Leu and SD/-Leu-Trp-His-Ade agar plates, followed by the examination of LacZ reporter gene with X-Gal as a substrate. Plates were incubated at 29°C for 3 to 5 days.

In vitro pull-down assay

His-TaNHX2-C protein—in combination with GST, GST-TaGAD1, GST-TaGAD1ΔC or GST–TaGAD1-C, respectively—were incubated and immunoprecipitated by GST resin (TransGen Biotech, ProteinIso GST Resin, DP201-01) at 4°C for 2 hours. The mixture was gathered by centrifugation at 500g for 5 min, followed by washing with phosphate-buffered saline buffer five times. Proteins were separated on 10% (w/v) SDS–polyacrylamide gel electrophoresis (SDS-PAGE) and detected with anti-GST (TransGen Biotech, ProteinFind Anti-GST Mouse Monoclonal Antibody, HT601) and anti-His (TransGen Biotech, ProteinFind Anti-His Mouse Monoclonal Antibody, HT501) antibodies. The coding sequences of TaNHX2-D and TaGAD1-A were used in pull-down assays.

GABA accumulation and GAD enzyme activity

Leaves of 1-week-old TaNHX2 overexpression, KO, and wild-type plants with drought treatment or normal growth were detached (0.1 g) for evaluation. The GABA contents were determined by the GABA Assay Kits (Su Zhou Grace Biotechnology Co. Ltd., Su Zhou, China, G1106W), and the GAD activity was determined by the GAD Assay Kits (Su Zhou Grace Biotechnology Co. Ltd., Su Zhou, China, G1102W). For pH dependent of TaGAD1 activity assay, purified recombinant GST-GAD1 protein was incubated in pH 5.7 and 7.0 incubation buffer, respectively, and then, the GAD activity was determined by the GAD Assay Kits (GAD assay kits, Su Zhou Grace Biotechnology Co. Ltd., Su Zhou, China, G1102W).

Measurements of the effect on truncating C terminus of TaGAD1 and TaNHX2 interaction on TaGAD1 enzymatic activity

The coding sequences of TaGAD1 and TaGAD1ΔC were cloned into the pGEX6p-1 to generate TaGAD1-GST and TaGAD1ΔC-GST. The coding sequence of TaNHX2C was also cloned into the pET-32a to produce TaNHX2C-HIS. The constructs were transformed into E. coli BL21 (DE3), different assay combinations were co-incubated to consistent OD₆₀₀, and the recombinant proteins were induced with isopropyl-β-D-thiogalactopyranoside at 4°C for 5 to 6 hours. The following eluted proteins were separated by 8% SDS-PAGE and subjected to immunoblotting with an anti-GST antibody. Primer pairs are listed in table S1.

The coding sequences of AtGAD1 and AtGAD1ΔC were cloned into the pSuper1300-GFP to produce the fusion protein vectors as positive control and then transformed into Agrobacterium GV3101 cells, which infiltrated into N. benthamiana leaves for GAD enzymatic activity assays.

The coding sequences of TaGAD1 and TaGAD1ΔC were cloned into the pSuper1300-GFP to produce the fusion protein TaGAD1-GFP and TaGAD1ΔC-GFP, respectively. The fusion constructs TaGAD1-GFP and TaGAD1ΔC-GFP were transformed into Agrobacterium GV3101 cells and co-infiltrated with TaNHX2-MYC, TaNHX2C-MYC, or MYC into N. benthamiana leaves according to different experimental combinations. After 48- to 72-hour cultivation, these leaves were harvested and ground in liquid nitrogen. Total protein was extracted and the immunoprecipitates were separated in 8% SDS-PAGE and detected by immunoblotting with an anti-GFP antibody. The coding sequences of TaNHX2-D and TaGAD1-A were used in these assays. Primer pairs are listed in table S1.

Supplementary Materials

This PDF file includes:

Figs. S1 to S29

Table S1