SCFTIR1/AFB‐auxin signalling regulates PIN vacuolar trafficking and auxin fluxes during root gravitropism
The EMBO Journal
(2012)
32: 260 - 274
The distribution of the phytohormone auxin regulates many aspects of plant development including growth response to gravity. Gravitropic root curvature involves coordinated and asymmetric cell elongation between the lower and upper side of the root, mediated by differential cellular auxin levels. The asymmetry in the auxin distribution is established and maintained by a spatio‐temporal regulation of the PIN‐FORMED (PIN) auxin transporter activity. We provide novel insights into the complex regulation of PIN abundance and activity during root gravitropism. We show that PIN2 turnover is differentially regulated on the upper and lower side of gravistimulated roots by distinct but partially overlapping auxin feedback mechanisms. In addition to regulating transcription and clathrin‐mediated internalization, auxin also controls PIN abundance at the plasma membrane by promoting their vacuolar targeting and degradation. This effect of elevated auxin levels requires the activity of SKP‐Cullin‐F‐boxTIR1/AFB (SCFTIR1/AFB)‐dependent pathway. Importantly, also suboptimal auxin levels mediate PIN degradation utilizing the same signalling pathway. These feedback mechanisms are functionally important during gravitropic response and ensure fine‐tuning of auxin fluxes for maintaining as well as terminating asymmetric growth.
Introduction
The phytohormone auxin is an important regulator of cell morphogenesis shaping and directing growth of organs within different developmental contexts and in response to environmental signals (Vanneste and Friml, 2009). To ensure optimal growth and development, plants have acquired elaborate mechanisms to control the local auxin homeostasis, including control of auxin metabolism (Cheng et al, 2006, 2007; Stepanova et al, 2008; Tao et al, 2008), subcellular compartmentalization (Mravec et al, 2009; Barbez et al, 2012; Ding et al, 2012) and directional auxin transport mediated by plasma membrane‐resident transporters, such as ABCB, PIN‐FORMED (PIN) and AUXIN‐RESISTANT 1 (AUX1) (Bennett et al, 1996; Geisler et al, 2005; Petrášek et al, 2006; Cho et al, 2007; Swarup et al, 2008; Jones et al, 2009). One of the prominent growth responses mediated by auxin transport is root gravitropism. Changes of the orientation relative to the gravity vector are perceived in the root tip, by the sedimentation of statoliths, defined as gravity‐sensing organelles (Harrison and Masson, 2008; Leitz et al, 2009; Morita, 2010). This process appears to induce the relocation of the auxin efflux carriers (Petrášek et al, 2006) PIN3 and PIN7 to the lower side of the gravity‐sensing cells, which presumably aligns auxin flux with gravity vector towards the lower side of the root tip (Friml et al, 2002; Harrison and Masson, 2008; Kleine‐Vehn et al, 2010). From there, another auxin efflux carrier, PIN2, which is apically (shootward, upper cell side) localized in the lateral root cap and epidermal cells, mediates the directional auxin flow from the root tip to the elongation zone where control of elongation occurs (Luschnig et al, 1998; Müller et al, 1998; Abas et al, 2006; Wiśniewska et al, 2006). Hence, the PIN‐mediated establishment of the asymmetric auxin distribution leads to a differential growth between the lower and the upper side of the root. As a consequence, root bends and re‐orients in respect to the gravity vector, allowing the efficient exploration of the soil (Firn et al, 2000; Swarup et al, 2005).
The mechanisms underlying the PIN3 and PIN7 polarization in gravity‐sensing columella cells and control of the PIN2 abundance at the plasma membrane for defined gravitropic response remain largely elusive. Nevertheless, some of the molecular processes controlling the subcellular localization of PIN proteins have been characterized (Grunewald and Friml, 2010). PIN proteins internalize continuously via a clathrin‐mediated endocytotic pathway (Dhonukshe et al, 2007; Kitakura et al, 2011) and cycle back to the plasma membrane as shown by pharmacological approaches with a vesicle‐budding inhibitor, Brefeldin A (BFA) (Geldner et al, 2001). This permanent cycling leads to a dynamic control of their polar localization and abundance at the plasma membrane (Kleine‐Vehn et al, 2008a), which in turn, determines the rate and direction of the auxin flow (Paciorek et al, 2005; Wiśniewska et al, 2006). The constitutive endocytic recycling enables also rapid switches in PIN polarity and, consequently, directionality of auxin fluxes in response to environmental signals, including light and gravity (Friml et al, 2002; Kleine‐Vehn et al, 2010; Ding et al, 2011; Rakusová et al, 2011).
Besides the control of the polar localization, PIN protein activity can be also regulated by degradation. Numerous studies reported the occurrence of PIN degradation in the vacuoles (Abas et al, 2006; Laxmi et al, 2008; Kleine‐Vehn et al, 2008b; Shirakawa et al, 2009; Leitner et al, 2012; Marhavý et al, 2011), to which they are targeted via a BFA‐sensitive canonical retrograde trafficking pathway, involving the retromer complex (Kleine‐Vehn et al, 2008b). Moreover, PIN2 turnover depends on the proteasomal activity (Sieberer et al, 2000; Abas et al, 2006) and sorting for vacuolar delivery was recently associated with the formation of the polyubiquitin chains linked to the specific lysine residues at the PIN2 hydrophilic loop (Leitner et al, 2012). Together, this data highlights the importance of post‐transcriptional regulations in auxin flux determination.
Notably, auxin itself modulates its own distribution by providing feedback on PIN biosynthesis and trafficking (Benjamins and Scheres, 2008). Short auxin treatments (⩽2 h) activate the transcription of different PIN genes (Peer et al, 2004; Heisler et al, 2005; Vieten et al, 2005; Scarpella et al, 2006) and can stabilize PIN at the plasma membrane by inhibiting clathrin‐mediated internalization (Paciorek et al, 2005; Robert et al, 2010). Recently, it was found that AUXIN‐BINDING PROTEIN 1 (ABP1) is a positive regulator of clathrin‐mediated endocytosis, which is inhibited upon auxin binding (Robert et al, 2010; Chen et al, 2012). In contrast, prolonged application of auxin also promotes the turnover of PIN proteins via an unknown mechanism (Sieberer et al, 2000; Vieten et al, 2005; Abas et al, 2006). How this duality of auxin action on endocytosis versus degradation is regulated is unknown.
The BFA fungal toxin is known to inhibit the activity of specific ADP‐ribosylation factor GTP‐exchange factors (ARF‐GEFs) (Peyroche et al, 1999; Sata et al, 1999; Geldner et al, 2003). In plants, the secretory pathway is readily inhibited by BFA, resulting in the intracellular accumulation of endocytosed plasma membrane proteins such as PIN proteins (Geldner et al, 2001). Upon inhibition of endocytosis (at ∼25 μM of BFA), PIN proteins no longer end up in such a BFA compartments (Paciorek et al, 2005; Men et al, 2008; Kitakura et al, 2011). Interestingly, it has recently been discovered that at higher concentrations (∼50 μM), BFA also inhibits vacuolar targeting and degradation of PIN proteins (Kleine‐Vehn et al, 2008b; Kleine‐Vehn and Friml, 2008; Robert et al, 2010). Thus, different concentrations of BFA allow discriminating between effects on endocytosis for recycling and targeting for degradation (Robert et al, 2010). Notably, the aforementioned BFA concentration cutoff should not be taken precisely as most likely specificity of the BFA towards specific ARF‐GEF's changes gradually.
Here, we show that PIN2 protein abundance is dynamically and differentially controlled at the upper and lower sides of a gravistimulated root. Both increased and decreased auxin levels change PIN2 stability by a post‐transcriptional regulation of its vacuolar targeting. Moreover, we provide additional data to clarify the involvement of SCFTIR1/AFB‐based signalling in auxin‐mediated PIN turnover. These findings link auxin‐mediated regulation of vesicle transport and asymmetric growth control during gravitropic response.
Results
Dynamic changes of auxin response and PIN2 abundance in gravistimulated roots
To better understand the regulation of auxin transport activity in response to gravity, we have investigated the dynamics of root bending, auxin redistribution and abundance of PIN2 in gravistimulated roots of Arabidopsis thaliana. We have indirectly visualized the auxin redistribution by monitoring the activity of the synthetic auxin‐responsive promoter DR5rev (Ulmasov et al, 1997) driving expression of a nuclearly localized VENUS protein (DR5rev::3xVENUS‐N7; Heisler et al, 2005). Consistent with previous observations, after 2 h of gravistimulation, Arabidopsis root bent visibly (Figure 1A–F) and an asymmetric increase in DR5, expression was observed at the less elongated (lower) root side (Ottenschläger et al, 2003; Paciorek et al, 2005), whereas at the upper side of the bending root, the DR5 response was reduced (Figure 1G–M). This asymmetry in auxin response was maintained throughout gravity‐induced root bending (Figure 1A–M). In time, the growth angle of the root became progressively parallel to the gravity vector (Figure 1F) and with a delay, a balanced DR5 expression was re‐establishing (Figure 1L and M). We have confirmed the formation of auxin lateral gradient in roots responding to the gravity with use of highly dynamic DII‐VENUS reporter system (see Supplementary Figure 1; Brunoud et al, 2012). This system was previously used to precisely place the timing of auxin accumulation during root gravitropic response (Band et al, 2012). It is important to note that the timing of onset and disappearance of the DR5rev::3XVENUS‐N7 signal lags behind the real kinetics of the auxin distribution due to the time needed for VENUS maturation and turnover. Nonetheless, in spite of the inherent shortcomings of this reporter, we were able to demonstrate a clear spatio‐temporal regulation of the auxin distribution during gravitropic bending.

To further characterize the regulation of the efflux carrier activity in response to gravity, we have investigated PIN2 abundance at the plasma membrane of gravistimulated roots. As previously suggested, following gravistimulation, PIN2 distribution became asymmetric between the upper and lower sides of the root, in concordance with an asymmetrical auxin distribution (Paciorek et al, 2005; Abas et al, 2006; Kleine‐Vehn et al, 2008b) (Figure 1N–T). We have quantified the plasma membrane‐localized PIN2 abundance at the lower and upper sides of horizontally placed roots at different time points after gravistimulation. Within 2 h of gravistimulation, an increase in PIN2 at the plasma membrane of cells on the lower root side was detected, which spatially correlated with an increase in auxin response (Figure 1H, M). Following this temporal stabilization, the PIN2 level at the lower side of the root started to decrease to supposedly re‐establish the pre‐stimulation levels after 12 h of gravistimulation (Figure 1O–S). Thus, at the lower root side, the PIN2 levels transiently increased before gradually decreasing to the pre‐stimulation values.
In parallel, at the upper side of the bending root, where auxin response initially decreases (Figure 1G–M), PIN2 protein levels at the plasma membrane steadily decreased in time (Figure 1N–P and S), probably because of higher rates of protein degradation due to an increased targeting to the vacuole (Kleine‐Vehn et al, 2008b; Figure 4A and B). Notably, after 4 h of gravistimulation, PIN2 started to accumulate again at the plasma membrane, reaching levels close to the initial pre‐stimulation levels at ∼12 h after gravistimulation (Figure 1P–S). Thus, at the upper root side, the PIN2 levels initially decrease, which is followed by an increase leading to re‐establishment of the pre‐stimulation values. The observed changes in signal intensity infer ∼12 and 14% change in PIN2 abundance on the lower and upper sides of gravistimulated root, respectively. Notably, the recovery of symmetry in PIN2 protein levels at the plasma membrane after 12 h of gravistimulation at both lower and upper sides of the root presumably reflects a re‐established symmetric auxin flow, resulting in vertical root growth (Figure 1).
Overall, our data shows that a spatio‐temporal regulation of the auxin distribution after gravistimulation correlates with complex and differential regulation of the PIN2 abundance at the lower and upper side of gravistimulated roots. Specifically, the increase in auxin response at the lower side of the root is accompanied with the initial increase in PIN2 abundance followed by its gradual decrease. On the other hand, at the upper side of the root, we have detected a decrease in auxin response that is accompanied with initial decrease in PIN2 abundance followed by its gradual increase. Importantly, the differential auxin accumulation in all observed cases preceeded changes in PIN2 abundance at the plasma membrane. The above findings also complement the observation of Luschnig et al (1998) that a missense pin2 allele fails to establish gravity‐induced lateral auxin gradient in the root.
Auxin promotes PIN2 degradation in the vacuoles at the lower side of the root
First, we have addressed the mechanisms underlying the regulation of PIN2 abundance at the lower side of the gravistimulated root. The initial, transient stabilization of PIN2 at the plasma membrane is presumably a result of a documented transient (⩽2 h) inhibitory effect of higher auxin levels on PIN internalization (Paciorek et al, 2005; Robert et al, 2010; Chen et al, 2012; Lin et al, 2012). On the other hand, the following decrease in PIN2 levels that still coincides with a DR5‐visualized local increase in auxin response (Figure 1) might be result of the long‐term effect of auxin on PIN stability (Sieberer et al, 2000; Vieten et al, 2005). Therefore, we have tested the effect of prolonged (⩾3 h) exogenous auxin application on PIN2 abundance at the plasma membrane. Following NAA treatment, we have observed a reduction of PIN2‐GFP levels (in PIN2::PIN2‐GFP (eir1‐1) transgenic seedlings) at the plasma membrane concomitantly with an increase in a diffused vacuolar GFP signal (Figure 2A–C; see Supplementary Figure 2). This observation was confirmed by a significant reduction of PIN2 abundance in membrane protein extracts from NAA‐treated seedlings as detected by western blots (Figure 2D).

We have then addressed the cellular mechanism of the auxin effect on PIN2 abundance. In general, protein abundance at the plasma membrane is expected to reflect a sum of transcription, translation, targeting and proteolysis. It has been shown previously that PIN2 transcription does not change dramatically in response to auxin (Sieberer et al, 2000; Shin et al, 2005). Consistently, PIN2 mRNA levels were shown to be induced by auxin with low amplitude and much slower kinetics than other PIN genes or other auxin inducible genes (Vieten et al, 2005; Lee et al, 2009). In agreement with those findings, in our experimental conditions, auxin treatment only mildly affected PIN2 transcription (see Supplementary Figure 3), suggesting that auxin regulates PIN2 levels via a post‐transcriptional mechanism. Moreover, auxin‐mediated decrease in PIN2 from the plasma membrane occurred regardless of whether PIN2 was expressed under its endogenous (Figure 2A–C) or constitutive, heterologous 35S promoter (Figure 2E–G), suggesting that the increased downregulation of PIN2 is not an indirect effect of an excess of PIN2 protein in the cell's endomembrane system.
It is proposed that PIN proteins are degraded in the vacuoles (Laxmi et al, 2008; Kleine‐Vehn et al, 2008b; Shirakawa et al, 2009; Marhavý et al, 2011), where GFP‐tagged proteins can be visualized after an incubation in the dark (Tamura et al, 2003). In these conditions, we have found a decrease in PIN2‐GFP at the plasma membrane and concomitant increase in fluorescence signal in the vacuoles in response to auxin treatment (Figure 2A–C; see Supplementary Figure 2K–M). This strongly suggests that auxin downregulates PIN2 abundance at the plasma membrane by enhancing PIN trafficking to the vacuole. Moreover, the auxin application destabilized both apical and basal PIN1 and PIN2 cargos from the plasma membrane (Figure 2H–J; see Supplementary Figures 2A–C and 4A–C) and to lesser extent also non‐polar integral plasma membrane proteins such as BRASSINOSTEROID INSENSITIVE1 (BRI1)‐GFP and PLASMA MEMBRANE INTRINSIC PROTEIN2 (PIP2)‐GFP (see Supplementary Figure 4D–I). In addition, we could demonstrate that auxin reduces PIN2 protein levels (Figure 2D), thereby strongly suggesting that the observed vacuolar targeting of PINs is associated with protein degradation.
To further confirm the auxin effect on the degradation of plasma membrane proteins, we have genetically manipulated the endogenous auxin concentrations in Arabidopsis seedlings. We have constitutively overexpressed the Agrobacterium tumefaciens indoleacetic acid‐tryptophan monooxygenase (iaaM) under the strong ribosomal promoter RPS5. The iaaM enzyme converts tryptophan into indole‐3‐acetamide, which is then hydrolysed to indole‐3‐acetic acid (IAA) in plant cells (Klee et al, 1987; Romano et al, 1995; Weijers et al, 2001, 2005). The transcription of PIN2 was not altered in the RPS5»iaaM transactivated line (see Supplementary Figure 5). We have then analysed the abundance and intracellular distribution of PIN2‐GFP marker crossed into the iaaM background. Similarly to exogenously applied, endogenously produced auxin promoted an increased PIN2 degradation as manifested by higher vacuolar GFP signal (Figure 2K–M). The iaaM expression was shown to elevate cellular auxin concentration 2‐ to 10‐fold (Klee et al, 1987; Romano et al, 1995), Therefore, considering that we have not used additional media supplementation, neither with tryptophan nor with auxin, it can be expected that the physiological threshold of auxin effect on increased PIN degradation is placed in the aforementioned range of auxin concentration change above normal/physiological level.
Taken together, this data shows that exogenously applied or endogenously produced auxin mediates the PIN targeting to the vacuole and promotes PIN2 degradation. This auxin effect presumably accounts for the decrease in PIN2 level at the lower side of the gravistimulated root after 4 h.
Auxin promotes PIN2 degradation by SCFTIR1/AFB‐mediated signalling
Next, we have assessed by which signalling pathway auxin promotes PIN2 degradation. We have previously shown that the inhibitory effect of auxin on PIN endocytosis is mediated by an ABP1‐dependent signalling. Whereas auxin inhibits endocytosis instantaneously without de novo protein biosynthesis and nuclear auxin signalling (Paciorek et al, 2005; Robert et al, 2010; Chen et al, 2012; Lin et al, 2012), the auxin‐induced PIN2 translocation to the vacuole for degradation required prolonged (⩾3 h) auxin treatments (Figure 2A–C, see Supplementary Figure 4A–C). Given the fact that the earliest auxin‐induced response proteins are detectable after ∼10–15 min of auxin application (Badescu and Napier, 2006), the auxin effect on the vacuolar targeting might require transcriptional regulation and de novo protein synthesis mediated by the SCFTIR1/AFB pathway (Kepinski and Leyser, 2005; Dharmasiri et al, 2005a, 2005b; Badescu and Napier, 2006; Tan et al, 2007). To test this hypothesis, we have used structural auxin analogues that can discriminate between ABP1‐ and SCFTIR1/AFB‐mediated signalling (Robert et al, 2010). We have observed that treatment with IAA and all the synthetic auxin analogues, which induced transcriptional auxin response (as monitored by DR5rev::GFP), also promoted the degradation of PIN proteins. Moreover, the compounds, which did not induce DR5rev::GFP expression, did not cause a decrease in PIN abundance at the plasma membrane (Figure 3A–J; see Supplementary Figures 6 and 7). This data suggests that the same auxin perception mechanism and downstream effectors mediates regulation of gene transcription and control the PIN stability at the plasma membrane.

Indeed, in the quadruple tir1afb1,2,3 mutant, auxin did not downregulate the PIN protein levels, showing a resistance to the auxin effect on PIN degradation (Figure 4A, B). Importantly, resistance could also be observed in double tir1afb1, tir1afb2, tir1afb3 and partially in the single tir1‐1 mutant background, which all show comparable PIN protein levels to the wild type in control (untreated) conditions (see Supplementary Figure 8). We have also tested the abp1‐5 allele that contained a point mutation in the auxin‐binding domain of ABP1 (Napier et al, 2002), and thus exhibited reduced auxin sensitivity (Robert et al, 2010; Xu et al, 2010). The auxin effect on PIN degradation in the abp1‐5 mutant was comparable to the one observed in the wild type (Figure 4C, F and G).

Next, we have attempted to identify downstream molecular components of the SCFTIR1/AFB pathway that are involved in the control of PIN2 degradation process. We have analysed the promoter expression of 23 ARF genes in the root meristem using transcriptional nuclear GFP fusions (Rademacher et al, 2011). We have identified ARF's 1, 2, 6, 9, 10, 16 and 19 as prominently expressed in epidermis of the root meristematic region where PIN2 is also specifically expressed (see Supplementary Figure 9). Subsequently, we have employed the 50 μM BFA and 20 μM NAA co‐treatment on the arf2, arf6, arf10arf16, arf19 and arf7arf19 mutant lines. We were able to observe an increased PIN2 accumulation in BFA induced agglomeration in the arf2 mutant when compared to the wild‐type control. This effect was not observed after treatment with lower concentration of BFA (see Supplementary Figure 10). This suggests that the mutation in the ARF2 gene disturbs vacuolar trafficking of PIN2 protein. We therefore propose that this transcription factor could be more specifically involved in the control of PIN2 vacuolar targeting.
Overall, these data suggest that SCFTIR1/AFB‐dependent signalling is required for auxin‐induced PIN2 degradation. Thus, at the lower side of the gravistimulated root, overlapping auxin effects on PIN2 endocytosis (ABP1‐mediated) and PIN2 vacuolar targeting (SCFTIR1/AFB‐mediated) presumably account for a transient increase in a PIN2‐mediated auxin flow as well as for its subsequent decrease to the pre‐stimulation levels.
Auxin depletion promotes PIN2 degradation at the upper side of the root
Next, we have examined the mechanisms underlying the regulation of PIN2 abundance at the upper side of the gravistimulated root. Here, PIN2 levels steadily decreased coinciding with reduced DR5‐visualized auxin response (Figure 1). As seen for endogenous PIN2 (Figure 1), a similar asymmetric distribution with decreased levels at the upper epidermal cell file was observed also for PIN2‐EosFP expressed under control of constitutive 35S promoter (see Supplementary Figure 11), suggesting that this decrease occurs independently of PIN transcriptional regulation. Furthermore, this decrease correlated with the increased PIN2 vacuolar targeting (Kleine‐Vehn et al, 2008b; Figure 4A and B) also consistent with post‐transcriptional regulation.
We have tested whether decrease in PIN2 abundance at the plasma membrane and increased vacuolar targeting might be possibly a consequence of prolonged reduction in auxin levels. In Arabidopsis seedlings, not only the young leaves but also the cotyledons have a high capacity for auxin biosynthesis (Ljung et al, 2001). We have therefore reduced auxin biosynthetic capacity of the seedlings by removal of the cotyledons and shoot apical meristem (decapitation). We have observed that 14 h after such a decapitation, the growth rate of the roots was decreased but roots were still graviresponsive (see Supplementary Figure 12; Rashotte et al, 2000). By using DR5rev::3XVENUS‐N7, we have detected a significant decrease in DR5‐monitored auxin response in the PIN2 expression domain 14 h after decapitation (Figure 5A–C). These results are in line with previously reported findings showing that the auxin maximum in the root tip is highly stable and a decrease in auxin levels in the elongation zone can be detected only when auxin depletion by decapitation is prolonged (Grieneisen et al, 2007). Importantly, as a consequence of decapitation, we have observed a decreased PIN2 abundance at the plasma membrane and enhanced targeting to the vacuole (Figure 5D–F). This effect was independent of transcriptional control (see Supplementary Figure 13) and could be reversed by exogenous auxin application (see Supplementary Figure 14). What is more, we have confirmed the reduction of PIN2 level by western blot analysis of membrane fractions isolated 14 h after decapitation (Figure 5G).

To further simulate auxin depletion, we have used two independent chemical biology‐based approaches. First, we have used the auxin‐antagonist α‐(phenyl ethyl‐2‐one)‐indole‐3‐acetic acid (PEO‐IAA) (Hayashi et al, 2008) that counteracts the auxin effect on transcription presumably by binding to the SCFTIR1 receptor (Nishimura et al, 2009). After 3 h of treatment, PEO‐IAA caused a drop in auxin signalling as reflected by reduced expression of DR5rev::3XVENUS‐N7 reporter in epidermal and lateral root cap cells of the root apical meristem (Figure 6A–C). Similar treatment increased vacuolar targeting of PIN2 protein (Figure 6D–F) and, to lesser extent also non‐polar integral plasma membrane proteins BRI1‐GFP and PIP2‐GFP (see Supplementary Figure 15). We have additionally observed that PEO‐IAA disturbed the formation of the lateral gradient of PIN2 and consequently gravitropic response of the roots (see Supplementary Figure 16). Importantly, the PEO‐IAA‐induced destablization of PIN2 from the plasma membrane could be counteracted by exogenous auxin application (see Supplementary Figure 17). Western blot analysis of membrane protein fractions confirmed reduced PIN2 levels after PEO treatment (Figure 6G). This suggests that a lower throughput of SCFTIR1/AFB‐mediated transcriptional auxin pathway decreases PIN stability at the plasma membrane. Second, we have interfered with Trp‐dependent auxin biosynthesis by compromising the activity of key enzymatic components of this pathway. We have used l‐Kynurenine, a competitive and specific inhibitor of TAA1/TAR enzymatic activity, which was shown to reduce DR5‐GUS expression in the Arabidopsis roots (He et al, 2011). We have observed increased vacuolar accumulation coinciding with decreased plasma membrane abundance of PIN2‐derived GFP signal after 24‐h treatment with l‐Kynurenine. Importantly, this effect could be reversed by co‐incubation with auxin (Figure 6H–K).

As a complementary approach, we have genetically reduced the transcriptional auxin signalling. We have used a HS::axr3‐1 that expresses a stabilized allele of IAA17 after heat shock, resulting in a strong dominant repression of SCFTIR1/AFB‐regulated transcripts (Knox et al, 2003). Importantly, while PIN2 transcript levels were unaffected (Figure 7A), heat shock diminished PIN2 levels in membrane protein fractions (Figure 7B) as revealed by western blot analysis. Consistently, heat shock caused an increase in vacuolar PIN2‐GFP fluorescence signal along with the decrease in the fluorescence levels at the plasma membrane and caused root agravitropism (Figure 7C–E; Robert et al, 2010). These data imply that the genetic interference with SCFTIR1/AFB auxin signalling promotes PIN protein degradation. We have also analysed the stability of PIN2 protein in decapitated HS::axr3‐1 seedlings. We could not observe an additive effect of decapitation on vacuolar targeting of PIN2 protein in this genetic background (see Supplementary Figure 18). This suggests that increased vacuolar targeting (and degradation as shown by western blot analysis) of PIN2 efflux carrier triggered by the removal of cotyledons and shoot apical meristem is most likely caused by the changes in auxin levels rather than by possible secondary effects of tissue wounding like changes in cytokinin or jasmonate activity (Crane and Ross, 1986; Wasternack, 2007; Sun et al, 2009; Marhavý et al, 2011).

Thus, decreasing auxin levels or interfering with SCFTIR1/AFB auxin signalling leads to destabilization of PIN2 from the plasma membrane and higher rate of its vacuolar targeting. Overall, our data suggest that both the auxin decrease below optimal as well as increase above optimal levels can destabilize PIN proteins at the plasma membrane and, subsequently, induce PIN trafficking to the vacuole for degradation. Hence, ‘optimal’ auxin levels are required to stabilize PIN2 proteins for their action in gravitropic response.
Discussion
Dual regulation of PIN vacuolar targeting and degradation by auxin levels
The data presented in this study indicate that both, a prolonged increase or decrease in cellular auxin levels induce targeting of PIN auxin transporters (Petrášek et al, 2006) to the vacuole, thereby regulating the abundance of the auxin carriers at the plasma membrane. It appears that an ‘optimal’ auxin concentration is required to maintain PIN protein levels and thus auxin transport capacity at the plasma membrane. These effects of opposite auxin concentrations on PIN trafficking to the vacuole apparently depend on the canonical auxin signalling pathway, involving auxin‐dependent degradation of Aux/IAA transcriptional repressor proteins (Kepinski and Leyser, 2005; Dharmasiri et al, 2005a). Given the known outlines of the PIN subcellular trafficking (Kleine‐Vehn and Friml, 2008), auxin acts most likely in the regulation of the balance between recycling of PIN proteins back to the plasma membrane versus trafficking to the vacuole, possibly by influencing these trafficking pathways or PIN sorting between them. The WEAK AUXIN RESPONSE1 WXR1/RUS2 protein might play a role in the auxin‐mediated decision between PIN recycling and vacuolar targeting since the corresponding mutant shows defects in both transcriptional auxin response and PIN turnover (Ge et al, 2010). How the same outcome is achieved by two seemingly opposite signals is unclear. Different sets of proteins transcriptionally regulated by different auxin levels might possibly target different subcellular trafficking processes. Such a notion can be supported by the results of microarray experiment in which transcription profiling was analysed in response to exogenous auxin and in conditional axr3 auxin signalling mutant (http://www.ebi.ac.uk/arrayexpress/experiments/E-MEXP-3283). These experimental conditions can be considered as increased and decreased auxin signalling environment, respectively.
Alternatively, different AFB auxin receptors might respond to different auxin levels in various cells and might have opposite effects on the downstream signalling, as recently suggested for AFB4 (Greenham et al, 2011). It is possible that the SCFTIR1/AFB signalling pathway induces downstream effectors that post‐transcriptionally modify PIN proteins. Similarly to the PIN phosphorylation by the Ser/Thr protein kinase PINOID (PID) that directly affects the PIN polar targeting (Friml et al, 2004; Michniewicz et al, 2007; Kleine‐Vehn et al, 2009; Huang et al, 2010; Zhang et al, 2010), other post‐transcriptional modifications, such as PIN ubiquitination (Abas et al, 2006; Leitner et al, 2012), might change the subcellular sorting and trafficking of PIN proteins, leading to their preferential targeting to and degradation in vacuoles. Finally, SCFTIR1/AFB signalling can potentially affect a more general trafficking regulator since not only PIN proteins but also other plasma membrane proteins (although less effectively) are rerouted to the vacuole upon fluctuations in cellular auxin levels. Such a master regulator of vacuolar targeting could be subject to proteasome modifications and in turn direct post‐translational modifications of PINs and other proteins. Such a hypothesis would integrate the involvement of both proteasomal and vacuolar lytic degradation in the regulation of PIN abundance. It would also clarify why PIN degradation is impaired in the presence of proteasome inhibitor (Abas et al, 2006) given the fact that the proteasome complex targets mainly soluble and not membrane proteins (Vierstra, 2009). Future work will address which trafficking pathways are targeted by this processes and whether an increase or a decrease in cellular auxin levels would activate a common or distinct pathway.
Auxin differentially regulates PIN2‐mediated fluxes during root gravitropic response
Auxin can modify its own transport by regulating PIN transcription (Peer et al, 2004; Heisler et al, 2005; Vieten et al, 2005; Scarpella et al, 2006) and inhibiting PIN internalization from the plasma membrane (Paciorek et al, 2005; Robert et al, 2010; Chen et al, 2012; Lin et al, 2012). Here, we propose an integration of another auxin‐regulated trafficking process, namely PIN turnover as a substantial element of the multilevel control mechanisms by which auxin orchestrates root re‐orientation in response to gravity stimulus. Our observations indicate that protein degradation is a significant part of the PIN regulatory network, particularly important during later phases of root gravitropic response.
It has been previously shown that root re‐orientation to horizontal position results in auxin transport along the gravity vector leading to an establishment of temporal lateral auxin gradient across the organ (Luschnig et al, 1998; Swarup et al, 2005). Our study suggests that this gradient does not only involve an increase in the auxin response at the lower side but also its decrease at the upper side of the gravistimulated root. This apparent auxin depletion at the upper side coincides with PIN destabilization at the plasma membrane most likely due to enhanced trafficking to the vacuole for degradation (Abas et al, 2006; Kleine‐Vehn et al, 2008b). As a consequence of this feedback regulation, lowered auxin transport capacity along the upper side of the root leads to decreased cellular auxin levels to ‘below optimal’. Fluctuations in auxin level would then trigger changes in rates of cellular elongation (Barbier‐Brygoo et al, 1991; Ishikawa and Evans, 1993; Evans et al, 1994, reviewed in Perrot‐Rechenmann, 2010) eventually leading to a differential growth between two sides of the bending root (Zieschang and Sievers, 1991; Ishikawa and Evans, 1993) according to the classical Cholodny‐Went hypothesis (Firn et al, 2000; Blancaflor and Masson, 2003). Interestingly, in the same developmental context similar cellular output (PIN2 degradation) although separated spatially and shifted temporarily is achieved by elevated auxin levels at lower side of the root. We are speculating that the transient stabilization of PIN2 observed there is the result of inhibitory auxin effect on clathrin‐mediated PIN internalization (Paciorek et al, 2005; Robert et al, 2010; Chen et al, 2012; Lin et al, 2012). Elevated auxin levels also inhibit expansion of epidermal cells in the elongation zone at the lower side of the root. The subsequent decrease in PIN2 levels could be the result of the promoting effect of prolonged increased auxin levels on PIN2 degradation proceeding with slower kinetics than that of endocytosis (Robert et al, 2010). The interplay between these two auxin‐mediated effects running with different kinetics would ultimately lead to re‐establishment of the evenly distributed auxin flux on both sides and consequently vertical growth of the root.
The studies presented in this work address specifically a part of events following gravistimulation, namely how auxin influences the turnover of PIN2 thus regulating auxin flow from the place of gravity perception (root tip) to the responsive tissues in the elongation zone. These events follow the initial establishment of auxin asymmetry in the root tip presumably mediated by the gravity‐induced relocation of PIN3 and PIN7 in the root columella cells (Friml et al, 2002; Harrison and Masson, 2008; Kleine‐Vehn et al, 2010). Beside PIN action in auxin transport, gravity‐induced auxin translocation requires a crucial involvement of auxin influx machinery (Bennett et al, 1996; Marchant et al, 1999) and ATP‐energized auxin transport utilizing ABCB transporters (Geisler et al, 2005; Blakeslee et al, 2007; Lewis et al, 2007; Mravec et al, 2008). The model of auxin action on auxin transport activity must be also integrated with other gravity‐induced cellular signalling processes; many of which involve signals other than auxin (Evans and Ishikawa, 1997; Moulia and Fournier, 2009). Finally, it is tempting to speculate that the auxin effect on PIN protein degradation besides regulating root gravitropism might contribute to other processes, such as the auxin transport‐mediated auxin maxima establishment during de novo organ formation (Benková et al, 2003; Reinhardt et al, 2003; Heisler et al, 2005; Vernoux et al, 2010) where PIN degradation has been recently shown to play an important role (Marhavý et al, 2011).
Regulation of PIN activity at the plasma membrane by different auxin signalling pathways
Auxin has been demonstrated to influence its own efflux in a dual manner by either increasing or decreasing the incidence of PIN auxin transporters at the plasma membrane. These effects are achieved by the inhibition of PIN endocytosis (Paciorek et al, 2005; Robert et al, 2010) or promotion of PIN degradation (Sieberer et al, 2000; Vieten et al, 2005; Abas et al, 2006; present work), respectively. The auxin inhibitory effect on PIN endocytosis was attributed to the nuclear auxin signalling pathway that depends on the SCFTIR1/AFB auxin receptors (Pan et al, 2009). This was, however, most likely an erroneous interpretation as it did not account for the fact that trafficking inhibitor BFA (that was used to indirectly visualize rate of PIN internalization) targets besides PIN recycling to the plasma membrane also its trafficking to the vacuole (Peyroche et al, 1999; Sata et al, 1999; Geldner et al, 2003; Kleine‐Vehn et al, 2008b; Robert et al, 2010). It seems, therefore, that authors unintentionally addressed a process of vacuolar trafficking rather than the effect on endocytosis. Several recent reports strongly support the idea that the auxin effect on endocytosis does not depend on SCFTIR1/AFB machinery but utilizes a direct, non‐transcriptional ABP1‐mediated signalling pathway that targets a general process of clathrin‐mediated endocytosis (Robert et al, 2010; Chen et al, 2012; Lin et al, 2012; Nagawa et al, 2012). It has been proposed that ABP1 might sense auxin in the extracellular space where a small portion of the protein was detected (Jones and Herman, 1993; Bauly et al, 2000) and where ABP1 is active in terms of auxin response (Barbier‐Brygoo et al, 1996; Gehring et al, 1998; Steffens et al, 2001). Thus, cell surface active ABP1 could activate a rapid signalling pathways depending on ROP GTPases to inhibit clathrin‐mediated endocytosis without involvement of nuclear auxin signalling (Robert et al, 2010; Chen et al, 2012; Lin et al, 2012; Nagawa et al, 2012).
In this work, we provide additional data to clarify the involvement of SCFTIR1/AFB pathway in PIN endocytosis versus vacuolar trafficking. We show by independent approaches that targeting of PINs to the vacuole for degradation is controlled by SCFTIR1/AFB mechanism explaining the results of Pan et al (2009). Our results support a model, in which auxin regulates its own flux via distinct signalling pathways, which are controlling processes with different kinetics and specificities. This multilevel mechanism for the regulation of PIN‐dependent, directional auxin flux presumably contributes to the adaptive plasticity of plant development.
Materials and methods
For the description of reagents, drug application, experimental conditions, image processing, statistical analysis and quantification index, see Supplementary data at http://www.embojournal.org.
Plant material and growth conditions
All Arabidopsis thaliana mutants and transgenic lines employed in this study are in the Columbia (Col‐0) background and have been described previously: PIN2::PIN2‐GFP (Xu and Scheres, 2005), DR5rev::3XVENUS‐N7 (Heisler et al, 2005), DR5rev::GFP (Friml et al, 2003), DII‐VENUS (Brunoud et al, 2012), BRI1::BRI1‐GFP (Russinova et al, 2004), 35S::PIP2‐GFP (Cutler et al, 2000), 35S::PIN2‐EosFP (Dhonukshe et al, 2007), tir1afb1, tir1afb2, tir1afb3, tir1afb1,2,3 (Dharmasiri et al, 2005b), HS::axr3‐1 (Knox et al, 2003), RPS5»iaaM (Weijers et al, 2005), abp1‐5 (Xu et al, 2010) and tir1‐1 (Ruegger et al, 1998), ARF promoter::GFP lines (Rademacher et al, 2011), arf2‐8 (Ellis et al, 2005), arf6‐2 (Nagpal et al, 2005), arf19‐1 (Okushima et al, 2005), arf7arf19 (Wilmoth et al, 2005), arf10arf16 (Wang et al, 2005). Surface‐sterilized seeds were sown on half‐strength Murashige and Skoog (0.5 MS) agar plates and stratified for 2 days at 4°C. Plants were grown on vertically oriented plates under continuous light conditions at 22°C for 4–5 days.
Root gravitropism assay
Arabidopsis 4‐day‐old seedlings grown in continuous light conditions were covered with a layer of solid 0.5 MS medium and placed in Lab‐Tek® II Chambered Coverglass (Nalge Nunc International). Chambers were gravistimulated by 90° rotation and transferred to darkness 2 h prior CLSM analysis. Ten Z‐sections spaced ∼1 and 4.5 μm apart for PIN2 and DR5 promoter analysis, respectively, were collected in the median root section. Single pictures were subsequently combined into the maximum intensity projection. For the specific quantification method used in each experiment, please see Quantification Index.
Trans‐activation experiment
RPS5::GAL4 and UAS::iaaM (both in wild‐type Col‐0 background) were used for the cross. F1 progeny of RPS5::GAL4 × UAS::iaaM was crossed with a homozygous PIN2::PIN2‐GFP (eir1‐1). F1 generation was analysed. F1 generation of the PIN2::PIN2‐GFP (eir1‐1) × Col‐0 was used as a control.
Heat‐shock induction
All the CLSM analyses using HS::axr3‐1 expressing line were performed after 2 h of heat‐shock induction at 37°C followed by 3 h incubation in continuous light at 22°C. In experiment visualized in Supplementary Figure 18, following decapitation, seedlings were subjected to three subsequent heat‐shock inductions spaced over a total time of 16 h. Rationale was to maintain reasonable expression of mutated axr3‐1 gene over this period of time.
Immunodetection and microscopy
Whole‐mount immunolocalization in Arabidopsis roots was done as described previously (Sauer et al, 2006). The rabbit anti‐PIN1 (Paciorek et al, 2005) and rabbit anti‐PIN2 (kindly provided by C Luschnig) primary antibodies were used at a dilution of 1:1000 and fluorochrome‐conjugated anti‐rabbit‐Cy3 secondary antibody (Dianova) was diluted 1:600.
Membrane protein extraction and gel blotting analysis
Approximately 15 mg of seeds was germinated vertically on solid 0.5 MS medium. Seven DAG seedlings were subjected to the treatment. Roots were cut at fixed distance from the root tip and collected for analysis. Microsomal membrane fraction was isolated as described previously (Abas and Luschnig, 2010). Equal amount of proteins was separated by 10% SDS–Urea PAGE as described (Abas et al, 2006) followed by either Coomassie Briliant Blue staining (for loading control) or blotting to ECL membranes (GE Healthcare). The membranes were subsequently treated with affinity‐purified anti‐rabbit PIN2 antibody (overnight at 4°C) and ECLTM‐anti‐rabbit IgG, horseradish peroxidase (GE Healthcare; 1:10 000) (1 h at RT). Besides the specific band detected for PIN2 (around 70 kDa), other peptides were detected on the western blot by PIN2 antibody. These are most likely conjugates or metabolites of PIN2 detected together with native protein, as reported and commented in Abas et al (2006), Figure 2 legend. The immunoreactive signals were detected using the ECL detection system (GE Healthcare).
Quantitative RT–PCR
Total RNA was extracted with the RNeasy kit (Qiagen). Poly(dT) cDNA was prepared from total RNA with Superscript III (Invitrogen). Quantitative RT–PCR was done with LightCycler 480 SYBR Green I Master reagents (Roche Diagnostics) and a LightCycler 480 Real‐Time PCR System (Roche Diagnostics). Targets were quantified with specific primer pairs designed with Beacon Designer 4.0 (Premier Biosoft International). Data were analysed with qBASE v1.3.4 (Hellemans et al, 2007). Expression levels were normalized to the non‐auxin‐responsive genes CDKA (At3g48750), EEF (At5g60390) and TUB2 (At5g62690). For the presentation, TUB2 reference gene was used. For primer sequences, see Supplementary Table 1.
Conflict of Interest
The authors declare that they have no conflict of interest.
Acknowledgements
We thank Mark Estelle (University of California, San Diego, CA, USA), Ottoline Leyser (University of Cambridge, Cambridge, UK), Christian Luschnig (University of Applied Life Sciences and Natural Resources, Vienna, Austria), Remko Offringa (Leiden University, Leiden, The Netherlands), Ben Scheres (University of Utrecht, Utrecht, The Netherlands), Chris Somerville (University of California, Berkeley, CA, USA), Sacco de Vries (Wageningen University, Wageningen, The Netherlands) and Dolf Weijers (Wageningen University, Wageningen, The Netherlands) for sharing published materials and Martine De Cock for help in preparing the manuscript. This work was supported by the Odysseus program of the Research Foundation Flanders (to JF), the Vienna Science and Technology fund (WWTF) (to JK‐V) and Vetenskapsråde and VINNOVA (to SR). SV is a Post‐doctoral Fellow of the Research Foundation‐Flanders.
Author contributions: JK‐V, SR and JF designed the research. PB, SR, JK‐V, SV, WG and UK performed the experiments. PB analysed the data. JF, SR and PB wrote the article.
Supporting Information
Supplementary Data (PDF document, 9.23 MB)
Review Process File (PDF document, 244.22 KB)
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Volume 32,Issue 2,Jan 2013Dettifoss waterfall, Jökulsárgljúfur National Park, Iceland. 45 metres high and 100 metres wide, Dettifoss is Europe's most powerful waterfall. Up to 500 cubic metres of water thunder over the edge each second, sending up a plume of spray that can be seen from more than a kilometre away. The photograph was taken by Miguel Mano of the ICGEB, Trieste, Italy.
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Received: 27 September 2012
Accepted: 19 October 2012
Published online: 4 December 2012
Published in issue: 23 January 2013
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