Abstract In Arabidopsis thaliana, the endosomal-localized Na+/H+ antiporters NHX5 and NHX6 regulate ion and pH homeostasis and are important for plant growth and development. However, the mechanism by which these endosomal NHXs function in plant development is not well understood. Auxin modulates plant growth and development through the formation of concentration gradients in plant tissue to control cell division and expansion. Here, we identified a role for NHX5 and NHX6 in the establishment and maintenance of auxin gradients in embryo and root tissues. We observed developmental impairment and abnormal cell division in embryo and root tissues in the double knockout nhx5 nhx6, consistent with these tissues showing high expression of NHX5 and NHX6. Through confocal microscopy imaging with the DR5::GFP auxin reporter, we identify defects in the perception, accumulation and redistribution of auxin in nhx5 nhx6 cells. Furthermore, we find that the steady-state levels of the PIN-FORMED (PIN) auxin efflux carriers PIN1 and PIN2 are reduced in nhx5 nhx6 root cells. Our results demonstrate that NHX5 and NHX6 function in auxin-mediated plant development by maintaining PIN abundance at the plasma membrane, and provide new insight into the regulation of plant development by endosomal NHX antiporters. Introduction The ability to regulate cellular ion and pH homeostasis is a basic requirement of all living organisms, particularly under abiotic stress. In part this is achieved through a family of specialized Na+/H+ exchangers (NHXs) which utilize an electrochemical gradient to transport an Na+ or K+ ion for H+ across a membrane (Blumwald 2000). NHXs are found in all eukaryotes and in addition to their roles in pH and ion homeostasis, also function in diverse processes including the regulation of cell shape and volume, vesicular trafficking, protein sorting and cellular stress responses (Brett et al. 2005a, Orlowski and Grinstein 2007, Ohgaki et al. 2011, Bassil et al. 2012, Chanroj et al. 2012). In Arabidopsis thaliana, two intracellular NHXs, NHX5 and NHX6, act redundantly, and have a significant role in plant growth and development (Bassil et al. 2011). Double knockout nhx5 nhx6 plants display growth abnormalities, including a dwarf phenotype, and reduced shoot and root growth due to slowed cell proliferation (Bassil et al. 2011). All sequenced plant species examined to date have NHX5 and NHX6 orthologs, demonstrating the importance of endosomal NHXs for plant function (Chanroj et al. 2012, Ford et al. 2012). Recent evidence has implicated plant NHXs in subcellular protein trafficking. In A. thaliana, NHX5 and NHX6 localize to the Golgi, trans-Golgi network/early endosome (TGN/EE) and the multivesicular body/pre-vacuolar compartment (MVB/PVC) where they regulate lumenal pH homeostasis (Bassil et al. 2011, Reguera et al. 2015). nhx5 nhx6 mutants fail to sort a vacuolar-targeted yeast carboxypeptidase Y (CPY)–green fluorescent protein (GFP) fusion protein correctly, and have delayed trafficking of the endocytic tracer dye FM4-64 to the vacuole, demonstrating a clear role for NHX5 and NHX6 in vacuolar trafficking (Bassil et al. 2011). Furthermore, NHX5 and NHX6 are required for normal processing and transport of seed storage proteins to the protein storage vacuole in embryos (Ashnest et al. 2015, Reguera et al. 2015). Similar vacuolar trafficking defects have been observed in the yeast nhx1 mutant (Bowers et al. 2000, Brett et al. 2005b), and in animal cells where the mammalian ortholog NHE6 and NHE8 genes have been silenced by RNA interference (RNAi) (Lawrence et al. 2010, Ohgaki et al. 2010), demonstrating that this class of NHX proteins has a conserved role in subcellular trafficking across multiple phyla. The strong reported expression of NHX6 in developing embryos (Ashnest et al. 2015), along with inhibited root and shoot growth in nhx5 nhx6 (Bassil et al. 2011), implicate a role for NHX5 and NHX6 in both embryonic and post-embryonic plant development. The phytohormone auxin is vital for normal plant development, and regulates essential processes including cell patterning, meristem identity and tropisms (Petrásek and Friml 2009). The asymmetric distribution of auxin in cells is generated through polar auxin transport, which drives the generation of defined auxin gradients and maxima in plant tissues. Auxin gradients are formed and maintained through directional intercellular auxin transport mediated by the auxin influx AUX1/LIKE-AUX1 (AUX1/LAX) and auxin efflux PIN FORMED (PIN) families of carrier proteins (Marchant et al. 2002, Wisniewska et al. 2006, Friml 2010). Auxin gradients generated through PIN and AUX carriers are essential for controlling cell division and differentiation in the root tip and developing lateral root primordia (LRPs), as well as during embryogenesis and shoot organogenesis (Möller and Weijers 2009, Petrásek and Friml 2009). Maintenance of AUX and PIN polarity at the plasma membrane is essential for directional intercellular auxin transport, and relies on the dynamic trafficking of these carriers through multiple subcellular pathways. PINs undergo constitutive endocytosis from the plasma membrane to the TGN/EE (Geldner et al. 2001, Dhonukshe et al. 2007), which acts as the hub of endocytic and secretory traffic in plant cells (Viotti et al. 2010). The polar recycling of PINs to the basal plasma membrane from the TGN/EE is mediated by a pathway involving the brefeldin A (BFA)-sensitive ARF guanine-nucleotide exchange factor (ARF-GEF) GNOM (Geldner et al. 2003). Conversely, apical trafficking of PIN2 and AUX1 occurs through a GNOM-independent pathway distinct from that of basal trafficking of PIN1 (Kleine-Vehn et al. 2006, Robert et al. 2008), highlighting that distinct basal and apical transport routes exist. PIN carriers also undergo vacuolar degradation which assists in controlling their abundance at the plasma membrane. Internalized PIN proteins targeted for degradation are trafficked to the MVB/PVC for sorting into endosomal intraluminal vesicles, before final delivery to the lytic vacuole for degradation (Kleine-Vehn et al. 2008, Spitzer et al. 2009). This vacuolar degradation pathway is regulated by the retromer complex, which is thought to enable the retrieval of PIN proteins from the MVB/PVC, thus maintaining PIN polarity and abundance at the plasma membrane (Jaillais et al. 2006, Kleine-Vehn et al. 2008, Nodzyński et al. 2013). We recently reported that the C-terminal cytosolic tail of NHX6 interacts with the retromer complex component SORTING NEXIN1 (SNX1), and NHX5 and SNX1 strongly co-localize in endosomal compartments in root cells, suggesting that NHX5 and NHX6 may be involved in retromer-mediated PIN recycling (Ashnest et al. 2015). Although phenotypic evidence supports a role for NHX5 and NHX6 in embryo and root development, the underlying mechanism behind their contribution has not been investigated. Here, through promoter–reporter assays, we show that NHX6 is expressed in the primary root and developing lateral roots. Furthermore, we identify defects in embryo and LRP development in nhx5 nhx6 mutants, and link these defects to a disruption in auxin signaling due to reduced PIN1 and PIN2 protein abundance at the plasma membrane. Finally, pharmacological interference of PIN1 and PIN2 recycling indicates that NHX5 and NHX6 function independently of polar PIN recycling pathways, and instead may be implicated in retromer-controlled PIN degradation. These findings demonstrate that NHX5 and NHX6 are important for functional auxin-mediated plant development. Results NHX6 is expressed in primary and developing lateral roots NHX5 and NHX6 have been reported to be expressed in whole-root tissue (Yokoi et al. 2002, Bassil et al. 2011). To examine the expression of NHX6 in roots more closely, we created a β-glucuronidase (GUS) reporter construct driven by 3 kb of sequence upstream of the start codon of NHX6 (Ashnest et al. 2015). In the primary root, GUS activity was strongly observed in the apical meristematic zone, stem cell niche and columella cells in multiple independent lines (Fig. 1A). This strong NHX6 expression in root tip columella cells is consistent with findings from microarray expression profiles (Brady et al. 2007). GUS activity was also detected in the central primordia cells of early stage (II–V) developing LRPs, ubiquitously in late stage (VII) primordia and confined to the vascular tissue in emerging and mature lateral roots (Fig. 1A). A similar construct containing up to 3 kb of the NHX5 promoter failed to produce any GUS staining, suggesting that additional elements may be required for NHX5 expression in the root. Fig. 1 View largeDownload slide Lateral root development and emergence are inhibited in nhx5 nhx6. (A) pNHX6::GUS expression pattern in the primary root tip and lateral root primordia (stages denoted by roman numerals; Em, emerged; LR, lateral root). (B) Confocal images of lateral root primordia expressing AUX1–YFP through early and late developmental stages. (C–E) Quantification of mean root length (C), lateral root primordia density (D) and distribution of lateral root primordia (E) from the primary root of 15-day-old seedlings expressing AUX1–YFP. For each seedling, developing lateral root primordia along the primary root were staged as per Malamy and Benfey (1997), and the proportion of each stage was quantified from the total amount of initiated primordia. Data are means ± SEM of >15 seedlings. Student’s t-test; *P < 0.05; **P < 0.01. Scale bars = 20 μm. Fig. 1 View largeDownload slide Lateral root development and emergence are inhibited in nhx5 nhx6. (A) pNHX6::GUS expression pattern in the primary root tip and lateral root primordia (stages denoted by roman numerals; Em, emerged; LR, lateral root). (B) Confocal images of lateral root primordia expressing AUX1–YFP through early and late developmental stages. (C–E) Quantification of mean root length (C), lateral root primordia density (D) and distribution of lateral root primordia (E) from the primary root of 15-day-old seedlings expressing AUX1–YFP. For each seedling, developing lateral root primordia along the primary root were staged as per Malamy and Benfey (1997), and the proportion of each stage was quantified from the total amount of initiated primordia. Data are means ± SEM of >15 seedlings. Student’s t-test; *P < 0.05; **P < 0.01. Scale bars = 20 μm. nhx5 nhx6 mutants exhibit disturbed lateral root development The double knockout nhx5-1 nhx6-1 was originally characterized with a dwarf plant phenotype and reduced primary root growth (Bassil et al. 2011). To identify the cause of this defect, we examined nhx5 nhx6 roots in detail using a second allelic combination which we reported shows identical seed storage protein phenotypes to nhx5-1 nhx6-1 (nhx5-2 nhx6-3; see Materials and Methods; Ashnest et al. 2015). Consistent with previous reports, the primary root length of nhx5-2 nhx6-3 seedlings was significantly reduced, with a large proportion of seedlings exhibiting arrested development shortly after cotyledon expansion (Fig. 1C;Supplementary Fig. S1A; Bassil et al. 2011). These phenotypes could be completely rescued by growing nhx5 nhx6 seedlings on medium supplemented with sucrose (Supplementary Fig. S1), a phenotype shared by mutants with defects in vacuolar trafficking (Shimada et al. 2006, Kleine-Vehn et al. 2008, Feraru et al. 2010). We next determined if LRP development was affected in nhx5 nhx6 mutants. Lateral roots develop through a series of highly co-ordinated cell divisions before eventually emerging from the primary root (Malamy and Benfey 1997, Vilches-Barro and Maizel 2015). We observed and quantified LRP development from initiation through to emergence by crossing the plasma membrane-localized pAUX1-AUX1-YFP (yellow fluorescent protein) marker into nhx5 nhx6. No gross morphological defects were present in LRPs from nhx5 nhx6 seedlings, indicating that their cellular patterning and organization are not dramatically affected (Fig. 1B). LRP density was increased in nhx5 nhx6 plants (Fig. 1D), consistent with a reduction in root length due to slowed cell expansion (Bassil et al. 2011), but normal LRP initiation. However, we occasionally observed the formation of ectopic LRPs (Supplementary Fig. S2), suggesting that the distribution of LRP initiation events along the primary root may be disrupted. Moreover, nhx5 nhx6 mutants showed an increase of early stage LRPs, but a decrease in late stage LRPs (Fig. 1E), indicating a delay to, or inhibition of, LRP development. An estradiol-inducible NHX6–GFP rescue construct could partially restore both primary root elongation and lateral root emergence in nhx5 nhx6 seedlings germinated and grown in the presence of the inducer, confirming that knock-out of endosomal NHXs is responsible for these phenotypes (Supplementary Fig. S4). Embryo patterning is altered in nhx5 nhx6 We previously reported NHX6 promoter–GUS activity in developing and mature stage embryos (Ashnest et al. 2015), indicating that NHX6 is expressed during embryo development. We thus investigated whether nhx5 nhx6 mutants may display abnormalities during embryogenesis. Morphological assessment of cleared embryos revealed low penetrance cell patterning defects throughout globular, trianglular and heart stages of development in nhx5 nhx6 (Fig. 2). Defects were apparent in the basal cell region corresponding to the embryonic root, along with cell patterning anomalies in the tips of the incipient cotyledons. Together with primary and lateral root data, these findings indicate that NHX5 and NHX6 function in growth and development of both the embryo and root, two tissues that are critically associated with auxin patterning. Fig. 2 View largeDownload slide NHX5 and NHX6 are involved in auxin-mediated embryo development. Morphology of cleared embryos from globular to heart stage. Arrows indicate the position and region of aberrant cell division and patterning in nhx5 nhx6 embryos compared with the wild type. Brackets indicate the basal embryo domain where cell division defects were most prevalent. The number of embryos examined and the penetrance of embryos displaying defects in morphology are indicated. Scale bars = 10 μm. Fig. 2 View largeDownload slide NHX5 and NHX6 are involved in auxin-mediated embryo development. Morphology of cleared embryos from globular to heart stage. Arrows indicate the position and region of aberrant cell division and patterning in nhx5 nhx6 embryos compared with the wild type. Brackets indicate the basal embryo domain where cell division defects were most prevalent. The number of embryos examined and the penetrance of embryos displaying defects in morphology are indicated. Scale bars = 10 μm. Auxin gradients and maxima are disrupted in nhx5 nhx6 The phytohormone auxin is critical for meristem growth and cellular patterning in the root tip, and for the initiation, development and emergence of lateral roots (Petrásek and Friml 2009). We questioned whether the disruption to LRP initiation and development in nhx5 nhx6 is a result of auxin-related defects. To investigate this, we assessed auxin response maxima in nhx5 nhx6 mutant plants homozygous for the auxin activity reporter pDR5rev::GFP(Benková et al. 2003). Visualization of DR5::GFP expression in the primary root tip of nhx5 nhx6 seedlings revealed strong expression in the quiescent center and columella root cap cells, similar to the expression in the wild type (Fig. 3A), indicating no obvious defect to auxin maxima at the root tip. The DR5 auxin response marker is also clearly expressed in the tip of LRPs from stage IV/V onwards (Dubrovsky et al. 2008). Interestingly, DR5 expression in late stage LRPs of nhx5 nhx6 mutants was diffuse, with less clearly defined auxin maxima at the primordia tip (Fig. 3B), indicating that auxin response is disrupted in these cells. Fig. 3 View largeDownload slide Auxin gradients are disrupted in nhx5 nhx6. (A) DR5::GFP expression in the primary root tip of 7-day-old seedlings. Counterstain in red is FM5-95. (B) DR5::GFP expression in late stage lateral root primordia. Note the diffuse expression and reduced DR5 auxin maxima in the tips of nhx5 nhx6 compared with the wild type (arrowheads). (C) DR5::GFP expression in triangular and heart stage embryos. Reduced DR5 expression is present in meristem-derived tissue (arrows), and in the apical tips (arrowheads) in nhx5 nhx6 compared with the wild type. The penetrance of auxin signaling defects in heart stage embryos is indicated. Scale bars = 20 μm. Fig. 3 View largeDownload slide Auxin gradients are disrupted in nhx5 nhx6. (A) DR5::GFP expression in the primary root tip of 7-day-old seedlings. Counterstain in red is FM5-95. (B) DR5::GFP expression in late stage lateral root primordia. Note the diffuse expression and reduced DR5 auxin maxima in the tips of nhx5 nhx6 compared with the wild type (arrowheads). (C) DR5::GFP expression in triangular and heart stage embryos. Reduced DR5 expression is present in meristem-derived tissue (arrows), and in the apical tips (arrowheads) in nhx5 nhx6 compared with the wild type. The penetrance of auxin signaling defects in heart stage embryos is indicated. Scale bars = 20 μm. Auxin plays a major role in embryogenesis and leaf development, with auxin gradients controlling the formation of the apical–basal axis and co-ordinating cell division in the embryo and leaf (Scarpella et al. 2006, Möller and Weijers 2009). Given the disruption to cellular patterning in nhx5 nhx6 embryos, we assessed whether defects in auxin response were present similar to those in the LRPs. In wild-type embryos from the triangular stage onwards, DR5 auxin response maxima are present in the basal cells and in the tips of the incipient cotyledons (Fig. 3C) (Benková et al. 2003). In triangular and heart stage nhx5 nhx6 embryos, DR5::GFP auxin reporter maxima were visibly reduced in the basal region, and undetectable in the apical embryo tips, demonstrating a disruption to auxin maxima in these cells. Furthermore, we also occasionally observed defects in leaf patterning in young nhx5 nhx6 seedlings which were accompanied by a reduction in DR5 expression. This finding is consistent with recent evidence reporting reduced DR5::GUS staining in nhx5 nhx6 rosettes (Fan et al. 2018). Taken together, these results show that NHX5 and NHX6 play a role in auxin-mediated plant development in multiple tissues. nhx5 nhx6 mutants exhibit auxin insensitivity and reduced gravitropic response The dwarf phenotype and inhibited root growth of nhx5 nhx6 mutants (Bassil et al. 2011), as well as the auxin-dependent defects reported here, suggest that their perception and/or response to auxin may be disrupted. To investigate this, we tested the response of nhx5 nhx6 seedlings to the synthetic auxin analog 1-naphthaleneacetic acid (1-NAA). Exogenous auxin treatment inhibits root elongation and causes rapid proliferation of LRPs. Root growth assays revealed that nhx5 nhx6 seedlings were less sensitive than the wild type to inhibition of root elongation by 1-NAA at concentrations of ≥250 nM (Fig. 4A), typical of mutants insensitive to auxin (Booker et al. 2003, Ambrose et al. 2013). Next, we assessed whether nhx5 nhx6 mutants were insensitive to the induction of lateral root initiation by 1-NAA treatment. Although LRPs were initiated in nhx5 nhx6 roots in response to 1-NAA, they were underdeveloped with abnormal spacing, often exhibited weak DR5 expression (Fig. 4B) and emerged at lower frequency than in wild-type plants (Fig. 4C). These abnormalities are consistent with the defects in auxin maxima in nhx5 nhx6 LRPs, and suggest that nhx5 nhx6 mutants have reduced auxin transport capability necessary to generate functional auxin gradients in the root. Fig. 4 View largeDownload slide Auxin perception and gravitropism response is altered in nhx5 nhx6. (A) Relative root elongation of seedlings grown on medium supplemented with 1-NAA. Data are means ± SEM of >20 seedlings. Student’s t-test; **P < 0.01. (B) Lateral root initiation in DR5::GFP seedlings after treatment with 10 μM 1-NAA for 72 h. Note the reduction of late stage developed LRPs (arrows) and the presence of underdeveloped LRPs (arrowheads) in nhx5 nhx6. (C) Quantification of LRP development from (B). LRPs were scored as developing (stage I to stage VII) or emerged. At least eight roots were examined for each genotype. (D) Time course of root curvature after gravity stimulus. Data are means ± SEM of >30 seedlings. Student’s t-test; *P < 0.05; **P < 0.01. (E) Auxin distribution in response to gravity stimulus visualized by DR5::GFP 2 h after reorientation. Scale bars = 50 μm (A), 20 μm (E). Fig. 4 View largeDownload slide Auxin perception and gravitropism response is altered in nhx5 nhx6. (A) Relative root elongation of seedlings grown on medium supplemented with 1-NAA. Data are means ± SEM of >20 seedlings. Student’s t-test; **P < 0.01. (B) Lateral root initiation in DR5::GFP seedlings after treatment with 10 μM 1-NAA for 72 h. Note the reduction of late stage developed LRPs (arrows) and the presence of underdeveloped LRPs (arrowheads) in nhx5 nhx6. (C) Quantification of LRP development from (B). LRPs were scored as developing (stage I to stage VII) or emerged. At least eight roots were examined for each genotype. (D) Time course of root curvature after gravity stimulus. Data are means ± SEM of >30 seedlings. Student’s t-test; *P < 0.05; **P < 0.01. (E) Auxin distribution in response to gravity stimulus visualized by DR5::GFP 2 h after reorientation. Scale bars = 50 μm (A), 20 μm (E). Root gravitropism requires the co-ordinated asymmetrical distribution of auxin in the root meristem (Petrásek and Friml 2009). As we identified a possible disruption to auxin transport in nhx5 nhx6 roots, we tested whether the root gravitropic response was affected. In response to gravity stimulus, nhx5 nhx6 seedlings showed slower root re-orientation compared with wild-type seedlings (Fig. 4D). Furthermore, we assessed whether auxin response and redistribution in nhx5 nhx6 may be altered in response to gravity. nhx5 nhx6 seedlings also showed a reduction in the redistribution of auxin to the lower root side after gravistimulation (Fig. 4E). Thus, the reduced gravitropic response in nhx5 nhx6 is likely to be due to a reduced ability to redistribute auxin in the root meristem. The co-ordinated asymmetrical distribution of auxin in root gravitropic response is dependent on PIN2 activity (Abas et al. 2006, Kleine-Vehn et al. 2008), indicating that normal PIN2 function could be disrupted in nhx5 nhx6. Taken together, these data support our findings suggesting that nhx5 nhx6 mutants have a reduced ability to perceive and transport auxin in the root. PIN1 and PIN2 abundance is reduced in nhx5 nhx6 roots To determine if the disruption to auxin gradients and maxima in nhx5 nhx6 tissues is the result of a disturbance in auxin carrier protein abundance or polarity, we first investigated the localization of the AUX/LAX carrier pAUX1:AUX1–YFP in the primary root meristem of nhx5 nhx6. AUX1 localizes to epidermal cells in the root meristem, as well as on the apical plasma membrane in the protophloem (Swarup et al. 2001, Kleine-Vehn et al. 2006). AUX1–YFP polarity and abundance in protophloem and epidermal cells was unaffected in nhx5 nhx6 (Fig. 5A;Supplementary Fig. S3), suggesting that the auxin-related defects may occur independently of AUX1 activity. Fig. 5 View largeDownload slide PIN1 and PIN2 abundance are reduced in nhx5 nhx6. (A–C) Maximum intensity projections of AUX1–YFP (A), PIN1–GFP (B) and PIN2–GFP (C) in the primary root tip of 7-day-old seedlings. (D) Quantification of relative AUX1–YFP, PIN1–GFP and PIN2–GFP fluorescence. Data represent the means ± SEM from n ≥9 seedlings. Student’s t-test; *P < 0.05, **P < 0.01. (E) Localization of PIN1–GFP in developing lateral root primordia. Note the reduction in PIN1–GFP levels in nhx5 nhx6 LRPs during stage V–VII. Scale bars = 20 μm. Fig. 5 View largeDownload slide PIN1 and PIN2 abundance are reduced in nhx5 nhx6. (A–C) Maximum intensity projections of AUX1–YFP (A), PIN1–GFP (B) and PIN2–GFP (C) in the primary root tip of 7-day-old seedlings. (D) Quantification of relative AUX1–YFP, PIN1–GFP and PIN2–GFP fluorescence. Data represent the means ± SEM from n ≥9 seedlings. Student’s t-test; *P < 0.05, **P < 0.01. (E) Localization of PIN1–GFP in developing lateral root primordia. Note the reduction in PIN1–GFP levels in nhx5 nhx6 LRPs during stage V–VII. Scale bars = 20 μm. We next asked whether the reduced gravitropic response and impaired ability to redistribute auxin in nhx5 nhx6 might be associated with changes in PIN distribution or abundance. In the primary root meristem, PIN1 localizes to the basal plasma membrane of stele cells to direct auxin towards the tip, while PIN2 localizes to the apical plasma membrane of epidermal and cortex cells to direct auxin towards the elongation zone (Blilou et al. 2005, Abas et al. 2006, Wisniewska et al. 2006). The expression domain and polarity of pPIN1:PIN1–GFP and pPIN2:PIN2–GFP remained unaffected in nhx5 nhx6 root tips (Fig. 5B, C), indicating that trafficking of PIN1 and PIN2 to the basal or apical plasma membrane, respectively, occurs normally. However, fluorescence intensity levels of PIN1–GFP and PIN2–GFP at the plasma membrane were markedly lower in nhx5 nhx6 compared with the wild type (Fig. 5B–D), demonstrating that the steady-state level of these proteins is disrupted. Furthermore, the signal intensity of PIN1–GFP was also reduced in the provascular cells of stage IV–VII LRPs in nhx5 nhx6 roots (Fig. 5E), consistent with the inhibited DR5 gradients in these cells. These findings suggest that the reduced PIN abundance in root tissue may be related to the disruption of auxin gradients and maxima in nhx5 nhx6. NHX5 and NHX6 are not involved in polar PIN trafficking To test whether NHX5 and NHX6 are directly involved in trafficking of PIN1 or PIN2 to the plasma membrane, we examined the response of PIN1 and PIN2 to BFA inhibition. BFA interferes with GNOM function, and results in the internalization of PIN1 and PIN2 protein into a core BFA compartment composed of aggregated TGN/EE vesicles (Geldner et al. 2001, Geldner et al. 2003, Naramoto et al. 2014). We pre-treated root epidermal cells expressing PIN1–GFP and PIN2–GFP with cycloheximide (CHX) to inhibit protein synthesis, followed by CHX + BFA treatment, in order to examine PIN1 and PIN2 cycling between the TGN/EE and the plasma membrane. Response to BFA treatment and washout of PIN1–GFP and PIN2–GFP in nhx5 nhx6 was similar to that in wild-type seedlings (Fig. 6), suggesting that NHX5 and NHX6 are not directly involved in the polar transport of either PIN1 or PIN2. This finding indicates that while NHX5 and NHX6 are important for the maintenance of PIN1 and PIN2 steady-state levels at the plasma membrane, they are not required for the establishment or maintenance of PIN1 or PIN2 polarity. Fig. 6 View largeDownload slide NHX5 and NHX6 are not directly required for polar PIN1 and PIN2 recycling. Root epidermal cells expressing PIN1–GFP and PIN2–GFP were pre-treated with cycloheximide (CHX) for 1 h to inhibit protein synthesis, then incubated with brefeldin A (BFA) + CHX for 1 h, before being washed in CHX for the indicated times. Note the presence of BFA bodies (arrowheads) after BFA treatment, and the absence of BFA bodies after washout in both wild-type and nhx5 nhx6 cells. Fig. 6 View largeDownload slide NHX5 and NHX6 are not directly required for polar PIN1 and PIN2 recycling. Root epidermal cells expressing PIN1–GFP and PIN2–GFP were pre-treated with cycloheximide (CHX) for 1 h to inhibit protein synthesis, then incubated with brefeldin A (BFA) + CHX for 1 h, before being washed in CHX for the indicated times. Note the presence of BFA bodies (arrowheads) after BFA treatment, and the absence of BFA bodies after washout in both wild-type and nhx5 nhx6 cells. Discussion In this study, we identified a role for NHX5 and NHX6 in auxin-mediated embryonic and post-embryonic growth and development. Detailed characterization of the double knockout nhx5 nhx6 revealed cell patterning and growth defects during embryogenesis and LRP development. Through confocal microscopy imaging, we identified a disruption in DR5 auxin perception, accumulation and redistribution in nhx5 nhx6 seedlings. Furthermore, in nhx5 nhx6 root cells we found that PIN1–GFP and PIN2–GFP abundance was altered, but not their polar trafficking to the plasma membrane. NHX5 and NHX6 are important for root and embryo development In addition to previously reported expression of pNHX6:GUS in the developing embryo (Ashnest et al. 2015), strong promoter activity was observed in the primary root meristem and throughout LRP development. These findings correlate with previous microarray and semi-quantitative RT-PCR experiments demonstrating that NHX5 and NHX6 are expressed in all root tissues (Yokoi et al. 2002, Brady et al. 2007, Bassil et al. 2011). Furthermore, the highly spatial and temporal expression in the early to mid LRPs suggests a specific role for NHX6 in lateral root development. This finding is consistent with the disturbed development and emergence of LRPs, and cellular patterning defects in radicle cells in developing embryos in nhx5 nhx6 double knockouts. NHX5 and NHX6 play roles in auxin-mediated plant development through maintaining PIN homeostasis We found that developmental patterning defects in the embryo and LRPs in nhx5 nhx6 were correlated with a reduction in DR5-GFP expression, indicating a disruption to the establishment of auxin gradients in these tissues. Furthermore, gravitropism and synthetic auxin (1-NAA) experiments revealed that auxin perception and redistribution in the nhx5 nhx6 root are impaired. The generation of functional auxin gradients is required for correct cellular division and patterning during embryogenesis, root meristem growth and LRP formation and development (Benková et al. 2003, Blilou et al. 2005, Möller and Weijers 2009, Robert et al. 2015). Our results demonstrate that the defects to cellular patterning and growth in nhx5 nhx6 are largely auxin dependent. Thus, loss of NHX5 and NHX6 appears to interfere with the generation and establishment of auxin gradients during plant tissue development, with auxin perception and signalling also likely to be affected. Interestingly, it has been reported that free IAA and IAA-conjugate metabolites are altered in nhx5 nhx6 Arabidopsis roots (Fan et al. 2018), suggesting that NHX5 and NHX6 may also affect auxin homeostasis and metabolism. Auxin gradients in plant tissues are established and maintained by the creation of an auxin reflux loop that is largely dependent on PIN-mediated auxin transport (Petrásek and Friml 2009). The reduced PIN1–GFP and PIN2–GFP abundance at the plasma membrane in nhx5 nhx6 root tip cells would limit the effective auxin efflux capacity in the root tip. These data are consistent with the slowed root bending and reduced auxin redistribution during root gravitropism in nhx5 nhx6, processes which are dependent on PIN2 activity (Abas et al. 2006, Kleine-Vehn et al. 2008). Furthermore, in nhx5 nhx6 LRPs, the limited and diffuse localization of PIN1–GFP would be sufficient to inhibit cell division and expansion, consistent with our reported defects in lateral root development and emergence. These data are also supported by evidence from single and multiple pin mutants which display similar defects in primordia patterning and development (Benková et al. 2003). Likewise, the low penetrance and type of cell patterning defects in nhx5 nhx6 embryos are strikingly similar to the phenotype of single pin1 and pin4 mutant embryos which have reduced auxin transport (Friml et al. 2002, Robert et al. 2015). Thus, our data suggest that NHX5 and NHX6 affect auxin gradient establishment through maintaining PIN homeostasis. NHX5 and NHX6 affect PIN homeostasis independently of polar PIN transport Published microarray analysis of gene expression in nhx5 nhx6 whole seedlings revealed no large changes to expression levels in any PIN genes (Bassil et al. 2011), suggesting a post-translational mechanism behind the reduction to steady-state PIN levels at the plasma membrane. Instead, the subcellular trafficking or recycling of PINs to or from the plasma membrane may be altered in nhx5 nhx6 cells. PINs are constitutively endocytosed to the TGN/EE, and are subsequently recycled back to the plasma membrane through polar transport pathways (Geldner et al. 2001, Dhonukshe et al. 2007, Friml 2010). BFA washout experiments indicate that the polar delivery and recycling of PIN1 and PIN2 to the plasma membrane are unaffected in nhx5 nhx6, also consistent with the lack of defects to PIN1 and PIN2 polarity at the plasma membrane. Furthermore, the non-polar delivery of AUX1 to the plasma membrane occurred normally, suggesting that the general recycling of auxin carriers to the plasma membrane is functional in nhx5 nhx6. Thus, NHX5 and NHX6 appear to impact PIN homeostasis through a pathway independent of polar PIN recycling. NHX5 and NHX6 may assist in retromer-mediated PIN retrieval While NHX5 and NHX6 are reported to function in vacuolar trafficking of soluble cargo proteins (Bassil et al. 2011, Ashnest et al. 2015, Reguera et al. 2015), their role in the vacuolar trafficking of membrane-bound receptors has not been investigated. PIN proteins targeted for degradation are transported to the vacuole via late endocytic pathways, and can be retrieved before degradation from this pathway through retromer complex-mediated targeting (Kleine-Vehn et al. 2008, Nodzyński et al. 2013). Thus, retromer activity maintains PIN levels at the TGN/EE for subsequent recycling back to the plasma membrane, assisting in the fine-tuning of PIN abundance at the plasma membrane. We hypothesized that the reduced PIN abundance in nhx5 nhx6 may be due to a disruption to PIN vacuolar trafficking pathways. Interestingly, NHX5 and NHX6 mutants share striking phenotypic similarities with retromer mutants. We previously reported that a key component of the retromer complex, SNX1, interacts with the cytosolic tail of NHX6, and co-localizes with NHX5 in endosomal compartments (Ashnest et al. 2015). Furthermore, retromer, as well as other mutants associated with defects in protein trafficking to the lytic and protein storage vacuole exhibit sucrose-conditional developmental arrest (Shimada et al. 2006, Kleine-Vehn et al. 2008, Silady et al. 2008, Feraru et al. 2010, Zwiewka et al. 2011), a phenotype attributed to defects in the late steps of the endocytic pathway. Moreover, retromer mutants snx1 and vps29 display defects in lateral root growth, reduced gravitropic response, and have reduced PIN1/PIN2 abundance but normal polar PIN recycling (Jaillais et al. 2006, Jaillais et al. 2007, Kleine-Vehn et al. 2008, Ambrose et al. 2013). The SNX1–NHX6 interaction, together with the reduced PIN abundance but functional polar PIN recycling in nhx5 nhx6 shown, here suggests that NHX5 and NHX6 may assist in the retromer-mediated retrieval of PINs. While we propose that NHX5 and NHX6 facilitate PIN homeostasis through the retromer complex, the mechanism of action is unclear. We hypothesize that NHX5 and NHX6 may be involved in PIN trafficking through the maintenance of pH homeostasis in endomembrane compartments. Recent evidence has shown that NHX5 and NHX6 antiporter activity regulates pH homeostasis of the Golgi, TGN/EE and PVC compartments (Reguera et al. 2015). Furthermore, the disruption to endosomal pH in nhx5 nhx6 leads to reduced association of a vacuolar sorting receptor with its cargo (Reguera et al. 2015). In analogy to this, endosomal pH defects in nhx5 nhx6 could potentially compromise retromer–PIN association, and hence limit PIN retrieval from vacuolar targeting for degradation. Furthermore, we speculate that the NHX6–SNX1 interaction could enable local pH adjustment at the site of retromer binding, which may facilitate PIN binding and retrieval. Future investigation of any pH sensitivity of retromer-mediated PIN retrieval, including structural analysis of this association, may address this hypothesis. In conclusion, our data provide new insights into the cellular mechanism of how endosomal NHX antiporters regulate plant growth. We show that NHX5 and NHX6 function during embryo and root development and are required for functional LRP development and emergence. Moreover, NHX5 and NHX6 mediate auxin gradients and maxima through the maintenance of steady-state PIN levels at the plasma membrane. We hypothesize that NHX5 and NHX6 influence PIN protein abundance through late endocytic trafficking pathways, probably involving retromer-mediated PIN retrieval. Materials and Methods Plant material and growth conditions Arabidopsis thaliana lines were all in the Columbia-0 (Col-0) accession background. Plant lines used have been previously described: nhx5-2 nhx6-3 (Ashnest et al. 2015); pDR5rev::GFP and pPIN1::PIN1-GFP (Benková et al. 2003); pAUX1::AUX1-YFP (Swarup et al. 2004); pPIN2::PIN2-GFP in eir1-4 (Xu and Scheres 2005); and pNHX6::GUS (Ashnest et al. 2015). Seeds were surface sterilized with 70% ethanol for 5 min and 10% bleach for 5 min, washed three times in ddH2O and grown on half-strength Murashige and Skoog (1/2 MS) medium containing 1.0% (w/v) agar, pH 5.8, without sucrose unless indicated. Seedlings were stratified for 48 h at 4°C in the dark, and grown in a 16 h light/8 h dark photoperiod at 22°C. Wild-type (Col-0) lines containing the pDR5rev::GFP, pAUX1-AUX1-YFP, pPIN1-PIN1-GFP and pPIN2-PIN2-GFP transgenes were crossed with NHX5/nhx5-2 nhx6-3/nhx6-3 lines. F1 plants from these initial crosses were selfed, and homozygous nhx5-2 nhx6-3 lines were identified in the segregating F2 population by PCR genotyping as previously described (Ashnest et al. 2015). Only lines homozygous for the appropriate reporter constructs were used for analysis. pNHX6:GUS reporter assays Whole roots from 10-day-old Arabidopsis seedlings carrying a 3 kb promoter fragment immediately upstream of the start codon of NHX6 (Ashnest et al. 2015) were fixed in cold 90% acetone for 30 min, then washed twice in 100 mM sodium phosphate buffer (pH 7.2) before staining in GUS solution [100 mM sodium phosphate (pH 7.2), 1 mM EDTA, 5 mM potassium ferricyanide, 5 mM potassium ferrocyanide, 1% Triton X-100 and 1 mg ml–1 X-Gluc] for 24 h. Stained tissue was cleared by mounting in Hoyer’s solution (70% chloral hydrate, 4% glycerol, 5% gum arabic) and incubating overnight before imaging using a Zeiss Axio Observer inverted microscope. Microscopy and image quantification Confocal microscopy was performed using a Zeiss LSM 510 or LSM 780 confocal laser scanning microscope (Carl Zeiss) with a C-Apochromat ×40/1.2 W or C-Apochromat ×40/1.3 W objective. Excitation and emission detection settings were as follows: GFP/YFP, 488 nm/490–560 nm, FM5-95, 561 nm/565–650 nm. For all quantification experiments, identical acquisition settings were used to acquire each image. Chemical stock solutions were made in dimethylsulfoxide (DMSO) at the following concentrations: BFA 50 mM, CHX 50 mM, FM5-95 (FM4-64 analog) 4 mM, 1-NAA 10 mM. For BFA treatments, 7-day-old seedlings were incubated in 6-well plates containing 1/2 MS with 50 μM CHX for 60 min, 50 μM BFA + 50 μM CHX for 60 min, and washed out in 50 μM CHX for the times indicated. For FM5-95 counterstaining, roots were incubated for 5 min at room temperature in 1/2 MS medium containing 2 μM FM5-95 dye, and washed twice in 1/2 MS before imaging. For the 1-NAA root elongation experiment, seedlings were grown on 1/2 MS vertical plates containing 1-NAA at the indicated concentration for 8 d before imaging and analysis. The root length of untreated plants was set at 100%. For the 1-NAA LRP induction experiment, seedlings were first grown on solid 1/2 MS plates for 4 d and then transferred to solid 1/2 MS containing 10 μM 1-NAA for 3 d before being imaged by confocal microscopy. Lateral root phenotyping experiments were performed using seedlings expressing the AUX1:YFP marker grown on 1/2 MS vertical plates for 15 d. For the nhx5 nhx6 mutants, only seedlings which did not arrest growth were used for analysis. For each seedling, the medial slice of each observable LRP was imaged using a Zeiss Imager M2 fluorescence microscope with a 525/50 nm GFP filter. Primordia were staged similarly to as previously described (Malamy and Benfey 1997, Lucas et al. 2013). For the root gravitropism experiments, 7-day-old seedlings were grown on 1/2 MS vertical plates before being rotated 90° for the indicated time. The angle of root curvature in at least 30 seedlings per genotype was measured in ImageJ. The pDR5:GFP images were captured by acquiring z-stacks of 6–8 slices at 2.5 μm per slice, from at least 12 seedlings for each genotype. Quantification of PIN1–GFP and PIN2–GFP fluorescence was obtained from 15–20 z-stack slices taken at 1 μm intervals, from which a maximum intensity projection was generated. Mean gray values were obtained from the PIN1–GFP or PIN2–GFP localization region, and subtracted from the mean gray value obtained from the background. Relative fluorescence was calculated with the mean wild-type gray value set as 100%. Post-processing of images was performed with Zeiss ZEN Black (v8.0) and ImageJ v1.48g. Statistical analysis was performed using Microsoft Excel. Inducible promoter The NHX6 β-estradiol-inducible rescue construct was generated by ligating the NHX6 open reading frame (ORF) into the pMDC7 β-estradiol-inducible construct (Curtis and Grossniklaus 2003). Primer sequences are listed in Supplementary Table S1. To induce NHX6 expression, transgenic nhx5 nhx6 seed harboring the inducible NHX6 rescue construct were germinated directly on 1/2 MS medium supplemented with 8 µM β-estradiol (Sigma-Aldrich, E8875). Root lengths and emerged lateral roots were quantified after 14 d. Control medium was supplemented with equivalent concentrations of DMSO. Embryo dissection and analysis Embryos were dissected from siliques of different developmental stages using hypodermic needles. Embryos were cleared on microscope slides in clearing solution (chloral hydrate:water:glycerol (7:3:1, by vol.) and incubated at 4°C for 1 h. Images were obtained with a Zeiss Axio Observer microscope using ×20 air and ×63 oil objectives. Embryos were scored by stage from late globular to mid heart according to ten Hove et al. (2015) in three biological repeats. Embryos were scored as defective if they displayed clear cellular patterning defects in the incipient cotyledon or root cells. For DR5:GFP localization, embryos were dissected from siliques onto microscope slides containing 1/2 MS liquid and imaged immediately on an LSM 780 confocal laser scanning microscope with a C-Apochromat ×40/1.3 W objective. Only clearly intact embryos were used for analysis. Supplementary Data Supplementary data are available at PCP online. Funding This work was supported by the Australian Federal Government [an Australian Postgraduate Award to J.D.]; the Grains Research and Development Corporation [Graduate Research Scholarships to J.A. (GRS 179) and B.F. (GRS 161)]; the Government of India, Ministry of Social Justice and Empowerment [National Overseas Scholarship to P.D.]; La Trobe University [Post-Graduate Research Scholarship to P.D.]; the Australian Research Council [Linkage Infrastructure, Equipment and Facilities grant (LE0989920).] Acknowledgments The authors would like to thank sincerely the ABRC, GABI-Kat, D. Smyth, C. Luschnig and T Gaude for supplying seeds, M. 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Plant and Cell Physiology – Oxford University Press
Published: May 16, 2018
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