New Results

Autophagy-mediated CTR1 turnover orchestrates the reciprocal interaction between autophagy and ethylene signaling

Hye Lin Park, Weiwei Zhang, Yuan-Chi Chien, Chanung Park, Gyeong Mee Yoon
doi: https://doi.org/10.1101/2024.07.31.606019

ABSTRACT

The phytohormone ethylene and autophagy are crucial for plant adaptation to various environmental stresses, yet the integration of these signaling networks into stress responses is not fully understood. Here, we report that ethylene signaling and autophagy reciprocally regulate each other through Constitutive Triple Response 1 (CTR1), a negative regulator of ethylene signaling. Autophagy facilitates the turnover of the CTR1 protein, which interacts with the key autophagy-related protein ATG8 as autophagic cargo. Impaired autophagy attenuates ethylene responses. Conversely, ethylene-insensitive mutants exhibit enhanced autophagic flux, while a constitutive ethylene response mutant is hypersensitive to carbon starvation stress, which induces autophagy. This suggests that ethylene suppresses autophagy during carbon limitation. We further elucidated that ethylene receptors with a receiver domain play a primary role in modulating autophagy, while receptor kinase activity is not essential. Our findings uncover that the autophagic control of CTR1 degradation allows reciprocal cross-regulation between autophagy and ethylene transduction cascades, optimizing stress responses and resilience.

Main text

When stressed, plants reprogram their cellular, biochemical, and physiological features to acclimate to prevailing stress conditions. During this process, it is critical to keep their stress responses within optimal threshold levels—strong enough to withstand yet restrained enough to recover when the stress subsides. Ethylene signaling and autophagy, a self-eating cellular degradation and recycling process, are two well-known cellular pathways largely responsible for plant acclimation to a broad range of stresses1–7. However, despite well-established signaling pathways and physiological outputs, there is still a limited understanding of how these two pathways optimize the levels of stress responses to confer adequate stress acclimation and resilience.

Ethylene is a key plant hormone that enables plants to cope with environmental stresses. It also plays an important role in plant growth and development by regulating fruit ripening, seedling germination, senescence, and root hair formation7. Ethylene is perceived by ethylene receptors located on the endoplasmic reticulum (ER) membrane8, 9. At low levels of ethylene, these receptors activate Constitutive Triple Response 1 (CTR1), a Raf-like protein kinase, which phosphorylates Ethylene Insensitive 2 (EIN2), a key positive regulator of ethylene signaling. Phosphorylation of EIN2 by CTR1 leads to its inactivation, likely through degradation mediated by EIN2-targeting protein 1 (ETP1) and ETP210–14. In the presence of ethylene, the ethylene receptors and CTR1 proteins are inactivated, which results in decreased phosphorylation of EIN2. This triggers the cleavage and transport of the C-terminal domain of EIN2 (EIN2-CEND) into the nucleus and processing body (P-body), where EIN2-CEND indirectly activates the EIN3 transcription factor and sequesters mRNA of EIN3-binding F-Box proteins (EBFs), respectively, to activate ethylene responses15, 16. Similarly, CTR1 is released from the ER to the cytoplasm upon ethylene binding to the receptors. CTR1 then translocates into the nucleus, where it enhances ethylene responses by stabilizing EIN3 proteins through direct interaction with EBFs6. The CTR1-EBF interactions presumably suppress EBF function, although the underlying mechanism is unknown.

The protein turnover of several ethylene signaling components, such as EIN2, EIN3/EIN3-like (EIL), and EBFs, is relatively well-established12, 14, 17–19. However, the mechanism governing CTR1 turnover remains elusive despite its critical role in the ethylene signaling pathway. Ethylene and 1-aminocyclopropane-1-carboxylic acid (ACC) have been shown to increase CTR1 protein levels in Arabidopsis plants, which facilitates the nuclear translocation of CTR1 and turns it into a positive regulator for the ethylene response6, 20. This suggests that cellular CTR1 levels, along with EIN2 and EIN3/EILs, play an important role in modulating plant stress responses. Although the mechanism underlying ethylene-induced stabilization of CTR1 is unknown, these results imply that ethylene negatively regulates the mechanism governing CTR1 protein turnover, thus contributing to optimal levels of ethylene responses in plants.

The diverse roles of ethylene in modulating various aspects of plant growth and mediating adaptive responses to biotic and abiotic stresses are underpinned by complex signaling crosstalk between ethylene and other cellular processes. One such process emerging as a key regulator of ethylene signaling is autophagy, a catabolic mechanism employing vacuolar degradation and recycling of cellular components to ensure plant survival under stress. Ethylene and autophagy regulate several key developmental transitions and stress response pathways, including senescence, hypoxia, drought tolerance, and nutritional stress adaptation4, 5, 21–29. This implies significant convergent signaling and physiological crosstalk between the two pathways. However, despite several potential nodes of interaction, the molecular mechanism underlying ethylene-autophagy crosstalk remains largely obscure.

In this study, we elucidate the reciprocal interaction between ethylene signaling and autophagy through the autophagic turnover of CTR1, revealing a crosstalk node integrating these pathways. Using autophagy-defective and ethylene signaling mutants, we demonstrate that autophagy positively regulates ethylene responses by destabilizing CTR1 proteins.

Conversely, ethylene negatively regulates autophagy by destabilizing autophagosomes and modulating the expression of autophagy-related (ATG) genes and Target of Rapamycin (TOR) signaling components under carbon starvation. Furthermore, ethylene receptors with a receiver domain are crucial for regulating autophagic responses independently of kinase activity.

Together, our work demonstrates how plants coordinate the control of CTR1 turnover to dynamically co-regulate the ethylene and autophagy pathways, optimizing stress responses and resilience to nutritional deficiencies while sustaining growth.

RESULTS

CTR1 protein levels are regulated by autophagy

CTR1 plays a critical role in modulating responsiveness to ethylene in multiple developmental and physiological responses6, 30–33. However, the mechanism regulating the level of CTR1 protein is unknown. Given potential links between autophagy and ethylene function, we examined whether autophagy plays a role in CTR1 protein turnover under carbon starvation, a well-known environmental condition for inducing autophagy22, 34. We transferred 7-day-old light- grown wild-type (WT) Arabidopsis thaliana seedlings to sucrose-deficient media and incubated them in darkness. Within 24 hours, the steady-state levels of CTR1 protein were substantially decreased and undetectable by 72 hours. However, treatment with concanamycin A (ConA), an inhibitor of vacuolar degradation, not only blocked CTR1 reduction but also led to gradual accumulation (Fig. 1a)34. Unlike in WT seedlings, genetic disruption of autophagy in the atg7-2 mutant seedlings abolished the CTR1 decrease under these conditions (Fig. 1a)35. Notably, we observed that CTR1 protein levels also started to reduce after 4 days of dark incubation in the atg7-2 seedlings, likely attributed to the intrinsic chlorosis phenotype of the atg7-2 mutant.

Figure 1.
Figure 1. CTR1 is degraded by the autophagy pathway in Arabidopsis.

a. Concanamycin A inhibits CTR1 degradation. Seven-day-old WT and atg7-2 mutant seedlings grown in light were transferred to sucrose-deficient liquid media with or without concanamycin A (ConA, 0.5 μM). The seedlings were then incubated in darkness for the indicated times to induce carbon starvation. Total proteins were extracted and analyzed by immunoblotting using anti-CTR1 antibodies. Anti-actin antibodies were used as a loading control. b. Time course of free GFP release from GFP-CTR1 during carbon starvation. ctr1-2 and atg7-2 seedlings expressing GFP- CTR1 were carbon starved for the indicated times. Total protein extracts were immunoblotted with anti-GFP antibodies. Arrows indicate the GFP-CTR1 fusions and the free GFP released upon degradation. c. Delivery of GFP-CTR1 to vacuoles is blocked in atg7-2. Five-day-old light- grown WT and atg7-2 seedlings expressing GFP-CTR1 were incubated in a sucrose-free medium with ConA under light or dark conditions for 16 hours. Root cells were examined for autophagic vesicles by confocal microscopy. Scale bars, 10 μm. One-way ANOVA with Tukey’s post-hoc test was performed, and the letters denote statistically significant differences between groups, p < 0.0001 (n ≥ 20 cells from five to six seedlings for each treatment) d. GFP-CTR1 colocalizes with mCherry-ATG8a. Seedlings co-expressing GFP-CTR1 and mCherry-ATG8a were incubated in a sucrose-free medium with ConA for 16 hours. Root cells were imaged by light sheet fluorescence microscopy to visualize the intracellular colocalization of GFP-CTR1 and mCherry-ATG8a. As a negative control, random colocalization was measured by rotating one image 180 degrees and repeating the analysis on the same image pair, assessing the expected colocalization due to chance. Scale Bar, 10 μm. One-way ANOVA with Tukey’s post- hoc test was performed, and the letters denote statistically significant differences between groups, p < 0.0001 (n ≥ 5 seedlings). Two different sections in the cells of each seedling were imaged and measured. e. CTR1 interacts with ATG8a in Nicotiana benthamiana. Leaves of N. benthamiana were co-infiltrated with agrobacteria carrying plasmids for nYFP-CTR1, cYFP- ATG8a, or cYFP-CIP8 (negative control), and bZIP-RFP (expression control). After 3 days, the reconstitution of YFP signals was observed by confocal microscopy, indicating CTR1-ATG8a interaction. f. Co-immunoprecipitation of CTR1 and ATG8a. Leaves of N. benthamiana were infiltrated with agrobacteria expressing GFP-CTR1 alone, myc-ATG8a alone or both GFP-CTR1 and myc-ATG8a. Total protein extracts were incubated with anti-GFP magnetic beads to pull down GFP-CTR1. The immunoprecipitants were subsequently immunoblotted with anti-GFP and anti-Myc antibodies to detect the co-immunoprecipitation of CTR1 and ATG8a.

Additionally, we utilized GFP-cleavage assays to evaluate the autophagic turnover of GFP-CTR1 in WT and atg7-2 mutant (Fig. 1b)34, 36, 37. This assay determines the vacuolar delivery of GFP fusion proteins by quantifying the accumulation of protease-resistant free GFP in vacuoles.

Under carbon starvation, we observed escalating levels of liberated GFP from the GFP-CTR1 while descending levels of GFP-CTR1 in a ctr1-2 mutant background. However, carbon starvation-triggered GFP release was absent in the atg7-2 mutants (Fig. 1b). These results demonstrate that CTR1 protein levels are regulated by autophagy-mediated trafficking and subsequent vacuolar degradation under carbon deficit stress.

As CTR1 protein levels are regulated by autophagy (Fig. 1a, 1b), we next examined if CTR1 is targeted to autophagosomes, the double-membraned autophagic vesicles that deliver cargos to the vacuole for degradation22. We used stable transgenic plants expressing GFP- CTR1 from its native promoter in a ctr1-2 null mutant background (CTR1p:GFP-CTR1/ctr1-2). Incubation of GFP-CTR1 seedlings with ConA increased the number of GFP-CTR1-containing vesicles after carbon starvation stress. However, the accumulation of GFP-CTR1 into vesicles was nearly eliminated in the atg7-2 mutant background, indicating that these vesicles are indeed autophagic vesicles (Fig. 1c). To confirm further that the ConA-dependent CTR1- containing vesicles are autophagic vesicles, we analyzed the intracellular colocalization of GFP- CTR1 with mCherry-ATG8a-containing autophagic vesicles in roots of stable transgenic Arabidopsis plants (Fig. 1d). Manders’ overlap coefficient values revealed that 56.6% of the GFP-CTR1 vesicles colocalized with mCherry-ATG8a vesicles, while 57.2% of mCherry-ATG8a autophagosomes colocalized with vesicles containing GFP-CTR1 (Fig. 1d)38. These results show that the GFP-CTR1 signal significantly colocalized with mCherry-ATG8a, consistent with the autophagic turnover of CTR1. Bimolecular fluorescence complementation (BiFC) and co- immunoprecipitation analyses further revealed that CTR1 interacts with ATG8a in plants (Fig. 1e, 1f), indicating that CTR1 is either a direct cargo of autophagy or acts as a selective autophagy receptor, mediating the degradation of unknown cargo. Notably, given that CTR1 does not contain a canonical ATG8-interacting motif (AIM) site, this result also suggests that CTR1 interacts with ATG8 through a non-canonical ATG8-binding site22, 39, 40. Additionally, CTR1 colocalizes with multiple ATG proteins involved in autophagosome formation, further indicating the functional link between CTR1 and autophagy (Supplementary Fig. 1).

Autophagy positively regulates ethylene responses in Arabidopsis by decreasing CTR1 protein levels

The autophagic turnover regulation of CTR1 protein suggests that autophagy modulates ethylene responses in plants. To elucidate the functional relationship between ethylene and autophagy, we examined the ethylene responsiveness of atg mutant seedlings under light and dark conditions. Ethylene and its biosynthetic precursor, ACC, have distinct effects on hypocotyl cell expansion in Arabidopsis seedlings, stimulating elongation in the light while inhibiting growth in darkness41, 42. By examining these morphological responses in atg mutants in response to ethylene or ACC, we aimed to determine whether disrupting autophagy perturbs ethylene responses, at least in the context of hypocotyl growth regulation. We selected six atg mutants (atg2, atg5, atg7, atg9, atg11, and atg18a), that are defective in autophagy43. In the absence of ACC, the hypocotyl lengths of all dark-grown atg mutants were significantly longer than those of WT seedlings (Fig. 2a). All etiolated atg mutants also exhibited a hyposensitive response to a broad range of ACC concentrations compared to the WT, displaying longer hypocotyls than the WT at different ACC concentrations (Fig. 2b). The measurement of ethylene biosynthesis in dark-grown atg seedlings showed comparable levels to the WT (Fig. 2c), suggesting that the altered hypocotyl growth observed in the atg mutants is likely due to an altered ethylene response rather than ethylene production. In the absence of ACC treatment, light-grown atg mutant seedlings exhibited hypocotyl lengths that were indistinguishable from those of WT seedlings. However, upon exposure to increasing concentrations of ACC, the responsiveness of these mutants to ACC was significantly diminished, leading to considerably shorter hypocotyls compared to WT seedlings (Fig. 2d, 2e). No difference in ethylene biosynthesis was observed in light-grown atg mutant seedlings compared to the WT (Fig. 2f).

Figure 2.
Figure 2. Autophagy positively regulates ethylene responses by modulating CTR1 levels.

a. atg mutants are less sensitive to ACC treatment in darkness. Seedlings were grown on MS medium with or without 10 μM ACC for 3 days in darkness and photographed. The graph presents quantification of the hypocotyl lengths of seedlings grown on MS without ACC. Scale bar, 5 mm. One-way ANOVA with Dunnett’s test was performed to determine statistical significance. Error bars, SD (n ≥ 38 seedlings). b. ACC dose-response curves for the hypocotyl length of 3-day-old dark-grown WT and atg mutant seedlings. Error bars, SD (n ≥ 38 seedlings for each ACC concentration). Two-tailed Student’s t-test. c. Etiolated atg mutant seedlings produced comparable levels of ethylene to the WT seedlings. Seedlings were grown on MS medium for 3 days in darkness, and the accumulated ethylene was measured using gas chromatography. Error bars, SD (n = 3 vials and n ≥ 216 seedlings in each vial). One-way ANOVA with Dunnett’s test was performed to determine statistical significance. d. atg mutants are less sensitive to ACC treatment in the light. Seedlings were grown on MS medium with or without 10 μM ACC for 5 days in the light and photographed. The graph presents the quantification of the hypocotyl lengths of seedlings grown on MS with 10 μM ACC. Scale bar, 5 mm. One-way ANOVA with Dunnett’s test was performed to determine the statistical significance. Error bars, SD (n ≥ 32 seedlings). e. ACC dose-response curves for the hypocotyl length of 5-day-old light-grown WT and atg mutant seedlings. Error bars, SD (n ≥ 34 seedlings for each ACC concentration). Two-tailed Student’s t-test. f. Light-grown atg mutant seedlings produced comparable levels of ethylene to the WT seedlings. Seedlings were grown on MS medium for 5 days in the light, and the accumulated ethylene was measured using gas chromatography. Error bars, SD (n = 4 vials and n ≥ 17 seedlings in each vial). One-way ANOVA with Dunnett’s test was performed to determine the statistical significance. g. atg mutant expressed higher levels of CTR1 protein compared to WT seedlings. Total protein extracts of 3-day-old etiolated and 5-day-old light-grown seedlings were immunoblotted using an anti-CTR1 antibody. An anti-actin antibody was used as a loading control. Rel. quantities represent the ratio of the intensity of the CTR1 band to actin band signals, and these values are expressed relative to the intensity of CTR1/actin in the WT, which was set to 1. h, i. Quantitative gene expression analysis of CTR1 and ERF1 in WT and atg mutant seedlings. Expression was normalized to an Actin2 control and is presented relative to the WT control. One-way ANOVA with Dunnett’s test was performed to determine statistical significance. Error bars, SD (n = 3 biological replicates).

Given that autophagy modulates the turnover of the CTR1 protein, the reduced ethylene sensitivity observed in atg mutants could partly stem from increased endogenous CTR1 protein levels, which results in an attenuation of ethylene responses. In support of this hypothesis, immunoblot analysis revealed significantly higher CTR1 protein accumulation in the atg mutant seedlings than in the WT under both light and dark growth conditions (Fig. 2g). However, transcript levels of CTR1 in the atg mutants exhibited either similar or reduced expression relative to those in WT seedlings under dark and light conditions, respectively (Fig. 2h). This suggests that the elevated CTR1 protein levels in atg mutants result from post-transcriptional effects due to impaired autophagic flux in these mutants. Supporting the enhanced CTR1 protein levels, both dark- and light-grown atg mutant seedlings expressed significantly lower levels of the ethylene-inducible ERF1 than WT seedlings (Fig. 2i)44.

Ethylene and autophagy have opposing effects on CTR1 turnover (Fig. 1)6, 20. Ethylene and its precursor ACC increase CTR1 levels, whereas autophagy promotes its protein turnover. To investigate whether autophagy inhibits the ethylene-induced stabilization of CTR1, thereby destabilizing CTR1, we examined the steady-state levels of CTR1 in response to increasing concentrations of ACC in atg5 and atg7 mutants, which are representative atg mutants with severe autophagy defects. Immunoblot analysis revealed that ACC gradually increases CTR1 levels in a concentration-dependent manner in WT seedlings, corroborating previous reports (Supplementary Fig. 2a)6, 20. The atg5 and atg7 mutants also exhibited a similar pattern of CTR1 accumulation in response to ACC, despite their significantly higher basal CTR1 levels compared to WT seedlings. This suggests that autophagy does not destabilize CTR1 by interfering with ethylene-induced CTR1 stabilization. Instead, an alternative pathway, likely the 26S proteasome-ubiquitin system, may regulate CTR1 turnover. Supporting this notion, treatment with Bortezomib (Btz), a proteasome inhibitor, prevented the degradation of endogenous CTR1 following cycloheximide treatment (Supplementary Fig. 2b), demonstrating that CTR1 protein turnover is also regulated by the 26S proteasome pathway. Collectively, these results demonstrate that autophagic flux positively regulates ethylene responses by degrading its key negative regulator, CTR1, thus potentiating ethylene signaling.

Ethylene signaling mutants exhibit altered carbon starvation-induced autophagic responses

Autophagy-deficient mutants exhibit reduced ethylene responses in both light- and dark-grown Arabidopsis seedlings (Fig. 2), suggesting the positive regulatory role of autophagy in the ethylene signaling-mediated developmental process. To further elucidate the functional link between autophagy and ethylene, we investigated whether the crosstalk between the two pathways is reciprocal. To this end, we assessed the autophagic responses of constitutive ethylene response and ethylene-insensitive mutants under carbon starvation. The constitutive ethylene response mutant ctr1-2 showed a hypersensitive response to carbon starvation, exhibiting severe chlorosis compared to WT and failing to recover in the light (Fig. 3a). The introduction of a CTR1 genomic transgene with the CTR1 promoter driving a GFP-CTR1 coding fusion complemented the carbon starvation hypersensitivity phenotype of ctr1-2, restoring autophagic phenotypes to WT levels. The ctr1-2 mutant displayed significantly reduced total protein levels and levels of the large subunit of Rubisco (RbcL) seven days after starvation.

Figure 3.
Figure 3. The ethylene signaling pathway negatively regulates autophagic responses under carbon starvation.

a, Carbon starvation-induced senescence phenotypes of WT, ctr1-2, CTR1p:GFP-CTR1/ctr1-2, and atg7-2 mutant seedlings. Seven-day-old light-grown seedlings were grown on MS medium without sucrose and transferred to darkness for 7 days, followed by recovery in the light for 7 days (7-rec). b, Total cellular proteins from seedlings during carbon starvation were resolved in SDS-PAGE, stained with Coomassie brilliant blue (CBB), or immunoblotted using antibodies against RbcL and actin (loading control). c, Carbon starvation- induced senescence phenotypes of WT, ein2-5, EIN2p:EIN2/ein2-5, ein2-5atg7-2, and atg7-2 mutant seedlings. d. Total cellular proteins from seedlings during carbon starvation were resolved in SDS-PAGE, stained with CBB, or immunoblotted using antibodies against RbcL and actin (loading control). e. Autophagosome formation is altered in ethylene signaling mutants.

Five-day-old light-grown seedlings expressing GFP-ATG8a in WT, ctr1-2, or ein2-5 were incubated in a sucrose-free medium with and without ConA under light or dark conditions for 16 hours. Root cells were examined for autophagic vesicles by confocal microscopy. Scale bars, 10 μm. One-way ANOVA with Tukey’s post-hoc test was performed, and different letters denote statistically significant differences between groups, p ≤ 0.0007. (n ≥ 20 cells from five to six seedlings for each treatment). f. Time course of free GFP release from GFP-ATG8a during carbon starvation. WT, ctr1-2 and ein2-5 seedlings expressing GFP-ATG8a were carbon starved for the indicated times. Total protein extracts were immunoblotted with anti-GFP antibodies. Arrows indicate the GFP-ATG8a fusions and the free GFP released upon degradation. Relative quantities represent the ratio of the GFP-ATG8a band intensity to the PBA1 band intensity. For each genotype, these ratios are normalized to the GFP-ATG8a/PBA1 ratio in the corresponding no-carbon starvation control, which is set to 1. g. The steady-state levels of GFP-ATG8a in WT, ctr1-2, and ein2-5 mutants. Total protein extracts of seedlings after the indicated starvation were immunoblotted using anti-GFP and anti-PBA1 antibodies. h. Quantification of GFP-ATG8a bands during starvation with and without ConA. Relative quantities of GFP-ATG8a bands are represented as the ratio of the GFP-ATG8a band intensity to PBA1 band intensity. For each genotype, these ratios are normalized to the GFP- ATG8s/PBA1 ratio in the corresponding no starvation control, which is set to 1. Error bars, SD (3 biological replicates). i, Quantitative gene expression analysis of ATGs, TOR, Raptor1b, and LST8-1 genes in WT, ctr1-2, and ein2-5 mutant seedlings. Expression was normalized to an Actin2 control and is presented relative to the WT control. One-way ANOVA with Tukey’s post- hoc test was performed, and the different letters denote statistically significant differences between groups. Error bars, SD (n = 4 biological replicates).

These defects were fully alleviated in the complementation lines harboring the CTR1 transgene, with total protein and RbcL levels restored to WT levels (Fig. 3b). In contrast, the ethylene- insensitive ein2-5 mutant displayed significantly delayed leaf chlorosis, which was sustained even after nine days in darkness (Fig. 3c). The ein2-5 mutant continued to express higher levels of total cellular proteins and RbcL protein compared to WT under carbon starvation.

Complementation of ein2-5 by expressing a genomic EIN2 transgene rescued the carbon starvation insensitivity of ein2-5 to WT levels (Fig. 3c, 3d), confirming that EIN2 is responsible for the carbon starvation-insensitive phenotypes of ein2-5. Furthermore, introducing an atg7 mutation into ein2-5 (ein2-5atg7-2 double mutant) inhibited the carbon starvation insensitivity of ein2-5, positioning autophagy downstream of ethylene signaling (Fig. 3c, d). Similar to ein2-5, the ethylene-insensitive mutants ein3eil1 and etr1-1 exhibited hyposensitive carbon starvation responses and maintained higher levels of total cellular proteins and RbcL compared to WT seedlings under carbon starvation conditions (Supplementary Fig. 3).

Ethylene signaling negatively regulates autophagy likely through destabilizing autophagosomes and modulating transcription

The dynamic engulfment and delivery of cellular components to the vacuoles via autophagosomes is a key process in autophagy22. To investigate the influence of ethylene signaling on autophagosome formation in response to carbon starvation, we examined this process in ein2-5 and ctr1-2 mutants using GFP-ATG8a reporter lines expressing GFP-ATG8a in the WT, ctr1-2, or ein2-5 mutant backgrounds. We selected ein2-5, ctr1-2, and WT lines with similar GFP-ATG8a protein levels for the carbon starvation experiments (Supplementary Fig. 4). Carbon starvation treatment of 35S:GFP-ATG8a/WT seedlings in the presence of ConA led to an elevated number of GFP-ATG8a-labeled autophagosomes compared to the ConA untreated control. This increase in autophagosome formation was even more pronounced in 35S:GFP-ATG8a/ein2-5 seedlings under the same starvation conditions compared to the WT (Fig. 3e). Conversely, 35S:GFP-ATG8a/ctr1-2 mutants exhibited only a marginal rise in the quantity of GFP-ATG8a autophagosomes when exposed to carbon starvation. (Fig. 3e).

Additionally, treatment with benzothiadiazole (BTH), a salicylic acid analog that induces autophagy, further increased autophagic flux in 35S:GFP-ATG8a/WT and 35S:GFP- ATG8a/ein2-5 seedlings after carbon starvation in the presence of ConA45, 46. However, this increase was not observed in 35S:GFP-ATG8a/ctr1-2 mutants (Supplementary Fig. 5). Next, we investigated whether ethylene signaling influences autophagic flux under carbon starvation conditions. Intriguingly, despite the contrasting effects on the number of GFP-ATG8a-labeled autophagosomes in the ctr1-2 and ein2-5 mutants compared to WT (Fig. 3e), GFP cleavage assays revealed a striking similarity in the release kinetics and amounts of free GFP among the mutants and WT after starvation (Fig. 3f). However, the steady-state levels of intact GFP- ATG8a significantly differed between ctr1-2 and ein2-5 compared to WT. In ctr1-2, GFP-ATG8a degraded faster upon starvation, while no apparent decrease in GFP-ATG8a levels was observed in ein2-5 (Fig. 3f). These results suggest that ethylene signaling does not control autophagosome degradation in the vacuole. Rather, the ctr1-2 mutation likely either accelerates ATG8 protein degradation or impedes autophagosome formation before vacuolar fusion, leading to fewer autophagosomes. To test these possibilities, we subjected 35S:GFP-ATG8a/WT, 35S:GFP-ATG8a/ctr1-2, and 35S:GFP-ATG8a/ein2-5 seedlings with or without ConA treatment during carbon starvation (Fig. 3g). ConA inhibits vacuolar (H+)-ATPases, thus preventing the degradation of autophagic cargos within vacuoles. In WT seedlings, ConA treatment maintained constant levels of GFP-ATG8a during starvation compared to untreated controls. However, in ctr1-2 mutants, GFP-ATG8a levels initially stabilized briefly but then rapidly degraded, similar to untreated samples. In contrast, ein2-5 mutants exhibited constant GFP-ATG8a levels during starvation, unaffected by ConA treatment (Fig.3g, 3h). To further investigate whether CTR1 directly regulates ATG8a stability, we examined the steady-state levels of endogenous ATG8a in ctr1-2 mutants and transgenic lines overexpressing GFP-CTR1 (CTR1ox) (Supplementary Fig. 6). Immunoblot analysis showed no significant differences in ATG8a levels among WT, ctr1-2, and CTR1ox lines with and without autophagy-inducer BTH treatment. This implies that CTR1 does not significantly impact overall ATG8a protein abundance. Instead, the role of CTR1 appears to be more specific to autophagosome stabilization or turnover. Collectively, these findings suggest that ethylene signaling inhibits autophagy by destabilizing autophagosomes before vacuolar fusion, while CTR1 counteracts this process.

To elucidate the molecular mechanism by which ethylene signaling negatively regulates autophagy at the transcriptional levels, we examined the transcriptional regulation of genes involved in autophagy and the TOR signaling pathway, which is a well-known negative regulator of autophagy47, 48. Under normal conditions, quantitative RT-PCR analysis showed similar transcript levels of various ATG genes in WT, ctr1-2, and ein2-5 mutants. However, upon carbon starvation, ctr1-2 mutants exhibited significantly higher expression of most ATG genes compared to WT and ein2-5, while WT and ein2-5 showed reduced or similar transcript levels of several ATG genes (Fig. 3i). Intriguingly, the transcript levels of TOR, RAPTOR1B, and LST8-1, which encode components of the TOR signaling pathway, were also significantly higher in ctr1-2 mutants after carbon starvation, with particularly notable increase in TOR and RAPTOR1B (Fig. 3i). The elevated expression of most ATG genes in ctr1-2 mutants likely represents a compensatory response to impaired autophagosome formation in an attempt to upregulate autophagy-related pathways to counteract the reduced autophagic activity. Concurrently, the upregulated expression of TOR signaling components supports the observed reduction in autophagic activity in ctr1-2 mutants. These results demonstrate that ethylene signaling also regulates autophagy in part through transcriptional regulation targeting key autophagy and TOR components, revealing an interplay between ethylene signaling, autophagy, and TOR pathway regulation.

Serine 924 of EIN2 plays a key role in regulating autophagic responses to carbon starvation

The constitutive ethylene response mutant ctr1-2 showed hypersensitive responses to carbon starvation. Thus, we further sought to elucidate the autophagic responses of constitutive nuclear localization of EIN2 and the role of Serine 645 (S654) and Serine 924 (S924), both of which lead to the cleavage of EIN2 and subsequent activation of ethylene signaling12, 13, 49. To investigate this, we examined carbon starvation phenotypes of transgenic Arabidopsis seedlings expressing WT full-length EIN2 (EIN2WT) or an EIN2 genomic fragment with S654A (EIN2S645A), S924A (EIN2S924A), or both substitutions (EIN2AA) under its native promoter in the ein2-5 background (Fig. 4a)12. The EIN2p-EIN2WT transgene alleviated the delayed leaf senescence of the ein2-5. In contrast, both EIN2p-EIN2AAand EIN2-EIN2S924A transgenes were unable to rescue the carbon starvation phenotypes of the ein2-5, instead conferring hypersensitive carbon starvation responses similar to those observed in ctr1-2 mutants. Notably, the EIN2p-EIN2S645A transgene fully complemented the phenotypes observed in the ein2-5 mutant. Under carbon starvation, EIN2p-EIN2S645Aplants maintained total protein and RbcL accumulation levels comparable to those of WT. In contrast, carbon-starved EIN2p-EIN2S924Aplants exhibited severe depletion of protein and RbcL levels, akin to the patterns seen in the atg7-2 mutant (Fig. 4b). These observations suggest that CTR1-mediated phosphorylation of EIN2 at S924, but not S645, plays a pivotal role in suppressing autophagy during carbon starvation conditions.

Figure 4.
Figure 4. CTR1-mediated EIN2 phosphorylation and ethylene receptors with a receiver domain play a role in autophagic responses during carbon starvation.

a. Carbon starvation-induced senescence phenotypes of WT, ein2-5, and ein2-5 complementation lines expressing EIN2WT, EIN2S645A, EIN2S924A, or EIN2AA under its native promoter. Seven-day-old light-grown seedlings were grown on MS medium without sucrose and transferred to darkness for 7 days, followed by recovery in the light for 7 days (7-rec). b. Total cellular proteins from seedlings during carbon starvation were resolved in SDS-PAGE, stained with CBB, or immunoblotted using antibodies against RbcL and actin (loading control). c. Carbon starvation- induced senescence phenotypes of WT and ethylene receptor mutants with or without a receiver domain or kinase activity. d. Total cellular proteins from seedlings during carbon starvation were resolved in SDS-PAGE, stained with CBB, or immunoblotted using antibodies against RbcL and actin (loading control).

Ethylene receptors with a receiver domain play a major role in regulating autophagy independently of kinase activity

CTR1 localizes to the ER as part of an ethylene receptor signaling complex through its direct interaction with the C-terminal domain of the receptors, which consists of a kinase domain with or without a receiver domain20, 50. The ER-localized ethylene receptors consist of five members: Ethylene Response 1 (ETR1), Ethylene Response Sensor 1 (ERS1), ETR2, ERS2, and EIN4. These receptors are classified into subfamilies I (ETR1 and ERS1) and II (ETR2, EIN4, and ERS2) based on the number of transmembrane domains and the homology of the histidine kinase domain 8, 51–55. Subfamily I members primarily exhibit histidine kinase activity, whereas subfamily II members may possess serine/threonine kinase activity. Of the five receptors, ETR1, ETR2, and EIN4 have a receiver domain as part of their C-terminus, while ERS1 and ERS2 lack this domain53, 56, 57. To investigate the role of the receiver domain and receptor histidine kinase activity in autophagy during carbon starvation, we examined the autophagic phenotypes of various ethylene receptor mutant combinations (Fig. 4c). The etr1-9ers1-3 mutant (lacking receptors with histidine kinase activity) and the ers1-2ers2-3 mutant (lacking receptors without a receiver domain) exhibited a WT-like chlorosis phenotype and reduced total protein abundance and RbcL levels in response to carbon starvation (Fig. 4d)58, 59. In contrast, the etr1-6etr2-3ein4- 4 mutant (lacking receptors with a receiver domain) exhibited hypersensitivity to starvation similar to ctr1-2 and atg7-2 mutants60. The etr2-3ers2-3ein4-4 triple mutant (lacking subfamily II receptors) also showed hypersensitivity, but with a lesser reduction in RbcL and total protein compared to the etr1-6etr2-3ein4-4 (Fig. 4d). These results suggest that receptors without a receiver domain are non-essential, while receptors with a receiver domain play a primary role in autophagy, regardless of kinase activity. Consistent with this, etr1-6 and etr1-7 loss-of-function mutants also displayed hypersensitive responses to carbon starvation (Supplementary Fig. 7)61, 62. Together, these results indicate that ethylene receptors with a receiver domain are essential for the autophagic response to carbon starvation, presumably via interaction with CTR1 through the receiver domain, while receptor kinase activity is nonessential63.

Discussion

CTR1 has long been recognized as a critical regulator of the ethylene signaling pathway. However, how its protein level is controlled has remained enigmatic. In this study, we demonstrate that autophagy degrades CTR1 proteins in response to carbon-deficient stress, identifying autophagy as a key regulator of ethylene responses by controlling CTR1 levels under stress conditions. We also reveal reciprocal crosstalk between ethylene signaling and autophagy mediated by autophagic CTR1 destabilization, wherein autophagy stimulates ethylene responses, while ethylene signaling inhibits autophagy. Our findings unveil that autophagy-mediated CTR1 degradation is a key mechanism linking and coordinating these two signaling networks that govern plant growth and stress responses.

While the biochemical and physiological stress responses of ethylene and autophagy pathways are well known, how plants balance these signaling pathways to control the magnitude and duration of stress responses is unclear. Preventing overshoot stress responses is crucial for avoiding plant damage and death while maintaining the capacity to resume growth after stress relief. Our finding that ethylene and autophagy reciprocally regulate each other through modulation of CTR1 levels provides insight into these balancing stress responses and proposes a model for the crosstalk between the two pathways (Fig. 5). When ethylene signaling becomes overactive, it suppresses autophagy, which is attenuated by the TOR kinase cascade under normal growth conditions. This suppression leads to the accumulation of CTR1, consequently alleviating further intensification of ethylene responses. Conversely, hyperactivated autophagy promotes the degradation of CTR1, releasing ethylene signaling from inhibition to restrain autophagic activity. Thus, CTR1 enables each pathway to shut off the other upon exceeding moderate thresholds while permitting balanced crosstalk to sustain optimal stress responses. Collectively, by leveraging CTR1 to modulate both pathways, ethylene signaling and autophagy may be able to activate protective stress responses while still preventing excessive responses that compromise the long-term viability of plants. This elegant homeostatic feedback loop modulates transient and defensive stress responses, followed by a coordinated recovery of growth homeostasis, unveiling molecular strategies to enhance environmental stress resilience in plants.

Figure. 5.
Figure. 5. Model for the reciprocal regulation between ethylene signaling and autophagy.

The survival of plants under stress conditions hinges on maintaining a delicate balance between mounting stress responses and preserving their recovery potential. Plants achieve this balance through the reciprocal regulation of ethylene signaling and autophagy with CTR1 acting as a crucial crosstalk node to prevent an overshoot of stress responses. By modulating CTR1 levels, plants can mitigate excessive activation of either pathway, thereby preventing an overshoot of stress responses while retaining the ability to recover once the stress has subsided. a. When ethylene signaling becomes overactivated, it suppresses autophagy, subsequently inhibiting the autophagy-mediated turnover of CTR1. The resulting accumulation of CTR1 then dampens the ethylene response, forming a negative feedback loop that mitigates the overshoot of ethylene signaling. b. Conversely, excessively activated autophagy further stimulates CTR1 turnover, leading to an inhibition of the CTR1-mediated constraint on ethylene signaling. The enhanced ethylene response then attenuates autophagy activity, ultimately mitigating the overactivation of autophagy. Through these interconnected negative feedback inhibition pathways, ethylene signaling and autophagy counterbalance each other when either process becomes dominant. This intricate interplay defends the plants against runaway activity from either a critical stress response or an excessive recovery program, ensuring their survival. The thickness of the lines represents the strength of inhibition, with thicker lines indicating stronger inhibition.

The CTR1-mediated phosphorylation of EIN2 regulates the proteolytic cleavage of EIN2, transducing ethylene signals from the ER to the nucleus12, 13, 49. Given the hypersensitive and insensitive autophagic responses of ctr1-2 and ein2-5 mutants, respectively, to carbon starvation, activated ethylene signaling through proteolytic cleavage of EIN2 regulated by two phosphorylation sites, S645 and S924, appears essential for suppressing autophagy. However, unlike EIN2S924A plants, which showed hypersensitive autophagy as expected, the S645A mutation did not confer a hypersensitive autophagic response to carbon starvation. This result somewhat echoes the substantially weaker ethylene responses of EIN2S645A plants compared to EIN2S924A plants in inhibiting leaf cell expansion of light-grown and etiolated seedlings12. The differential autophagy impacts of S645A and S924A mutations likely stem from multifaceted regulatory mechanisms. The absence of S645A effects on autophagy raises questions about potential redundancy from the modification of additional residues obscuring S645A effects. Notably, the TOR kinase phosphorylates EIN2 at S657, blocking EIN2 nuclear translocation and ethylene signaling activation64. Given the close proximity of S645 to S657 and the interplay between autophagy and TOR signaling, TOR-mediated regulation of EIN2 may compensate for the S645A mutation. Specifically, TOR-induced phosphorylation of EIN2 at S657 could potentially mask the effects of the S645A mutation, inhibiting EIN2 nuclear translocation. This compensatory mechanism may account for the higher expression of transcripts encoding TOR components observed in ctr1-2 mutants (Fig. 3i), representing a negative feedback loop to inhibit ethylene signaling when the EIN2 function is highly activated. Moreover, S645 and S924 may transduce distinct downstream signals, with S924 directly influencing autophagy machinery and S645 affecting peripheral pathways. Finally, localization or stability changes from the mutations could alter EIN2 activity. Further elucidating site-specific signaling outcomes and dissecting this intricate phosphorylation-based control of EIN2 and autophagy will be essential to explain the divergence between S645A and S924A mutations.

CTR1 interacts with ATG8 despite lacking a canonical ATG8-interacting motif (AIM) (Fig. 1e and f). This suggests that non-canonical binding sites may facilitate CTR1-ATG8 interaction. Intriguingly, CTR1 contains a motif (EESYQLQLALALRLSSE) with high sequence homology to the ubiquitin-interacting motif (UIM), a novel class of alternative ATG8 interaction sites that bind the UIM-docking site (UDS) present in ATG8 proteins39. The identification of the conserved UIM- UDS interface expands the range of selectively targeted autophagy cargoes that can be recruited independently of AIM sites39. The putative UIM-like sequence in CTR1 could facilitate its previously unknown interaction with ATG8 through this alternative UIM-UDS binding mode, providing a mechanism for the AIM-independent autophagic turnover of CTR1. Further biochemical and genetic characterization are needed to confirm whether this putative UIM motif directly mediates CTR1-ATG8a binding via the predicted UDS interface.

The seemingly contradictory results between GFP-ATG8a cleavage assays and the hypersensitive carbon starvation response in the ctr1-2 mutant (Fig. 3) suggest inefficient autophagy that leads to inadequate recycling of cellular components in the ctr1-2. GFP cleavage assays revealed similar release kinetics of free GFP across WT, ctr1-2, and ein2-5 backgrounds, indicating that autophagosome degradation proceeds similarly once initiated in all genotypes. However, while ConA treatment stabilized GFP-ATG8a in WT seedlings, it only led to a transient increase in GFP-ATG8a levels in ctr1-2 seedlings, followed by rapid degradation under starvation. In contrast, ein2-5 mutants maintained consistent GFP-ATG8a levels regardless of ConA treatment (Fig. 3g, 3h). These results suggest that CTR1 may stabilize ATG8a proteins, and ethylene signaling counteracts this process. However, endogenous ATG8a levels did not significantly differ among WT, CTR1ox, and ctr1-2 mutants (Supplementary Fig. 6), indicating that the role of CTR1 is specific to autophagosome stabilization rather than affecting ATG8a protein abundance. CTR1 colocalizes with multiple ATG proteins in addition to ATG8a (Supplementary Fig. 1), implying that it may participate in early autophagosome formation by potentially phosphorylating these proteins, similar to how mammalian mTORC1 kinase influences autophagy through phosphorylation of ATG proteins such as ATG1 and ATG1365, 66. These findings propose a model where CTR1 stabilizes autophagosomes, preventing their premature degradation, while ethylene signaling promotes autophagosome destabilization, thereby negatively regulating autophagy. This complex interplay between stabilization and destabilization processes likely determines the overall efficiency of autophagy under carbon starvation conditions.

The differential gene expression regulation of ATGs and TOR signaling components in ctr1-2 and ein2-5 mutants compared to WT suggests the presence of compensatory mechanisms. Under carbon starvation conditions, ctr1-2 mutants exhibited increased transcript levels of TOR components, including TOR and RAPTOR 1B, indicating enhanced TOR signaling activity. Simultaneously, ctr1-2 expressed higher levels of ATG genes compared to WT and ein2-5 mutants during starvation (Fig. 3i). This upregulation of ATG gene expression suggests a positive feedback mechanism to enhance autophagy-related processes, possibly in reaction to stress conditions. However, the hypersensitivity of ctr1-2 mutants to carbon starvation implies that increased ATG gene expression alone may not confer effective stress tolerance. We hypothesize that elevated TOR signaling in ctr1-2 likely inhibits autophagy initiation by phosphorylating and inactivating key autophagy regulators such as Atg13 and ULK1, despite increased expression of ATG genes 65, 67. Together with the potential role of ethylene signaling in destabilizing autophagosomes, the enhanced TOR signaling in ctr1-2 could partly explain its hypersensitivity to carbon starvation and decreased autophagosome numbers, despite showing similar levels of autophagic flux markers. Furthermore, the role of TOR signaling in coordinating nutrient responses and growth may disrupt cellular metabolic balance, potentially exacerbating the sensitivity of ctr1-2 mutants to nutrient deprivation. Further research is needed to understand the specific mechanisms through which ethylene and TOR signaling modulate autophagy and their implications for cellular adaptation to stress.

MATERIALS AND METHODS

Plant materials and growth conditions

Arabidopsis thaliana Col-0 was used as the WT reference throughout the study. All plants were grown in long-day conditions at 22 °C ± 2 °C or in vitro on half-strength Murashige and Skoog (MS) basal medium supplemented with 0.8 % plant agar (pH 5.7) with or without sucrose in a continuous light chamber at 22 °C ± 2 °C. All transgenic plants were homozygous and were identified by the segregation of specific antibiotic resistance, followed by the confirmation of protein expression via immunoblot analyses. Arabidopsis mutants and transgenic plants used in this study include ctr1-2, ein2-5, ein2-5atg7-2, etr1-1, etr1-6, etr1-7, ein3eil1, etr1-9ers1-3, etr1- 2ers2-3, etr1-6etr2-3ein4-4, etr2-3ers2-3ein4-4, atg2-1, atg5-1, atg7-2, atg9-3, atg11-1, atg18a-1, EIN2p:EIN2WT/ein2-5, EIN2p:EIN2S645A/ein2-5, EIN2p:EIN2S924A/ein2-5, EIN2p:EIN2AA/ein2-5, CTR1p:GFP-CTR1/ctr1-2, CTR1p:GFP-CTR1/atg7-2, CTR1p:GFP-CTR1;35S:mCherry- ATG8a/ctr1-2, 35S:GFP-ATG8a/WT, 35S:GFP-ATG8a/ctr1-2, 35S:GFP-ATG8a/ein2-5, 35S:GFP-CTR1/WT.

Plasmid construction

All molecular cloning was performed using Infusion cloning (Takara Bio USA) and Gateway (Invitrogen) strategies. The creation of the CTR1p-GFP-gCTR1 construct was described in Park et al. (2023). To create the 35S:GFP-CTR1 construct, coding sequences of full-length CTR1 were cloned into the pENTR entry vector (Invitrogen) and subsequently moved to the binary pSITE-2CA Gateway vector. To generate the 35S:Myc-ATG8a, 35S:GFP-ATG8a and 35S:mCherry-ATG8a constructs, coding sequences of full-length ATG8a were cloned into the pENTR vector and transferred to either binary pEarleygate 203, pSITE-2CA, or a modified pEarleygate 100 with mCherry, respectively. To generate the constructs for BiFC, the coding sequences of CTR1, ATG8a and CIP8 in the pENTR Gateway entry vector were transferred into the pCL112 or pCL113 BiFC binary vector.

Hypocotyl measurements

To measure the hypocotyl lengths of seedlings in response to ACC, Arabidopsis Col-0 and autophagy mutants were planted on an MS medium containing different concentrations of ACC. After cold treatment for 72 hours at 4 °C in darkness, the plates were placed vertically at 22 °C in either darkness or light for 3 or 5 days, respectively. The seedlings were photographed at the indicated times, and the hypocotyl length was measured using Image J software.

Confocal microscopy

Imaging of GFP, YFP, RFP, and mCherry fluorescence was performed using either a laser- scanning confocal microscope (Zeiss LSM880 upright) or a spinning-disk confocal microscope.

Samples of N. benthamiana leaves were directly mounted on glass slides in water, and epidermal cells were imaged by merging z-stack images acquired at 2 µm intervals. Arabidopsis samples were mounted similarly, and cells within the root elongation zone were imaged. For imaging Arabidopsis stable transgenic seedlings expressing GFP-ATG8a or co-expressing GFP-CTR1 and mCherry-ATG8a, seedlings were grown in liquid Gamborg (GM) medium under light for 5 days. For ConA treatment, seedlings were washed three times in liquid GM medium without sucrose and treated with 0.5 µM ConA dissolved in DMSO in the dark for 16 hours. Imaging was performed using a spinning-disk confocal system consisting of an Olympus IX-83 microscope equipped with a Yokogawa scanning unit (CSU-X1-A1; Hamamatsu Photonics) and an Andor iXon Ultra 897BV EMCCD camera (Andor Technology). The root elongation zone was imaged with a 60x (1.40 NA) or 100x (1.45 NA) oil objective (UPlanSApo; Olympus), and z-series were collected at 0.5 µm step sizes. For BiFC imaging, N. benthamiana leaves were co-infiltrated with Agrobacterium strains expressing nYFP-CTR1, bZIP-RFP, and either cYFP-ATG8a or cYFP-CIP8a. For colocalization analysis, leaves were co-infiltrated with strains expressing GFP-CTR1 and mCherry-ATG8s. After a 3-day incubation in constant light, leaf epidermal cells were imaged for YFP, GFP, RFP, and/or mCherry signals using a Zeiss LSM880 upright confocal microscope. For all multi-channel imaging experiments, cells expressing single fluorescent markers were imaged initially to ensure no fluorescence bleed- through occurred between channels.

Colocalization analysis

Spatial colocalization of GFP and mCherry fluorescent signals from labeled autophagosomes was quantified using Manders’ overlap coefficient analysis implemented in the JaCoP (Just Another Colocalization Plugin) plug-in for ImageJ 38. Regions of interest containing fluorescently labeled autophagosomes were cropped from the acquired images. The cropped images were processed with the automatic thresholding function of JaCoP to remove background signals.

Colocalization percentages were then determined by comparing the GFP (green) images with the mCherry (red) images, and vice versa. To account for random colocalization events, a rotation control was performed. One image from each pair was rotated 180 degrees, and the colocalization analysis was repeated on the rotated and original images to quantify the expected level of colocalization due to chance overlap.

Measurements of ethylene production

Surface-sterilized seeds were placed on 3 mL of half-strength Murashige and Skoog (MS) medium supplemented with 1% sucrose and 0.8% agar in 22 mL gas chromatography (GC) vials. After 3 day-stratification at 4°C in the dark, the vials were capped, and the seedlings were incubated in the dark for an additional three days. The accumulated ethylene was then measured using a Shimadzu GC2010 Plus capillary gas chromatography system equipped with an HS-20 headspace autosampler. To assess ethylene production in light-grown seedlings, stratified seeds were transferred to light conditions and grown for 4 days, followed by capping and incubation in the light for 2 days. Ethylene concentration was calculated as pL per seedling for 2-day incubation periods. All genotypes and treatments were evaluated using three biological replicates with the average and standard deviation (SD) were calculated.

Co-immunoprecipitation analysis

N. benthamiana leaves were infiltrated with Agrobacteria co-transformed with the plasmids of interest and incubated for 3 days in a growth chamber. Total proteins were extracted from the infiltrated leaves and homogenized in a co-immunoprecipitation buffer (25 Tris, pH 7.5, 150 mM NaCl, 5 mM EDTA, pH 8.0, 0.1% Tween-20, 1 mM DTT, 40 µM MG132, 1 mM PMSF, and 0.5x protease inhibitor cocktail) by a 10-min centrifugation at 5,000 RPM. Following the centrifugation, the supernatant was removed, and the proteins were homogenized and extracted from the pellets in a RIPA buffer (50 mM Tris, pH 7.5, 50 mM NaCl, 2 mM EDTA, pH 8.0, 0.1%

SDS, 5 mM DTT, 40 µM MG132, 1 mM PMSF, 1x protease inhibitor cocktail) by a 10-min centrifugation at 7,000 RPM. After quantifying the protein concentration with a Bradford assay, an equal amount of total protein extracts was incubated with ChromaTek GFP-Trap® Magnetic Agarose (Proteintech) according to the manufacturer’s instructions. The samples were eluted with boiled 2x sodium dodecyl sulfate (SDS) sample buffer, and subjected to immunoblotting analysis with anti-GFP and anti-Myc-POD antibodies.

GFP cleavage analysis

To analyze the GFP cleavage assay, seedlings were grown on MS medium without sucrose in the light for 7 days, followed by 2-to-9-day incubation in darkness for carbon starvation stress. The harvested seedlings were weighed and homogenized with a pestle in liquid nitrogen. The 2x SDS sample buffer was added to the seedlings in proportion to their weight, and the total protein extract from each sample was boiled for 3 minutes. Subsequently, the samples were centrifuged and resolved through SDS-polyacrylamide gel electrophoresis (SDS-PAGE), followed by immunoblotting with an anti-GFP or anti-Actin antibody. Signals were detected with the SuperSignalTM Femto Maximum Sensitivity Substrate or Pico PLUS Chemiluminescent Substrate.

Arabidopsis carbon starvation stress

For the fixed carbon starvation treatment, seedlings were grown on MS medium without sucrose in the light for 7 days and subsequently transferred to darkness for the indicated days. After the dark treatment, the seedlings were moved to recover for one week, during which time they were photographed. To measure the protein contents of the sample, whole seedlings were harvested for the indicated days, weighed, frozen, and ground in liquid nitrogen. The ground samples were resuspended in 2x SDS sample buffer in proportion to their weight. Subsequently, the samples were centrifuged and resolved through SDS-PAGE, followed by Coomassie brilliant blue (CBB) staining and immunoblotting with an anti-RbcL or anti-Actin antibody. Signals were detected with SuperSignalTM Pico PLUS Chemiluminescent Substrate.

Real-time quantitative PCR analysis

Total RNA was prepared using the RNeasy Plant Mini Kit (QIAGEN) and reverse transcribed using SuperScript II reverse transcriptase (Invitrogen) according to the manufacturer’s instructions. Quantitative RT-PCR was performed using PowerUPTM SYBRGreen Master Mix (Applied Biosystems). The primers used are listed in Supplementary Table 1. Three biological replicates were analyzed with three technical replicates per sample. The relative expression of candidate genes was normalized to Actin2.

Immunoblot analysis

Three-day-old dark-grown or seven-day-old light-grown seedlings of WT and autophagy mutants were grown in MS media without sucrose, followed by treatment with ACC for 2 hours. The harvested seedlings were blotted to remove excess liquid and weighed. Subsequently, the seedlings were frozen in liquid nitrogen and ground to a fine powder. The ground samples were resuspended in 2× SDS sample buffer in proportion to their weight. An equal amount of total protein extract from each sample was boiled for 3 minutes and centrifuged to extract the total protein. The total proteins were resolved by SDS-PAGE, followed by immunoblotting with anti- CTR1 or anti-Actin antibodies. The signals were detected using SuperSignal™ Pico PLUS Chemiluminescent Substrate.

AUTHOUR CONTRIBUTION

GMY and HLP conceived idea and designed research; HLP performed most of the experiments; WZ, YC, and CP performed autophagosome localization and quantification; GMY wrote original draft; GMY, HLP, and WZ edited the manuscript.

CONFLITS OF INTEREST

All authors declared no conflicts of interest

DATA AVAILABILITY

All data is available through the main text and supplementary information

AKOWLEDGEMENTS

This work was supported by the National Science Foundation (IOS-2245525 to GM.Y.). W.Z. was supported by the EMBRIO institute, contract #2120200, a National Science Foundation (NSF) Biology Integration Institute. We thank Dr. Taijoon Chung (Busan National University, South Korea) for providing the autophagy mutants and technical advice and Dr. Zhixiang Chen (Purdue University) for critical reading of the manuscript.

References

  1. 1.↵
    Yagyu, M. & Yoshimoto, K. New insights into plant autophagy: molecular mechanisms and roles in development and stress responses. J. Exp. Bot. 75, 1234–1251 (2023).
    OpenUrl
  2. 2.
    Sedaghatmehr, M. & Balazadeh, S. Autophagy: a key player in the recovery of plants from heat stress. J. Exp. Bot. 75, 2246–2255 (2024).
    OpenUrl
  3. 3.
    Sun, J. et al. An Arabidopsis Rab18 GTPase promotes autophagy by tethering ATG18a to the ER in response to nutrient starvation. Dev. Cell 58, 2947–2958.e2945 (2023).
    OpenUrl
  4. 4.↵
    Huang, L., Wen, X., Jin, L., Han, H. & Guo, H. HOOKLESS1 acetylates AUTOPHAGY-RELATED PROTEIN18a to promote autophagy during nutrient starvation in Arabidopsis. Plant Cell 36, 136–157 (2023).
    OpenUrl
  5. 5.↵
    Lee, H.Y., Park, H.L., Park, C., Chen, Y.C. & Yoon, G.M. Reciprocal antagonistic regulation of E3 ligases controls ACC synthase stability and responses to stress. Proceedings of the National Academy of Sciences of the United States of America 118 (2021).
  6. 6.↵
    Park, H.L. et al. Ethylene-triggered subcellular trafficking of CTR1 enhances the response to ethylene gas. Nat Commun 14, 365 (2023).
  7. 7.↵
    Abeles, F.B., Morgan, P.W. & Saltveit, M.E.J. Ethylene in plant biology. San Diego, CA: Academic Press. (1992).
  8. 8.↵
    Hua, J. et al. EIN4 and ERS2 are members of the putative ethylene receptor gene family in Arabidopsis. Plant Cell 10, 1321–1332 (1998).
    OpenUrlAbstract/FREE Full Text
  9. 9.↵
    Johnson, P.R. & Ecker, J.R. The ethylene gas signal transduction pathway: a molecular perspective. Annu. Rev. Genet. 32, 227–254 (1998).
    OpenUrlCrossRefPubMedWeb of Science
  10. 10.↵
    Alonso, J.M., Hirayama, T., Roman, G., Nourizadeh, S. & Ecker, J.R. EIN2, a bifunctional transducer of ethylene and stress responses in Arabidopsis. Science (New York, N.Y.) 284, 2148–2152 (1999).
    OpenUrl
  11. 11.
    Kieber, J.J., Rothenberg, M., Roman, G., Feldmann, K.A. & Ecker, J.R. CTR1, a negative regulator of the ethylene response pathway in Arabidopsis, encodes a member of the raf family of protein kinases. Cell 72, 427–441 (1993).
    OpenUrlCrossRefPubMedWeb of Science
  12. 12.↵
    Ju, C. et al. CTR1 phosphorylates the central regulator EIN2 to control ethylene hormone signaling from the ER membrane to the nucleus in Arabidopsis. Proceedings of the National Academy of Sciences of the United States of America 109, 19486–19491 (2012).
    OpenUrlAbstract/FREE Full Text
  13. 13.↵
    Qiao, H. et al. Processing and subcellular trafficking of ER-tethered EIN2 control response to ethylene gas. Science (New York, N.Y.) 338, 390–393 (2012).
    OpenUrl
  14. 14.↵
    Qiao, H., Chang, K.N., Yazaki, J. & Ecker, J.R. Interplay between ethylene, ETP1/ETP2 F-box proteins, and degradation of EIN2 triggers ethylene responses in Arabidopsis. Genes Dev. 23, 512–521 (2009).
    OpenUrlAbstract/FREE Full Text
  15. 15.↵
    Merchante, C. et al. Gene-specific translation regulation mediated by the hormone-signaling molecule EIN2. Cell 163, 684–697 (2015).
    OpenUrlCrossRefPubMed
  16. 16.↵
    Li, W. et al. EIN2-directed translational regulation of ethylene signaling in Arabidopsis. Cell 163, 670–683 (2015).
    OpenUrlCrossRefPubMed
  17. 17.↵
    Guo, H. & Ecker, J.R. Plant responses to ethylene gas are mediated by SCF(EBF1/EBF2)- dependent proteolysis of EIN3 transcription factor. Cell 115, 667–677 (2003).
    OpenUrlCrossRefPubMedWeb of Science
  18. 18.
    Binder, B.M. et al. The Arabidopsis EIN3 binding F-Box proteins EBF1 and EBF2 have distinct but overlapping roles in ethylene signaling. Plant Cell 19, 509–523 (2007).
    OpenUrlAbstract/FREE Full Text
  19. 19.↵
    Hao, D. et al. The RING E3 ligase SDIR1 destabilizes EBF1/EBF2 and modulates the ethylene response to ambient temperature fluctuations in Arabidopsis. Proceedings of the National Academy of Sciences of the United States of America 118 (2021).
  20. 20.↵
    Shakeel, S.N. et al. Ethylene Regulates Levels of Ethylene Receptor/CTR1 Signaling Complexes in Arabidopsis thaliana. J. Biol. Chem. 290, 12415–12424 (2015).
    OpenUrlAbstract/FREE Full Text
  21. 21.↵
    Chen, L. et al. Autophagy contributes to regulation of the hypoxia response during submergence in Arabidopsis thaliana. Autophagy 11, 2233–2246 (2015).
    OpenUrlCrossRefPubMed
  22. 22.↵
    Marshall, R.S. & Vierstra, R.D. Autophagy: The Master of Bulk and Selective Recycling. Annu. Rev. Plant Biol. 69, 173–208 (2018).
    OpenUrlCrossRefPubMed
  23. 23.
    Nolan, T.M. et al. Selective Autophagy of BES1 Mediated by DSK2 Balances Plant Growth and Survival. Dev. Cell 41, 33–46.e37 (2017).
    OpenUrlCrossRefPubMed
  24. 24.
    Avila-Ospina, L., Moison, M., Yoshimoto, K. & Masclaux-Daubresse, C. Autophagy, plant senescence, and nutrient recycling. J. Exp. Bot. 65, 3799–3811 (2014).
    OpenUrlCrossRefPubMedWeb of Science
  25. 25.
    Lee, H.Y., Chen, Z., Zhang, C. & Yoon, G.M. Editing of the OsACS locus alters phosphate deficiency-induced adaptive responses in rice seedlings. J. Exp. Bot. 70, 1927–1940 (2019).
    OpenUrl
  26. 26.
    Hartman, S. et al. Ethylene-mediated nitric oxide depletion pre-adapts plants to hypoxia stress. Nature Communications 10, 4020 (2019).
    OpenUrl
  27. 27.
    Ugalde, J.M. Every breath you don’t take, I’ll be helping you: Ethylene promotes hypoxia tolerance. Plant Physiol. 190, 1085–1087 (2022).
    OpenUrl
  28. 28.
    Iqbal, N. et al. Ethylene Role in Plant Growth, Development and Senescence: Interaction with Other Phytohormones. Frontiers in plant science 8, 475 (2017).
  29. 29.↵
    García, M.J., Angulo, M., Lucena, C., Pérez-Vicente, R. & Romera, F.J. To grow or not to grow under nutrient scarcity: Target of rapamycin-ethylene is the question. Frontiers in plant science 13, 968665 (2022).
  30. 30.↵
    Li, Q. et al. The SOS2-CTR1 module coordinates plant growth and salt tolerance in Arabidopsis. J. Exp. Bot. (2023).
  31. 31.
    Chien, Y.C. & Yoon, G.M. Subcellular dynamics of ethylene signaling drive plant plasticity to growth and stress: Spatiotemporal control of ethylene signaling in Arabidopsis. Bioessays, e2400043 (2024).
  32. 32.
    Yasumura, Y. et al. An Ancestral Role for CONSTITUTIVE TRIPLE RESPONSE1 Proteins in Both Ethylene and Abscisic Acid Signaling. Plant Physiol. 169, 283–298 (2015).
    OpenUrlAbstract/FREE Full Text
  33. 33.↵
    Lin, Z., Alexander, L., Hackett, R. & Grierson, D. LeCTR2, a CTR1-like protein kinase from tomato, plays a role in ethylene signalling, development and defence. Plant J. 54, 1083–1093 (2008).
    OpenUrlCrossRefPubMedWeb of Science
  34. 34.↵
    Bassham, D.C. Methods for analysis of autophagy in plants. Methods 75, 181–188 (2015).
    OpenUrlCrossRefPubMed
  35. 35.↵
    Kim, J. et al. Autophagy-related proteins are required for degradation of peroxisomes in Arabidopsis hypocotyls during seedling growth. Plant Cell 25, 4956–4966 (2013).
    OpenUrlAbstract/FREE Full Text
  36. 36.↵
    Nair, U., Thumm, M., Klionsky, D.J. & Krick, R. GFP-Atg8 protease protection as a tool to monitor autophagosome biogenesis. Autophagy 7, 1546–1550 (2011).
    OpenUrlCrossRefPubMedWeb of Science
  37. 37.↵
    Torggler, R., Papinski, D. & Kraft, C. Assays to Monitor Autophagy in Saccharomyces cerevisiae. Cells 6 (2017).
  38. 38.↵
    Bolte, S. & Cordelières, F.P. A guided tour into subcellular colocalization analysis in light microscopy. J. Microsc. 224, 213–232 (2006).
    OpenUrlCrossRefPubMedWeb of Science
  39. 39.↵
    Marshall, R.S., Hua, Z., Mali, S., McLoughlin, F. & Vierstra, R.D. ATG8-Binding UIM Proteins Define a New Class of Autophagy Adaptors and Receptors. Cell 177, 766–781.e724 (2019).
    OpenUrlCrossRefPubMed
  40. 40.↵
    Liu, W. et al. ATG8-Interacting Motif: Evolution and Function in Selective Autophagy of Targeting Biological Processes. Frontiers in plant science 12, 783881 (2021).
  41. 41.↵
    Harkey, A.F., Yoon, G.M., Seo, D.H., DeLong, A. & Muday, G.K. Light Modulates Ethylene Synthesis, Signaling, and Downstream Transcriptional Networks to Control Plant Development. Frontiers in plant science 10, 1094 (2019).
    OpenUrl
  42. 42.↵
    Smalle, J., Haegman, M., Kurepa, J., Van Montagu, M. & Straeten, D.V. Ethylene can stimulate Arabidopsis hypocotyl elongation in the light. Proceedings of the National Academy of Sciences of the United States of America 94, 2756–2761 (1997).
    OpenUrlAbstract/FREE Full Text
  43. 43.↵
    Kang, S., Shin, K.D., Kim, J.H. & Chung, T. Autophagy-related (ATG) 11, ATG9 and the phosphatidylinositol 3-kinase control ATG2-mediated formation of autophagosomes in Arabidopsis. Plant Cell Rep. 37, 653–664 (2018).
    OpenUrl
  44. 44.↵
    Azhar, B.J. et al. Basis for high-affinity ethylene binding by the ethylene receptor ETR1 of Arabidopsis. Proceedings of the National Academy of Sciences of the United States of America 120, e2215195120 (2023).
    OpenUrl
  45. 45.↵
    Lin, Y. et al. Exocyst-Positive Organelles and Autophagosomes Are Distinct Organelles in Plants. Plant Physiol. 169, 1917–1932 (2015).
    OpenUrlAbstract/FREE Full Text
  46. 46.↵
    Zhuang, X. et al. ATG9 regulates autophagosome progression from the endoplasmic reticulum in Arabidopsis. Proceedings of the National Academy of Sciences of the United States of America 114, E426–e435 (2017).
    OpenUrlAbstract/FREE Full Text
  47. 47.↵
    Neufeld, T.P. TOR-dependent control of autophagy: biting the hand that feeds. Curr. Opin. Cell Biol. 22, 157–168 (2010).
    OpenUrlCrossRefPubMed
  48. 48.↵
    Russell, R.C., Yuan, H.-X. & Guan, K.-L. Autophagy regulation by nutrient signaling. Cell Res. 24, 42–57 (2014).
    OpenUrlCrossRefPubMed
  49. 49.↵
    Wen, X. et al. Activation of ethylene signaling is mediated by nuclear translocation of the cleaved EIN2 carboxyl terminus. Cell Res. 22, 1613–1616 (2012).
    OpenUrlCrossRefPubMedWeb of Science
  50. 50.↵
    Gao, Z. et al. Localization of the Raf-like kinase CTR1 to the endoplasmic reticulum of Arabidopsis through participation in ethylene receptor signaling complexes. J. Biol. Chem. 278, 34725–34732 (2003).
    OpenUrlAbstract/FREE Full Text
  51. 51.↵
    Chang, C., Kwok, S.F., Bleecker, A.B. & Meyerowitz, E.M. Arabidopsis ethylene-response gene ETR1: similarity of product to two-component regulators. Science (New York, N.Y.) 262, 539–544 (1993).
    OpenUrl
  52. 52.
    Hua, J., Chang, C., Sun, Q. & Meyerowitz, E.M. Ethylene insensitivity conferred by Arabidopsis ERS gene. Science (New York, N.Y.) 269, 1712–1714 (1995).
    OpenUrl
  53. 53.↵
    Gamble, R.L., Coonfield, M.L. & Schaller, G.E. Histidine kinase activity of the ETR1 ethylene receptor from Arabidopsis. Proceedings of the National Academy of Sciences of the United States of America 95, 7825–7829 (1998).
    OpenUrlAbstract/FREE Full Text
  54. 54.
    Sakai, H. et al. ETR2 is an ETR1-like gene involved in ethylene signaling in Arabidopsis. Proceedings of the National Academy of Sciences of the United States of America 95, 5812–5817 (1998).
    OpenUrlAbstract/FREE Full Text
  55. 55.↵
    Moussatche, P. & Klee, H.J. Autophosphorylation activity of the Arabidopsis ethylene receptor multigene family. J. Biol. Chem. 279, 48734–48741 (2004).
    OpenUrlAbstract/FREE Full Text
  56. 56.↵
    Gamble, R.L., Qu, X. & Schaller, G.E. Mutational analysis of the ethylene receptor ETR1. Role of the histidine kinase domain in dominant ethylene insensitivity. Plant Physiol. 128, 1428–1438 (2002).
    OpenUrlAbstract/FREE Full Text
  57. 57.↵
    Chen, T. et al. Effects of tobacco ethylene receptor mutations on receptor kinase activity, plant growth and stress responses. Plant Cell Physiol. 50, 1636–1650 (2009).
    OpenUrlCrossRefPubMedWeb of Science
  58. 58.↵
    Binder, B.M., O’Malley, R.C., Wang, W., Zutz, T.C. & Bleecker, A.B. Ethylene stimulates nutations that are dependent on the ETR1 receptor. Plant Physiol. 142, 1690–1700 (2006).
    OpenUrlAbstract/FREE Full Text
  59. 59.↵
    Hall, B.P. et al. Histidine kinase activity of the ethylene receptor ETR1 facilitates the ethylene response in Arabidopsis. Plant Physiol. 159, 682–695 (2012).
    OpenUrlAbstract/FREE Full Text
  60. 60.↵
    Kim, H. et al. Ethylene receptor ETHYLENE RECEPTOR1 domain requirements for ethylene responses in Arabidopsis seedlings. Plant Physiol. 156, 417–429 (2011).
    OpenUrlAbstract/FREE Full Text
  61. 61.↵
    Hua, J. & Meyerowitz, E.M. Ethylene Responses Are Negatively Regulated by a Receptor Gene Family in Arabidopsis thaliana. Cell 94, 261–271 (1998).
    OpenUrlCrossRefPubMedWeb of Science
  62. 62.↵
    Cancel, J.D. & Larsen, P.B. Loss-of-function mutations in the ethylene receptor ETR1 cause enhanced sensitivity and exaggerated response to ethylene in Arabidopsis. Plant Physiol. 129, 1557–1567 (2002).
    OpenUrlAbstract/FREE Full Text
  63. 63.↵
    Clark, K.L., Larsen, P.B., Wang, X. & Chang, C. Association of the Arabidopsis CTR1 Raf-like kinase with the ETR1 and ERS ethylene receptors. Proceedings of the National Academy of Sciences of the United States of America 95, 5401–5406 (1998).
    OpenUrlAbstract/FREE Full Text
  64. 64.↵
    Fu, L. et al. The TOR-EIN2 axis mediates nuclear signalling to modulate plant growth. Nature 591, 288–292 (2021).
    OpenUrl
  65. 65.↵
    Puente, C., Hendrickson, R.C. & Jiang, X. Nutrient-regulated Phosphorylation of ATG13 Inhibits Starvation-induced Autophagy. J. Biol. Chem. 291, 6026–6035 (2016).
    OpenUrlAbstract/FREE Full Text
  66. 66.↵
    Chan, E.Y. mTORC1 phosphorylates the ULK1-mAtg13-FIP200 autophagy regulatory complex. Science signaling 2, pe51 (2009).
  67. 67.↵
    Jung, C.H. et al. ULK-Atg13-FIP200 complexes mediate mTOR signaling to the autophagy machinery. Mol Biol Cell 20, 1992–2003 (2009).
    OpenUrlAbstract/FREE Full Text
Back to top
Posted August 01, 2024.
Download PDF

Supplementary Material

Email
Autophagy-mediated CTR1 turnover orchestrates the reciprocal interaction between autophagy and ethylene signaling
Hye Lin Park, Weiwei Zhang, Yuan-Chi Chien, Chanung Park, Gyeong Mee Yoon
bioRxiv 2024.07.31.606019; doi: https://doi.org/10.1101/2024.07.31.606019
Twitter logo Facebook logo LinkedIn logo Mendeley logo
Citation Tools

Subject Area