Arabidopsis thaliana seeds of WT (Col-0) or other indicated genotypes were sterilized with 33% (v/v) bleach and 0.1% (v/v) Triton X-100 (Sigma) for 20 min, followed by five washes of 5 min each with sterile water. Sterilized seeds were stored at 4°C in darkness for at least 2 days to allow stratification before plating on solid ½ MS medium (2.22 g/L Murashige-Skoog vitamin and salt mixture [Caisson Laboratory, MSP09], 1% [w/v] sucrose, 0.6% [w/v] Phytoblend agar [Caisson Laboratory], 2.4 mM 2-morphinolino-ethanesulfonic acid [MES, Sigma], pH 5.7). Seedlings were grown under long-day conditions (16 h light) at 22°C for 7 days. Plants for transient expression in leaf protoplasts were grown in soil in a humidity-controlled growth chamber with 50% humidity at 20–23°C under long-day conditions for 4–6 weeks. T-DNA insertion mutants used in this study are: raptor1a (SALK_043920c), raptor1b (SALK_078159) (Anderson et al., 2005), S7817 (SALK_147817), G166 (GABI_166C06), G548 (GABI_548G07) (Deprost et al., 2007), and atg7 (GABI_655B06) (Chung et al., 2010). Transgenic plants used in this study are: GFP-ATG8e (Xiong et al., 2007b), TOR-OE1 and TOR-OE2 (Ren et al., 2011).

For sucrose and nitrogen starvation, 7-day-old seedlings grown on solid ½ MS medium were transferred to solid ½ MS medium lacking sucrose or nitrogen for an additional 3 days (Doelling et al., 2002). Sucrose starvation plates were kept in the dark after transfer. For salt and mannitol treatment, 7-day-old seedlings grown on solid ½ MS medium were transferred to liquid ½ MS medium with 0.16 M NaCl or 0.35 M mannitol for 6–8 h. For oxidative and ER stress, 7-day-old seedlings grown on solid ½ MS medium were transferred to liquid ½ MS medium with 5 mM H₂O₂ (Sigma) for 2–3 h, or with 2 mM dithiothreitol (DTT, Fisher) or 5 μg/mL tunicamycin (Sigma) for 6–8 h. For AZD8055 treatment, 7-day-old seedlings grown on solid ½ MS medium were transferred to solid ½ MS medium with 2 μM AZD8055 (LC Laboratories) for 1 day, or liquid ½ MS medium with 1 μM AZD8055 for 2–3 h.

For auxin treatment, 7-day-old seedlings were transferred to solid ½ MS medium supplemented with 20 nM 1-naphthaleneacetic acid (NAA, Sigma-Aldrich, N0640) with or without starvation for an additional 3 days, or in liquid ½ MS medium with 20 nM NAA for 6–8 h with or without stress treatments as described above. For BTH treatment, 7-day-old seedlings were transferred to liquid ½ MS medium supplemented with 100 μM acibenzolar-S-methyl (BTH, Sigma-Aldrich, 32820) with or without 20 nM NAA for 8 h.

For concanamycin A treatment, 7-day-old GFP-ATG8e seedlings were transferred to liquid ½ MS medium with DMSO or 1 μM concanamycin A (Sigma) with or without other stress or drug treatments for 6–8 h.

Arabidopsis seedling roots were stained with monodansylcadaverine (MDC) as described previously (Contento et al., 2005). MDC-stained seedlings were observed with a Zeiss Axio Imager.A2 upright microscope (Zeiss) equipped with Zeiss Axiocam BW/color digital cameras using a DAPI-specific filter at the Iowa State University Microscopy and Nanoimaging Facility. GFP-ATG8e transgenic seedlings were observed and photographed with the same microscope system with a GFP-specific filter. Cells within the root elongation zone were photographed and the number of autophagosomes in each image was counted and averaged from at least 10 images per sample. Confocal microscopy images of autophagosomes in root cells and leaf protoplasts were taken using a Leica SP5 × MP confocal/multiphoton microscope system (Leica) with a 63x/1.4 oil immersion objective at the Iowa State University Roy J. Carver High Resolution Microscopy Facility (Pu and Bassham, 2016).

GFP-ATG8e was transiently expressed in Arabidopsis leaf protoplasts as previously described (Sheen, 2002; Liu et al., 2012). 25–30 μg of GFP-ATG8e plasmid DNA was introduced into protoplasts using 40% (w/v) polyethylene glycol (PEG, Sigma-Aldrich). Protoplasts were washed and incubated in W5 solution (154 mM NaCl, 125 mM CaCl₂, 5 mM KCl, 2 mM MES, pH 5.7). For starvation treatment, protoplasts were incubated in W5 solution without sucrose or with 0.5% (w/v) sucrose as control at room temperature in darkness for 2 days in total. For other stress treatments, protoplasts were incubated in W5 solution with treatments as described in the Stress and Auxin Treatment section. Protoplasts were observed by fluorescence microscopy (Nikon Eclipse E200) using a FITC filter, and protoplasts with more than three visible autophagosomes were counted as active for autophagy (Yang et al., 2016). A total of 100 protoplasts were observed per genotype for each condition, and the percentage of protoplasts with induced autophagy was calculated and averaged from three independent experimental replicates.

The RAPTOR1B cDNA sequence was divided into two fragments, and each fragment was amplified from Col-0 cDNA using CloneAmp HiFi PCR Premix (Takara). The 5′ fragment of RAPTOR1B was amplified with forward primer 5′-CACCGAGCTCGAATTCATGGCATTAGGAGACTTAATGGTGTCTC-3′ (inserted SacI restriction site underlined), and reverse primer 5′-GTCAAACCCAATATCAAGCAAGGTACCCA-3′, digested with SacI and KpnI (within the RAPTOR1B cDNA sequence), and ligated into the pPZP212 binary vector (Hajdukiewicz et al., 1994; Li et al., 2009), which has a 35S promoter sequence at the 5′ end and a MYC tag sequence at the 3′ end of the insert. The 3′ fragment was amplified with forward primer 5′-TGGGTACCTTGCTTGATATTGGGTTTGAC-3′ and reverse primer 5′-CACCGTCGACTCTTGCTTGCGAGTTGTCGTGGGTG-3′ (inserted SalI restriction site underlined), digested with KpnI and SalI, and ligated into the pPZP212 vector containing the 5′ fragment to complete the full sequence. The entire construct was confirmed by sequencing.

Sequence data from this article can be found in the Arabidopsis Genome Initiative under the following accession numbers: TOR, AT1G50030; RAPTOR1A, AT5G01770, RAPTOR1B, AT3G08850; ATG8e, AT2G45170; ATG7, AT5G45900.

We have shown previously that decreased TOR expression via RNA interference induces autophagy in Arabidopsis, suggesting that TOR is a negative regulator of autophagy in plants (Liu and Bassham, 2010). To confirm that autophagy is induced by inhibition of TOR kinase activity (Montane and Menand, 2013), we examined autophagy activity after application of the asTORis AZD8055 (Dong et al., 2015). WT and atg7 seedlings, a previously characterized knockout mutant that is unable to form autophagosomes (Doelling et al., 2002), were grown under standard conditions for 7 days, followed by 1 μM AZD8055 treatment in liquid ½ MS medium for 2–3 h. Roots of seedlings were stained with monodansylcadaverine (MDC), an acidotropic dye that can stain autophagosomes (Biederbick et al., 1995; Contento et al., 2005), and examined by fluorescence microscopy. Autophagosomes appear as rapidly moving fluorescent puncta, and the number of visible puncta in each image were counted for quantification. As expected, compared to the basal level of autophagy in the control, inhibition of TOR activity by AZD8055 led to a significant increase in the number of autophagosomes (Figure 1A), while no autophagosomes were detected in the atg7 mutant. This confirmed that TOR negatively regulates autophagy in Arabidopsis, and that the kinase activity of TOR is critical for this regulation.

Previous studies have shown that down-regulation of TOR or its binding partners RAPTOR and LST8 leads to defects in plant growth and development (Anderson et al., 2005; Deprost et al., 2007; Moreau et al., 2012; Montane and Menand, 2013), suggesting that RAPTOR and LST8 are critical for TOR-regulated plant growth. To test whether inhibition of TOR complex activity by disruption of RAPTOR also induces autophagy, WT, raptor1a and raptor1b knockout mutant seedlings (Anderson et al., 2005) were grown on ½ MS medium with sucrose for a week, and autophagy in root cells was examined by MDC staining followed by fluorescence microscopy (Figures 1B,C). Compared to the basal level of autophagy in WT seedlings, the number of autophagosomes in the raptor1a mutant appeared slightly higher, but this difference was not statistically significant, possibly due to the variability between seedlings. The raptor1b mutant had a significantly higher number of autophagosomes, suggesting that the raptor1b mutant has constitutive autophagy, and that RAPTOR1A and RAPTOR1B may not function equally in autophagy regulation.

To confirm that the raptor1b mutant has increased basal autophagy under standard conditions, the autophagosome marker GFP-ATG8e was expressed transiently in WT, raptor1a and raptor1b leaf protoplasts (Figure 1D). The GFP-ATG8e fusion protein has been used extensively as a specific marker of autophagosomes and autophagic bodies (Yoshimoto et al., 2004; Contento et al., 2005; Pu and Bassham, 2016), and active autophagy is defined as more than three visible autophagosomes in a protoplast (Yang et al., 2016). WT protoplasts maintain a basal level of autophagy with a low percentage with active autophagy. Consistent with the MDC staining results, the percentage of raptor1b protoplasts with active autophagy was significantly higher than that of WT protoplasts (Figure 1E). However, raptor1a also had a significantly higher percentage of active autophagy in leaf protoplasts, although significantly lower than raptor1b. RAPTOR1A may therefore be more important for autophagy regulation in leaves than in roots.

To confirm that the constitutive autophagy in the raptor1b mutant is specifically due to the mutation in RAPTOR1B, the RAPTOR1B cDNA was transiently expressed under a 35S promoter together with GFP-ATG8e in raptor1b knock out mutant leaf protoplasts. Autophagy was assessed as described above using fluorescence microscopy (Figure 1F). Note that the number of autophagosomes in the raptor1b mutant is variable between experiments, depending most likely on the growth conditions and age of the plants, room temperature, etc. The percentage of protoplasts with active autophagy in the raptor1b mutant expressing the RAPTOR1B cDNA was substantially lower than for the mutant protoplasts alone, and was not significantly different from WT. This indicates that the increased basal autophagy observed in the raptor1b mutant was suppressed by expression of the RAPTOR1B cDNA, confirming that the constitutive autophagy phenotype is indeed due to the disruption of RAPTOR1B.

Previous studies and our data have shown that the TOR complex negatively regulates autophagy in Arabidopsis (Liu and Bassham, 2010) (Figure 1), but the conditions under which TOR is important in plants are unknown. In other organisms TOR is well-described as regulating autophagy in response to nutrients (Dobrenel et al., 2016), with a decrease in TOR activity during nutrient deficiency leading to activation of autophagy. We therefore hypothesized that overexpression of TOR might prevent activation of autophagy by nutrient deficiency, and that autophagy induction by other stresses might be TOR-independent. To test this hypothesis, we obtained several previously characterized Arabidopsis lines with T-DNA insertions in the TOR upstream region (S7817, G166, and G548) and two transgenic lines with TOR expressed from a 35S promotor (TOR-OE1 and TOR-OE2). All lines have overexpression of TOR and enhanced growth (Deprost et al., 2007; Ren et al., 2011), with the exception of S7817, which has decreased TOR expression in leaves and overexpression of TOR in roots (Deprost et al., 2007). Seeds of WT and the five TOR overexpression lines were germinated and grown on solid ½ MS medium plus sucrose for 1 week, followed by transfer to solid ½ MS medium plus or minus nitrogen in the light, or minus sucrose in the dark for an additional 3 days. Autophagosomes were detected by MDC staining followed by fluorescence microscopy (Figures 2A,B). Representative images of one of the TOR overexpression lines are shown in Figure 2A. Quantification of autophagosomes indicated that both WT and the TOR overexpression lines had a low basal level of autophagy under control conditions. The average number of autophagosomes in WT seedlings after sucrose or nitrogen starvation was significantly higher than in control conditions, whereas the TOR overexpression lines had no significant activation of autophagy. This indicates that overexpression of TOR can repress autophagy induced by nutrient starvation, suggesting that repression of TOR activity is required for activation of autophagy in response to nutrient depletion.

While previous studies have shown that TOR is involved in nutrient sensing (Dobrenel et al., 2016), the extent to which TOR regulates stress responses other than nutrient deficiency is not known, although a link to osmotic stress resistance in Arabidopsis has been suggested (Mahfouz et al., 2006; Deprost et al., 2007). Autophagy is activated in Arabidopsis by salt and osmotic stresses (Liu et al., 2009). Therefore, we also tested whether overexpression of TOR affects autophagy induced by salt or osmotic stress (Figures 2A,C). WT and the TOR overexpression lines were germinated and grown on solid ½ MS medium for 1 week, and then transferred to liquid ½ MS medium containing 0.16 M NaCl or 0.35 M mannitol for 6–8 h. Autophagy in seedling roots was assayed by MDC staining followed by fluorescence microscopy (Figure 2C). Autophagy in salt or mannitol treated WT seedlings was significantly higher than the basal level of autophagy seen under control conditions. As for nutrient deficiency, autophagy was not induced in TOR overexpression lines under salt or osmotic stress, indicating that TOR can also repress autophagy induced by these stresses.

To confirm these results, we measured autophagy by transient expression of GFP-ATG8e in leaf protoplasts from WT and TOR overexpressing plants under sucrose starvation, salt and osmotic stresses (Figures 2D,E). As the protoplast incubation buffer contains nitrogen, it was not possible to test nitrogen deficiency using our standard protocol. After transformation with GFP-ATG8e constructs, protoplasts were incubated with or without sucrose for 2 days (Figure 2D), or plus or minus 0.16 M NaCl or 0.35 M mannitol for 1 day (Figure 2E), after which autophagy was observed using fluorescence microscopy. The percentage of protoplasts with active autophagy was calculated, with 100 protoplasts observed per genotype for each condition. WT and TOR overexpression lines had a low level of autophagy under control conditions, except for the S7817 line which had constitutive activation of autophagy. In this line, TOR expression is decreased in leaves, potentially explaining this observation (Deprost et al., 2007). While WT protoplasts had a significantly higher level of autophagy under sucrose starvation, salt and osmotic stresses, autophagy in the TOR overexpression lines, with the exception of S7817, remained at a low basal level indistinguishable from that in control conditions. We conclude that TOR is a regulator of autophagy in response to salt and osmotic stress in addition to nutrient deficiency.

Autophagy is also induced by oxidative stress and ER stress in plants (Xiong et al., 2007b; Liu et al., 2012; Yang et al., 2016). Oxidative stress is triggered when cells accumulate excessive reactive oxygen species (ROS), and oxidized proteins and lipids are degraded through autophagy (Xiong et al., 2007a,b). ER stress is generated when unfolded or misfolded proteins exceed the capacity of protein folding or degradation systems, causing accumulation of proteins in the ER (Howell, 2013). It can be triggered by heat stress, or experimentally by chemicals such as dithiothreitol (DTT) or tunicamycin (Howell, 2013). To determine whether TOR regulates autophagy upon oxidative or ER stress, 7-day-old WT and TOR overexpression lines were transferred to liquid ½ MS medium plus or minus 5 mM H₂O₂ for 2–3 h to cause oxidative stress, or plus 2 mM DTT or 5 μg/mL tunicamycin for 6–8 h to cause ER stress. Autophagy in seedling roots was detected by MDC staining followed by fluorescence microscopy (Figures 3A–C). Representative images of one of the TOR overexpression lines are shown in Figure 3A. WT and TOR overexpression lines had a low level of autophagy under control conditions, and WT seedlings had significantly higher autophagy induction after oxidative or ER stress treatment. Unlike nutrient, salt or osmotic stresses, TOR overexpression had no effect on autophagy induction, as overexpression lines remained able to activate autophagy under these stresses, suggesting that autophagy is activated via a pathway that does not require inhibition of TOR activity.

To confirm that autophagy remains induced in TOR overexpression lines under oxidative and ER stress, GFP-ATG8e was transiently expressed in leaf protoplasts of WT and TOR overexpression lines. Protoplasts were incubated with or without 5 mM H₂O₂, 2 mM DTT, or 5 μg/mL tunicamycin for 1 day, and observed using fluorescence microscopy (Figures 3D,E). WT protoplasts had a significantly higher level of autophagy after oxidative or ER stress treatment. In accordance with the MDC staining results, TOR overexpression lines also had a significantly higher percentage of protoplasts with active autophagy after oxidative or ER stress treatment, with no significant difference compared to WT under the same stress conditions. This demonstrates that overexpression of TOR is unable to repress autophagy induced by oxidative or ER stress, suggesting that oxidative- or ER stress-induced autophagy might be regulated through a TOR-independent pathway.

The effects of oxidative stress on autophagy can be difficult to interpret, as autophagy is also triggered by signaling ROS produced by NADPH oxidase (Liu et al., 2009). Salicylic acid (SA) has also been shown to enhance ROS signaling and induce autophagy in plants (Yoshimoto et al., 2009); we therefore tested whether SA-induced autophagy is dependent on TOR. Seven-day-old WT and TOR overexpression lines were transferred to liquid ½ MS medium with 100 μM benzo-(1,2,3)-thiadiazole-7-carbothioic acid S-methyl ester (BTH), an SA agonist, or 80% ethanol as control for 8 h. Autophagy in seedling roots was detected by MDC staining followed by fluorescence microscopy (Supplementary Figure 1A). WT and TOR overexpression lines had a low level of autophagy under control conditions, while both WT and TOR overexpression lines had significantly increased autophagy induction upon BTH treatment, suggesting that SA-induced autophagy, as for H₂O₂-induced autophagy, is not TOR dependent.

Target of rapamycin activity in Arabidopsis can be enhanced by exogenous addition of the auxin 1-naphthaleneacetic acid (NAA) (Schepetilnikov et al., 2013), indicating that auxin might regulate plant growth and development via the TOR signaling pathway. We hypothesized that auxin might repress stress-induced autophagy in plants through the TOR pathway. To confirm the existence of TOR-dependent and -independent pathways for activation of autophagy, NAA was added exogenously to GFP-ATG8e transgenic plants to activate TOR, and the effect on autophagy under different conditions was assessed (Figures 4A–D and Supplementary Figure 1B). For nutrient deficiency, 7-day-old GFP-ATG8e transgenic seedlings were transferred to solid ½ MS medium with or without sucrose or nitrogen and plus 20 nM NAA or DMSO for an additional 3 days. For salt, osmotic and ER stress, and BTH treatment, 7-day-old seedlings were transferred to liquid ½ MS medium plus 0.16 M NaCl, 0.35 M mannitol, 2 mM DTT or 5 μg/mL tunicamycin, or 100 μM BTH and plus 20 nM NAA or DMSO for 6–8 h. For oxidative stress, 7-day-old seedlings were transferred to liquid ½ MS medium plus 20 nM NAA or DMSO for 6–8 h, with 5 mM H₂O₂ added only during the last 2–3 h to avoid cell death. To more clearly observe GFP-ATG8e-labeled autophagic bodies in the vacuoles by confocal microscopy, 1 μM concanamycin A was added to block degradation of autophagic bodies prior to imaging of the vacuoles (Dröse et al., 1993; Liu and Bassham, 2010) (Figure 4A). In control conditions, root cells had few autophagic bodies within the vacuole, whereas all stresses tested led to accumulation of large numbers of autophagic bodies. In the presence of auxin, autophagic body accumulation was inhibited in nutrient deficiency, salt and osmotic stress, but accumulation was still observed in oxidative and ER stress and upon BTH treatment. These results also indicate that NAA reduces the number of autophagosomes observed by blocking autophagosome formation, rather than by accelerating autophagosome degradation. Autophagy was quantified by counting the number of autophagosomes under each condition, averaged from 10 images per genotype for each condition (Figures 4B–D and Supplementary Figure 1B). Compared to the basal level of autophagy under control conditions, autophagy was significantly higher after stress treatments. In the presence of NAA, autophagy was still significantly induced by oxidative and ER stress conditions and in the presence of BTH, but no significant difference compared to control conditions was observed under nutrient starvation, salt and osmotic stresses. This suggests that NAA represses autophagy induced by sucrose and nitrogen starvation, salt and osmotic stresses, but not oxidative stress or ER stress, consistent with the results from overexpression of TOR.

To further confirm that addition of auxin represses stress-induced autophagy through activation of TOR, we examined whether auxin can inhibit the constitutive autophagy seen upon disruption of the TOR signaling pathway by chemical inhibition or genetic mutation (Figures 4E–G). To inhibit TOR kinase activity, 7-day-old GFP-ATG8e seedlings were transferred to liquid ½ MS medium with or without 20 nM NAA for 6–8 h, with DMSO or 1 μM AZD8055 added during the last 2–3 h of treatment. GFP-labeled autophagic bodies in roots after concanamycin A treatment were examined using confocal microscopy (Figure 4A). AZD8055 as expected led to a high accumulation of autophagic bodies in the vacuole, and NAA had no effect on this accumulation, suggesting that NAA acts upstream of TOR in the autophagy pathway. The extent of autophagy was quantified by counting root autophagosomes, and AZD8055 caused accumulation of autophagosomes both in the presence and absence of NAA, with no significant difference in autophagy induction (Figure 4E).

As an alternative approach, the effect of NAA upon inhibition of TOR complex function via knockout of RAPTOR1B was tested. Seven-day-old WT and raptor1b seedlings were transferred to liquid ½ MS medium with or without 20 nM NAA for 6–8 h, followed by MDC staining and autophagy detection by fluorescence microscopy (Figures 4F,G). NAA had no significant effect on the constitutive autophagy seen in the raptor1b mutant. Taken together, these results suggest that auxin acts upstream of TOR in the regulation of autophagy.