Involvement of the Membrane Trafficking Factor PATROL1 in the Salinity Stress Tolerance of Arabidopsis thaliana

Abstract

The Arabidopsis thaliana stomatal complex contains a pair of guard cells surrounded by subsidiary cells, which assist in turgor-driven stomatal movement and receive water and ions. This transport, driven by environmental signals, involves a translocation factor of the plasma membrane proton pump H⁺-ATPase AHA1, PATROL1. In this study, we investigated the responses of PATROL1 to salinity and hyperosmotic stresses. Specifically, we analyzed the effects of 125 mM NaCl or 231 mM mannitol on the cotyledon pavement cell cortexes in transgenic A. thaliana seedlings expressing green fluorescent protein (GFP)-tagged PATROL1. Cells treated with NaCl had few GFP-PATROL1-labeled dot-like structures but contained unusual labeled large bodies and rod-like structures. Cells treated with mannitol had similar large bodies, but not rods, indicating that the rod-like structures form specifically under salinity stress conditions. Dual observations of GFP-PATROL1 and red fluorescent protein (RFP)-tagged AHA1 in stress-treated cells revealed that the latter did not accumulate in the stress-induced GFP-PATROL1 structures, suggesting that the stress-induced GFP-PATROL1 structures are not involved in RFP-AHA1 localization. Additionally, the primary root growth of the patrol1 mutant was more sensitive to NaCl treatment than was that of wild type. Thus, PATROL1 appears to contribute to salinity stress tolerance, possibly by regulating membrane trafficking.

The exocytotic pathway is required for the deposition of plasma membrane and cell wall components during cell growth and differentiation in plants (Wang et al. 2016, Vukašinović et al. 2017). It is also involved in altering these components in response to biotic (Collins et al. 2003, Kwon et al. 2008) or abiotic (Zhu et al. 2002, Drakakaki et al. 2012) stresses. Recent studies on the Munc13-like protein H⁺-ATPase translocation control 1 (PATROL1) revealed that the exocytotic pathway may contribute to stomatal movement in response to various environmental cues (Hashimoto-Sugimoto et al. 2013, Higaki et al. 2014). Additionally, PATROL1 may affect stomatal opening via the localization of the H⁺-ATPase AHA1 in the plasma membrane, in which AHA1 is activated in response to various signals, such as light, and induces guard cell plasma membrane hyperpolarization (Higaki et al. 2014). The AHA1-mediated hyperpolarization triggers a potassium influx into guard cells, resulting in the stomatal opening (Higaki et al. 2014). Before PATROL1 was identified, the mechanism of activation of plasma membrane H⁺-ATPases was a major topic of interest among researchers focused on the mechanisms underlying stomatal opening. The discovery of PATROL1 indicated that the membrane trafficking of AHA1 is important for controlling stomatal movement.

PATROL1 was originally identified as the gene associated with the defective stomatal opening of an Arabidopsis thaliana mutant in response to a low CO₂ concentration (Hashimoto-Sugimoto et al. 2013). PATROL1 has a MUN domain that is crucial for the synaptic vesicle priming of the Munc13 protein in animal neurons (Augustin et al. 1999). GFP-PATROL1 localizes on dot-like structures (GFP-PATROL1 dots) just beneath plasma membranes (Hashimoto-Sugimoto et al. 2013). These GFP-PATROL1 dots are stained by the endocytic tracer FM4-64 in leaf epidermal cells of A. thaliana (Hashimoto-Sugimoto et al. 2013), as is Munc13 in mouse brain cells (Kalla et al. 2006). Time-sequential imaging using a variable-angle epifluorescence microscope indicated that GFP-PATROL1 dots appear and remain at the same site for 2–10 s despite the existence of cytoplasmic streaming, suggesting that the dots are tethered to the plasma membranes and participate in membrane fusion (Higaki et al. 2014, Higaki 2015). A previous study on the patrol1 mutant revealed the abnormal internalization of the plasma membrane H⁺-ATPase AHA1, which triggers guard cell plasma membrane hyperpolarization and the subsequent potassium influx into the guard cells of opening stomata (Higaki et al. 2014). Interestingly, the other plasma membrane proteins involved in stomatal movement, including the potassium channel KAT1, the slow anion channel SLAC1, and aquaporin PIP2a, are localized normally, even in the patrol1 mutant (Hashimoto-Sugimoto et al. 2013). These findings imply that PATROL1 has specific functions related to the translocation of AHA1 to the plasma membrane (Hashimoto-Sugimoto et al. 2013). Most of the molecular mechanisms underlying the localization of the other stomatal movement-related plasma membrane proteins are unknown. Additionally, PATROL1 is ubiquitously expressed, suggesting that it also affects processes other than stomatal movement. The possible involvement of PATROL1 in the delivery of the cellulose synthase complex to the plasma membrane in hypocotyl cells has been reported (Zhu et al. 2018).

Using time-lapse confocal imaging, the dynamic behaviors of GFP-PATROL1 dots in response to short-term (up to 30 min) drought treatments have been observed. The dot density increases in guard cells in response to leaf detachment or an abscisic acid treatment (Hashimoto-Sugimoto et al. 2013, Higaki et al. 2014). In contrast, a light treatment decreases the density of GFP-PATROL1 dots in guard cells (Hashimoto-Sugimoto et al. 2013). These observations imply that the GFP-PATROL1 dot density dynamically changes with stomatal movement. Moreover, subsidiary cell responses differ from those of guard cells, with the GFP-PATROL1 dot density decreasing and increasing when stomata open and close, respectively (Higaki et al. 2014). These observations suggest that the cooperative dynamics of GFP-PATROL1 dots contribute to the smooth transport of ions and water between guard and subsidiary cells because of AHA1 localization during stomatal movement in response to environmental cues (Higaki et al. 2014). However, the long-term responses of PATROL1 to abiotic stresses have not been examined.

We herein report that a long-term (24 h) salinity stress treatment induced the formation of unusual rod-like structures labeled with GFP-PATROL1. Furthermore, we found that the patrol1 mutant exhibited decreased tolerance to salinity stress. The results of our cell biology and physiological analyses suggest that PATROL1 is involved in plant salinity tolerance.

Materials and methods

Plant materials and growth conditions

All A. thaliana plants used in this study were derived from a wild-type line with the Col-0 background. Using a thermal imaging screening system to detect transpiration, the patrol1 mutant was previously isolated as a CO₂-mediated stomatal movement-deficient mutant from EMS-mutagenized Col M2 seeds purchased from Lehle Seeds (Round Rock, TX, USA) (Hashimoto-Sugimoto et al. 2013). The patrol1 mutant has a point mutation that introduces a stop codon, and the resulting substantial decrease in the PATROL1 (At5g06970) transcript level was confirmed by RT-PCR analysis (Hashimoto-Sugimoto et al. 2013). To visualize the intracellular localization of PATROL1, the patrol1 mutant was transformed with the GFP-tagged PATROL1 construct under the control of the Cauliflower mosaic virus (CaMV) 35S promoter by Agrobacterium tumefaciens-mediated transformation (Hashimoto-Sugimoto et al. 2013). The PATROL1 transcript level in the patrol1/GFP-PATROL1 line was revealed to be similar to that of the wild-type Col-0 line by RT-PCR analysis (Hashimoto-Sugimoto et al. 2013). Moreover, the stomatal phenotypes were recovered in the patrol1/GFP-PATROL1 line (Hashimoto-Sugimoto et al. 2013). In this study, homozygous T3 generation patrol1/GFP-PATROL1 plants were used to visualize the PATROL1 intracellular localization and evaluate salinity stress tolerance in a rescue line. The transgenic A. thaliana plants expressing GFP-PATROL1 and RFP-AHA1 in the patrol1 mutant background, both under the control of the CaMV 35S promoter, were also analyzed. This dual-labeling line was established by inserting the RFP-AHA1 construct into the GFP-PATROL1 line via A. tumefaciens-mediated transformation (Hashimoto-Sugimoto et al. 2013). The homozygous T3 generation was used in this study. Sterilized seeds of the transgenic lines were sown on half-strength Murashige–Skoog (MS) gellan gum medium in a square plastic plate (140×100×14.5 mm) and incubated in a growth chamber at 23.5°C with a 16-h light/8-h dark photoperiod (86.2 µmol m⁻² s⁻¹ light-emitting diode for plant growth) (Plantflec LH-241PFP-S, NK system, Tokyo, Japan).

Stress treatment

As controls, we used cotyledons from 6–7-day-old seedlings. For the salinity or hyperosmotic stress treatment, 5-day-old seedlings were transferred to half-strength MS gellan gum medium supplemented with 125 mM NaCl or 231 mM mannitol, respectively, and then analyzed with a confocal microscope 24 h later (Dou et al. 2018).

Confocal microscopy

Cotyledons were sampled from the seedlings. They were placed on glass slides (76×26 mm) (S1112; Matsunami, Osaka, Japan) and mounted in half-strength MS liquid medium with a cover glass (18×18 mm) (Matsunami, Osaka, Japan). The pavement cells and guard cells of the cotyledon abaxial sides were observed by spinning disk confocal microscopy. For time-sequential observations of GFP-PATROL1, we used a fluorescence microscope (IX-70; Olympus, Tokyo, Japan) equipped with a spinning disk confocal laser scanning unit (CSU-X1; Yokogawa, Tokyo, Japan) and a cooled CCD camera (CoolSNAP HQ2; Photometrics, Arizona, USA). For dual observations of GFP-PATROL1 and RFP-AHA1, we used a fluorescence microscope (IX-70; Olympus, Tokyo, Japan) equipped with a spinning disk confocal laser scanning unit (CSU-W1; Yokogawa, Tokyo, Japan) and a complementary metal-oxide semiconductor camera (Zyla; Andor, Belfast, UK). We used a PlanApo×60/1.10 objective lens (Olympus, Tokyo, Japan). Additionally, GFP and RFP were excited at 488 nm and 561 nm, respectively, with emission wavelengths of 510–550 nm and 624–640 nm, respectively, through a band-pass filter. All microscopy and image acquisition settings were fixed so that the images could be compared.

Microscopy image processing and analysis

All microscopy images were processed using the ImageJ/Fiji software (Schneider et al. 2012). The motility of the stress-induced intracellular structures was evaluated using the ‘Temporal-Color Code’ ImageJ plugin (Fiji–Image–Hyperstacks–Temporal-Color Code). All image processing parameters were fixed so that the images could be compared.

Root growth assay

The wild-type (Col-0), patrol1 mutant, and the rescue line expressing GFP-PATROL1 (Hashimoto-Sugimoto et al. 2013) were treated with and without 125 mM NaCl or 231 mM mannitol as described above. The plants in the plates were time-sequentially photographed using a digital single-lens reflex camera (EOS Kiss X9; Canon, Tokyo, Japan). The primary root length of the living seedlings, which grew from 3 to 5 days after treatments, was manually measured using the ‘Freehand Line selection tool,’ the ‘ROI manager,’ and the ‘Measurement’ function in ImageJ/Fiji software (Schneider et al. 2012). The data were analyzed using Tukey–Kramer tests and Mann–Whitney U-tests with R software (version 3.6.1; https://www.r-project.org/).

Results

Formation of unusual GFP-PATROL1 structures under salinity or hyperosmotic stress conditions

To examine the behavior of GFP-PATROL1 in cells under salinity or hyperosmotic stress conditions, we analyzed the cortexes of cotyledon pavement cells in GFP-PATROL1-expressing transgenic A. thaliana seedlings treated with 125 mM NaCl or 231 mM mannitol for 24 h. In the untreated control seedlings, many small GFP-PATROL1 dots were observed against the cytosolic background that often negatively visualized organelles (Fig. 1A), as previously reported (Hashimoto-Sugimoto et al. 2013, Higaki et al. 2014). However, in cells treated with 125 mM NaCl, normal GFP-PATROL1 dots were rarely observed (Fig. 1B). Instead, we found unusual GFP-PATROL1-labeled structures (GFP-PATROL1 bodies) (Fig. 1B, left) that were larger than normal GFP-PATROL1 dots (Fig. 1A). We also found rod-like GFP-PATROL1-labeled structures (GFP-PATROL1 rods) (Fig. 1B, right). In the cells treated with 231 mM mannitol (having the same osmolarity as 125 mM NaCl), similar GFP-PATROL1 bodies were observed (Fig. 1C, left), but not GFP-PATROL1 rods, suggesting that these GFP-PATROL1 rods are specific to salinity stress. Additionally, we found large spherical bodies labeled with GFP-PATROL1 (GFP-PATROL1 spherical bodies) (Fig. 1C, right).

Fig. 1. Effects of salinity or hyperosmotic stress on GFP-PATROL1 localization. The cortexes of cotyledon pavement cells from 6–7-day-old A. thaliana seedlings treated without (A) or with 125 mM NaCl (B) or 231 mM mannitol (C) for 24 h are shown. Broken lines indicate the cell periphery. Scale bars=10 µm.

Motility of the stress-induced intracellular structures labeled with GFP-PATROL1

We next determined the motility of the GFP-PATROL1-labeled intracellular structures based on time-sequential observations. In the control, GFP-PATROL1 dots remained at the same position near the plasma membrane for 2–10 s and then disappeared (Fig. 2A), as previously reported (Higaki et al. 2014). Unlike the GFP-PATROL1 dots, the stress-induced structures did not appear or disappear during at least 30 s of time-sequential observations (Fig. 2B, C). Relatively few positional or morphological changes to GFP-PATROL1 bodies were observed over 30 s in cells subjected to salinity or hyperosmotic stress, whereas the cytoplasmic background moved, possibly because of cytoplasmic streaming (Fig. 2B, C, top). Although salinity stress-induced movement of GFP-PATROL1 rods, their motility was unremarkable (Fig. 2B, bottom). However, the hyperosmotic stress-induced GFP-PATROL1 spherical bodies moved dramatically, possibly because of cytoplasmic streaming (Fig. 2C, bottom).

Fig. 2. Motility of salinity or hyperosmotic stress-induced GFP-PATROL1-labeled intracellular structures in cotyledon pavement cells from 6–7-day-old A thaliana. The time-sequential images of the cells treated without (A) or with 125 mM NaCl (B) or 231 mM mannitol (C) were captured for 30 s at 1 s intervals. Snapshots (left) and temporal color-coded images (right) are shown. The colored bar indicates the timing of the signal detection. Scale bars=10 µm.

Dual observations of GFP-PATROL1 and RFP-AHA1

PATROL1 reportedly regulates the plasma membrane localization of the H⁺-ATPase AHA1, which contributes to stomatal opening (Hashimoto-Sugimoto et al. 2013). In plasmolyzed guard cells exposed to 530 mOsM, RFP-tagged AHA1 (RFP-AHA1) and GFP-PATROL1 colocalize at the tips of Hechtian strands within the cells (Hashimoto-Sugimoto et al. 2013). Therefore, we inspected the colocalization of GFP-PATROL1 and RFP-AHA1 in the stress-treated pavement cells. In the control, GFP-PATROL1 localized on the dots near the plasma membranes and RFP-AHA1 evenly localized in plasma membranes in addition to a few vesicle-like structures (Fig. 3A), as reported for guard cells (Hashimoto-Sugimoto et al. 2013). In the stress-treated cells, the localization of RFP-AHA1 did not appear to be significantly altered (Fig. 3B, C). RFP-AHA1 did not accumulate in the stress-induced GFP-labeled structures (i.e., the GFP-PATROL1 bodies, rods, and spherical bodies) (Fig. 3B, C), suggesting that the stress-induced GFP-PATROL1 structures were not directly linked with the regulation of AHA1 localization.

Fig. 3. Dual observation of GFP-PATROL1 and RFP-AHA1 in A. thaliana cotyledon pavement cells treated with salinity or hyperosmotic stress. Images of GFP-PATROL1 (left) and RFP-AHA1 (middle) in the cells treated without (A) or with 125 mM NaCl (B) or 231 mM mannitol (C) are presented. Merged images are on the right. Arrows and arrowheads indicate GFP-PATROL1 bodies and RFP-AHA1-labeled vesicle-like structures, respectively. To focus on the GFP-PATROL1 spherical bodies, the confocal image shows the region approximately 1 µm inside the cell cortex (C, bottom). Scale bars=10 µm.

We also checked the localization of GFP-PATROL1 and RFP-AHA1 in guard cells (Fig. 4). In the control, the GFP-PATROL1 dots were observed on the plasma membranes that were labeled with RFP-AHA1 (Fig. 4A), as previously reported (Hashimoto-Sugimoto et al. 2013). In both stress-treated guard cells, the GFP-PATROL1 dots were rarely observed (Fig. 4B, C), like in stress-treated pavement cells (Figs. 1B, C, 3B, C). However, unlike the pavement cells, the stress-treated guard cells contained few GFP-PATROL1 bodies, rods, and spherical bodies (Fig. 4B, C). In addition, RFP-AHA1 localization was not significantly affected by salinity or hyperosmotic stress (Fig. 4B, C), like in pavement cells (Fig. 3).

Fig. 4. Dual observation of GFP-PATROL1 and RFP-AHA1 in A. thaliana cotyledon guard cells treated with salinity or hyperosmotic stress. Images of GFP-PATROL1 (left) and RFP-AHA1 (middle) in the cells treated without (A) or with 125 mM NaCl (B) or 231 mM mannitol (C) are presented. Merged images are on the right. Scale bars=5 µm.

Effects of salinity stress on root growth of the patrol1 mutant

To examine the roles of PATROL1 in salinity tolerance, we monitored the seedling root growth of the wild type (Col-0), patrol1 mutant, and rescue line expressing GFP-PATROL1 (Hashimoto-Sugimoto et al. 2013) on MS medium with and without 125 mM NaCl or 231 mM mannitol supplementation (Fig. 5). At 5 days after treatments, the primary roots under both salinity and hyperosmotic stress conditions were significantly shorter than those under control conditions (without hyperosmotic stress) in all lines (p＜0.01, Tukey–Kramer test, N=9–15) (Figs. 5A, 6A), indicating that hyperosmotic stress suppressed root growth in a PATROL1-independent manner. After 11 days, the primary roots of the patrol1 mutant on the NaCl-containing medium were significantly shorter than those of the mutant on the mannitol-containing MS medium [p=0.048 (11 days), 0.036 (13 days), 0.026 (15 days), U-test, N=10–15] (Figs. 5B, 6C), whereas there were no significant differences in primary root length between the wild type and rescue lines growing under salinity and hyperosmotic stress conditions (p＞0.17, U-test, N=9–14) (Figs. 5B, 6B, D). In addition, as an indicator of inhibition of primary root growth by salinity stress, the ratio of the length of the primary root in the 125 mM NaCl treatment to the average value of the length of the primary root in the 231 mM mannitol treatment () was determined (Fig. 6E). The ratio was significantly lower in patrol1 than in wild type (Col-0) or the rescue line, suggesting that the patrol1 mutant was sensitive to salinity stress (Fig. 6E).

Fig. 5. Effects of salinity and hyperosmotic stress treatments on the seedling root growth of the wild-type A. thaliana (Col-0), patrol1 mutant, and rescue line (patrol1/GFP-PATROL1). Representative photographs of plates at 5 (A) and 15 days after transfer (B) to MS medium supplemented with 125 mM NaCl (middle) or 231 mM mannitol (bottom) are shown. Controls were grown in medium without NaCl or mannitol supplementation (top). Scale bars=10 mm.

Fig. 6. Effects of the patrol1 mutation on primary root growth under salinity and hyperosmotic stress conditions. (A) Seedling primary root length of the wild-type A. thaliana Col-0, patrol1 mutant, and rescue line (patrol1/GFP-PATROL1) at 5 days after treatments without (white column; control) and with 125 mM NaCl (black column; salinity stress) or 231 mM mannitol (gray column; hyperosmotic stress). Data are the mean±standard error. Statistical significance was determined by Tukey–Kramer test (p＜0.01). N=9–15. (B–D) Seedling primary root length of Col-0 (B), patrol1 (C), and patrol1/GFP-PATROL1 (D) at 11, 13, and 15 days after treatments with 125 mM NaCl (black column; salinity stress) or 231 mM mannitol (gray column; hyperosmotic stress). Data are the mean±standard error. Statistical significance was determined by Mann–Whitney U-tests (* p＜0.05). N=9–15. (E) Quantitative evaluation of salinity stress-induced inhibition of primary root growth. As an indicator of root growth inhibition, primary root length in the 125 mM NaCl treatment divided by the mean primary root length in the 231 mM mannitol treatment () was calculated for 11 to 15-day-old seedlings. Statistical significance was determined by Tukey–Kramer test (p＜0.04). N=12–15.

Discussion

In this study, we found unusual GFP-PATROL1-labeled intracellular structures in cells subjected to either salinity or hyperosmotic stress. Normal GFP-PATROL1 dots, which localize just beneath plasma membranes and remain at the same position for 2–10 s (Figs. 1A, 2A, 3A, 4A) (Higaki et al. 2014), were undetectable in cells under salinity or hyperosmotic stress conditions, suggesting that these stresses attenuate the PATROL1-mediated AHA1 localization to plasma membranes (Figs. 1B, C, 2B, C, 3B, C, 4B, C). This is reasonable because endocytosis and exocytosis are up-and down-regulated, respectively, in response to hyperosmotic stress (Zwiewka et al. 2015). Instead of GFP-PATROL1 dots, GFP-PATROL1 bodies were observed in cells treated with 231 mM mannitol (Fig. 1C, left). They had low motility at the cell cortex (Fig. 2C, top), implying the membranous structures were tethered to the plasma membranes, as previously suggested for the normal GFP-PATROL1 dots (Higaki et al. 2014). Additionally, the larger GFP-PATROL1 spherical bodies moved rapidly, possibly because of cytoplasmic streaming (Fig. 2C, bottom). These observations support the hypothesis that a long-term hyperosmotic treatment inhibits exocytosis and causes the GFP-PATROL1-labeled plasma membrane-anchored vesicles to aggregate, resulting in immobile GFP-PATROL1 bodies despite cytoplasmic streaming. These bodies finally detached from the plasma membranes, leading to the formation of motile spherical bodies. To further test this hypothesis, additional studies including ultrastructural observations of the membrane structures with electron microscopy are required.

The GFP-PATROL1 bodies were commonly observed in cells treated with 125 mM NaCl (Fig. 1B, left), as in the cells exposed to hyperosmotic stress, suggesting that the bodies formed in response to hyperosmotic stress. However, in the NaCl-treated cells, GFP-PATROL1 rods were observed, but not GFP-PATROL1 spherical bodies (Fig. 1B, right). Thus, the GFP-PATROL1 rods formed specifically in response to salinity stress. Unfortunately, the formation mechanisms remain unclear. Our results indicated that RFP-AHA1 did not accumulate in the stress-induced GFP-PATROL1-labeled intracellular structures (Fig. 3), reflecting the limited involvement of AHA1 in the stress-induced structures. In addition, we rarely observed stress-induced GFP-PATROL1-labeled intracellular structures in guard cells, suggesting that the formation of these structures is cell type-specific (Fig. 4). Further studies involving multi-color imaging of other membrane trafficking or cytoskeleton markers are needed to clarify the formation mechanisms. However, the primary root growth of the patrol1 mutant was sensitive to salinity stress (Figs. 5, 6). These results suggest that PATROL1 contributes to salinity stress tolerance, possibly by regulating membrane trafficking through the formation of rod-like membranous structures. Unfortunately, there is no available information regarding the key molecules involved in PATROL1-mediated salinity stress tolerance, but plasma membrane proteins confirmed to be involved in salinity stress tolerance, including the SOS1 Na⁺/H⁺ antiporter (Shi et al. 2002), are possible candidates. Additional research on the localization of candidate proteins may provide clues about the molecular mechanisms underlying PATROL1-mediated plant salt tolerance. Furthermore, although this study focused on primary root growth, more detailed studies on the various harmful effects of salinity on plants, such as inhibition of shoot growth and plant death, will help to clarify the role of PATROL1 in plant salt tolerance.

Acknowledgments

We thank Dr. Mimi Hashimoto-Sugimoto of Nagoya University and Prof. Seiichiro Hasezawa of The University of Tokyo for their helpful suggestions. This work was supported by the Japan Society for the Promotion of Science (JSPS) KAKENHI to T.H. (18H05492 and 20H03289). We thank Edanz Group (https://en-author-services.edanz.com/ac) for editing a draft of this manuscript and helping to draft the abstract.

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