Liquidity Is a Critical Determinant for Selective Autophagy of Protein Condensates

Article

Liquidity Is a Critical Determinant for Selective Autophagy of Protein Condensates Graphical Abstract

Authors Akinori Yamasaki, Jahangir Md. Alam, Daisuke Noshiro, ..., Kuninori Suzuki, Yoshinori Ohsumi, Nobuo N. Noda

Correspondence [email protected]

In Brief Selective autophagy contributes to cellular homeostasis through clearance of biomolecular condensates. Yamasaki et al. find that the liquidity of protein condensates is a critical determinant for its selective membrane sequestration and that selective autophagy of liquid droplets is mediated by a receptor with floatability.

Highlights d

Ape1 undergoes phase separation to form semi-liquid droplets

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Floatability of Atg19 mediates condensation of Atg19 on the surface of Ape1 droplets

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Atg19 and Atg8-PE are sufficient for selective membrane sequestration of Ape1 droplets

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Liquidity of Ape1 droplets is important for selective autophagy

Yamasaki et al., 2020, Molecular Cell 77, 1–13 March 19, 2020 ª 2019 Elsevier Inc. https://doi.org/10.1016/j.molcel.2019.12.026

Please cite this article in press as: Yamasaki et al., Liquidity Is a Critical Determinant for Selective Autophagy of Protein Condensates, Molecular Cell (2019), https://doi.org/10.1016/j.molcel.2019.12.026

Molecular Cell

Article Liquidity Is a Critical Determinant for Selective Autophagy of Protein Condensates Akinori Yamasaki,1,2 Jahangir Md. Alam,1 Daisuke Noshiro,1 Eri Hirata,3 Yuko Fujioka,1 Kuninori Suzuki,3,4,5 Yoshinori Ohsumi,2 and Nobuo N. Noda1,6,* 1Institute

of Microbial Chemistry (BIKAKEN), Tokyo 141-0021, Japan Biology Center, Institute of Innovative Research, Tokyo Institute of Technology, Yokohama 226-8503, Japan 3Department of Integrated Biosciences, Graduate School of Frontier Sciences, University of Tokyo, Kashiwa 277-8562, Japan 4Life Science Data Research Center, Graduate School of Frontier Sciences, University of Tokyo, Kashiwa 277-8562, Japan 5Collaborative Research Institute for Innovative Microbiology, University of Tokyo, Tokyo 113-8657, Japan 6Lead Contact *Correspondence: [email protected] https://doi.org/10.1016/j.molcel.2019.12.026 2Cell

SUMMARY

Clearance of biomolecular condensates by selective autophagy is thought to play a crucial role in cellular homeostasis. However, the mechanism underlying selective autophagy of condensates and whether liquidity determines a condensate’s susceptibility to degradation by autophagy remain unknown. Here, we show that the selective autophagic cargo aminopeptidase I (Ape1) undergoes phase separation to form semi-liquid droplets. The Ape1-specific receptor protein Atg19 localizes to the surface of Ape1 droplets both in vitro and in vivo, with the ‘‘floatability’’ of Atg19 preventing its penetration into droplets. In vitro reconstitution experiments reveal that Atg19 and lipidated Atg8 are necessary and sufficient for selective sequestration of Ape1 droplets by membranes. This sequestration is impaired by mutational solidification of Ape1 droplets or diminished ability of Atg19 to float. Taken together, we propose that cargo liquidity and the presence of sufficient amounts of autophagic receptor on cargo are crucial for selective autophagy of biomolecular condensates.

INTRODUCTION Phase separation of biomolecules is a widespread phenomenon in cells that mediates the generation of biomolecular condensates (also referred to as membraneless organelles). These condensates play a range of physiological roles, such as accelerating particular biochemical reactions and isolating specific proteins (Alberti et al., 2019; Banani et al., 2017; Shin and Brangwynne, 2017). One distinguishing feature of biomolecular condensates formed by phase separation is their liquid-like nature, which is critical to their function. It is also known that liquid-like biomolecular condensates, also called as liquid droplets, can solidify into harmful aggregates or amyloids due to stress or muta-

tion, which can lead to neurodegenerative diseases and otherwise compromise health (Aguzzi and Altmeyer, 2016; Taylor et al., 2016). Many cellular components can be degraded by autophagy, which mediates not only nonselective bulk degradation of cytoplasmic components but also selective degradation of targeted biomolecules and organelles (Mizushima and Komatsu, 2011). Selective autophagy facilitates the clearance of biomolecular condensates, thereby contributing to cellular homeostasis and potentially preventing aggregation-induced disease states (Deng et al., 2017). Recent studies have shown that PGL granules and p62 bodies, which are formed through phase separation, are degraded by selective autophagy (Sun et al., 2018; Wang and Zhang, 2019; Zaffagnini et al., 2018; Zhang et al., 2018). However, it is unclear whether the fluidity of biomolecular condensates determines their susceptibility to selective autophagy and how sequestration of such cargos by autophagic membranes proceeds. The budding yeast vacuolar hydrolase Ape1 is a well-established target of selective autophagy, undergoing constitutive transport to the vacuole by a form of selective autophagy known as the cytoplasm-to-vacuole targeting (Cvt) pathway (Lynch-Day and Klionsky, 2010; Yamasaki and Noda, 2017). After synthesis, Ape1 immediately forms a dodecamer that further assembles into the Ape1 complex in a manner depending on its N-terminal propeptide (Morales Quinones et al., 2012; Yamasaki et al., 2016). Atg19 recognizes the propeptide of Ape1 using its coiled-coil (CC) domain, thereby functioning as the specific receptor for Ape1 (Leber et al., 2001; Scott et al., 2001). Atg19 also interacts with Atg8 and Atg11, key components of the selective autophagy mechanism, leading to the generation of an isolation membrane (IM) that expands along the surface of the Ape1 complex to exclusively sequester the complex into a specialized autophagosome termed the Cvt vesicle (Shintani et al., 2002; Suzuki et al., 2002). Importantly, Cvt vesicle formation requires the Ape1 complex, whereas bulk autophagosome formation does not require the presence of any specific cargo (Shintani and Klionsky, 2004). As a model system of selective autophagy, the Cvt pathway has been extensively studied by various methods. Together with work in mammalian cells assessing selective autophagy of Molecular Cell 77, 1–13, March 19, 2020 ª 2019 Elsevier Inc. 1

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the cargo protein p62, a general mechanism of selective autophagy has been established: a specific receptor/adaptor protein links targeted cargo to the IM via receptor-cargo and receptorAtg8 interactions (Noda et al., 2010; Rogov et al., 2014). This mechanism has been applied not only to selective autophagy of biomolecules (Bjørkøy et al., 2005; Noda et al., 2008; Scott et al., 2001) but also to various organelles, such as mitochondria (mitophagy), the endoplasmic reticulum (ER) (ER-phagy), and peroxisomes (pexophagy) (Anding and Baehrecke, 2017). However, it is still unclear how selective sequestration of cargos by the IM proceeds and how cargo properties, such as the liquidity of condensates, affects sequestration. In vitro reconstitution experiments using natural cargos are required to gain a complete understanding of the process. Here, we show that Ape1 undergoes phase separation to form spherical, semi-liquid condensates (Ape1 droplets) both in vitro and in vivo. In vitro reconstitution experiments indicate that Atg19 and lipidated Atg8 are necessary and sufficient for selective sequestration of Ape1 droplets. Moreover, mutational studies reveal that the liquidity of Ape1 droplets and the abundance of the autophagic receptor Atg19 on droplets are crucial to drive selective autophagy. These results provide critical insights into the mechanism of selective degradation of condensates and are likely applicable to a range of cargos, including both membraneless and membrane-bound organelles. RESULTS Ape1 Undergoes Phase Separation to Form Semi-liquid Droplets Our previous analyses revealed that while recombinant glutathione S-transferase (GST)-tagged Ape1 is soluble, removal of the GST tag using a protease results in the formation of spherical Ape1 assemblages (Yamasaki et al., 2016). Other groups have also reported similar spherical assemblages for intact Ape1 purified from yeast cells (Morales Quinones et al., 2012), leading us to hypothesize that the Ape1 complex forms a phase-separated liquid-like droplet. To investigate this possibility, we first addressed whether full-length Ape1 undergoes liquid-liquid phase separation in vitro. Consistent with our previous findings, purified

GST-Ape1-mCherry was soluble and not characterized by the formation of assemblages, whereas the digestion of the GST and mCherry tags by protease resulted in the formation of spherical droplets (Figures 1A and S1A). In order to visualize the protein but minimize the effect of tagging on phase separation, we used a mixture of protease-cleavable and non-cleavable Ape1-mCherry fusions at a 9:1 ratio throughout this study (Figure S1B). Droplet formation was dependent on Ape1 concentration and resistant to high salt concentrations (Figure S1C). Next, we monitored the number and size of Ape1 droplets by time-course analysis (Figure 1B). Droplet formation began immediately upon protease addition (the number of droplets; large-size droplet formation at 0 min is an artifact of the sample preparation method). The number of droplets reached a maximum at ~20 min and then decreased, whereas droplet size steadily increased throughout incubation, reflecting droplet coalescence (Figures 1B and 1C). After coalescence, droplet circularity approached ~1.0, reflecting the propensity of liquid-like droplets to minimize surface area by assuming a spherical topology. We also observed the coalescence of Ape1 complexes in vivo (Figure S1D), although the frequency was very low, because most cells contain only one Ape1 condensate. Moreover, 1,6-hexanediol, which is known to disrupt liquid-liquid phase separation (Kroschwald et al., 2017), dispersed not only Ape1 droplets in vitro but also the Ape1 complex in vivo (Figures 1D, 1E, and S1E). We also found that Ape1 complexes dispersed by 1,6-hexanediol treatment reformed after washing out of 1,6-hexanediol, again forming multiple Ape1 complexes that coalesced in cells (Figure 1F). In cells, liquid-like biomolecular condensates rapidly exchange their constituent molecules with the surrounding cytosol or nucleoplasm (Banani et al., 2017). To determine the exchange rate of Ape1 between Ape1 droplets and the surrounding solution, we performed fluorescence recovery after photobleaching (FRAP) experiments. Approximately 30% of fluorescence was recovered within 100 s for fresh Ape1 droplets (within 10 min of protease addition), whereas recovery was markedly decreased or nearly completely lost in older droplets (Figures 1G and S1F). We noticed that fluorescence recovery was first observed in the peripheral area of the droplet, whereas recovery in the central region was slow

Figure 1. Ape1 Undergoes Phase Separation to Form Semi-liquid Droplets (A) 10 mM Ape1 forms spherical droplets in vitro upon protease treatment. DIC, differential interference contrast. Bar, 10 mm. (B) The number and size of Ape1 droplets were monitored after protease treatment. Ratios of droplet number to maxima are presented. Data are presented as mean ± SD of three independent experiments. (C) Representative images of droplet coalescence. Numbers indicate the circularity of the droplet. Bar, 5 mm. (D) 10% 1,6-hexanediol treatment of Ape1 droplets. Bars, 20 mm. (E) Cells expressing Ape1WT-mCherry were treated with 5% 1,6-hexanediol for 20 min. Bars, 5 mm. (F) Time lapse imaging of Ape1 complex coalescence as monitored by mCherry fluorescence. Bars, 3 mm. Total mean volume of Ape1 complexes calculated from each image (assuming sphericity) are shown with ± SDs (12 and 60 frames for Before and After). p (N.S., not significant) > 0.05. (G and I) FRAP analyses of Ape1 droplets in vitro (G) and in vivo (I). y axis values of 100% and 0% are Ape1 droplet signal intensity before and immediately after photobleaching, respectively. Monitored areas of fluorescence intensity are shown as broken circles. (G) Mean signal intensities ± SD (n = 4) at each time point are shown for variously aged (min) Ape1 droplets, as indicated at the right of each panel. Insets are representative images following 9-min protease treatment. Bars, 1 mm. (I) Mean signal intensities ± SD (n = 5) of Ape1 complexes are shown for each time point. Insets are representative images. Bars, 1 mm. (H) Time-lapse imaging of Ape1 droplets after photobleaching. Line profiling was performed over the range of broken lines. Bars, 5 mm. (J) FRAP analysis of a giant Ape1 complex. Line profiles were performed over the range of broken lines. Bars, 1 mm. (K) Ape1 droplet formation when using the PURE system. Bar, 20 mm. (L) Quantification of soluble and total amounts of Ape1. The graph shows the mean concentration with ± SDs (n = 3). See also Figure S1 and Video S1.

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Figure 2. A P22L Mutation Solidifies Semiliquid Ape1 Droplets (A) Droplet formation assay. 10 mM Ape1WT or Ape1 mutants were incubated in the presence of protease. Bars, 5 mm. (B) 10% 1,6-hexanediol treatment of Ape1P22L condensates. Bars, 20 mm. (C) Cells expressing Ape1P22L-mCherry were treated with 5% 1,6-hexanediol for 20 min. Arrowheads indicate 1,6-hexanediol resistant Ape1 complexes. Bars, 5 mm. (D) The graph shows the mean proportion of cells containing Ape1 complexes resistant to 1,6hexanediol, ± SDs (n = 3). **p < 0.01. (E) FRAP analysis of Ape1P22L condensates in vitro. Representative images of condensates incubated in the presence of protease for 9 min are shown. Broken circles delineate analyzed regions. Bars, 2 mm. Data are quantified in Figure S2E. (F) In vitro comparison of Ape1WT (Figure 1G) and Ape1P22L (Figure S2E) by FRAP analysis. The graph shows the mean recovery of fluorescence with ± SDs (n = 4). ***p < 0.001, p (N.S.) > 0.05. See also Figure S2.

(Figure 1H). Intriguingly, while the coalescence of bleached and non-bleached droplets resulted in the formation of a spherical droplet, non-bleached Ape1 molecules were not able to disperse throughout the bleached region of this droplet (Figure 1H; Video S1). These results suggest that the fluidity of the central area of the Ape1 droplets is reduced in comparison to the peripheral area, which may explain the partial recovery of the FRAP signal. We also performed FRAP experiments on the Ape1 complex in vivo, which showed ~20% fluorescence recovery within 100 s (Figure 1I) and replicated the low fluidity of the central region of the Ape1 complex (Figure 1J), both of which are consistent with the in vitro data. In order to determine whether nascent Ape1 translated from ribosomes also forms liquid droplets in vitro, we established a cell-free translation system to express tag-free Ape1 using the protein synthesis using recombinant elements (PURE) system (Shimizu et al., 2001, 2005). In this cell-free system, Ape1 was successfully expressed in a time-dependent manner and started to form spherical droplets as the concentration of Ape1 increased (Figure 1K). We quantified the concentration of soluble and total Ape1 and found that the concentration of soluble Ape1 remained consistently between 0.6 and 0.7 mM after phase separation started at 20 min, although the total concentration of Ape1 continued to increase up to 6 mM (Figure 1L). In yeast cells, several groups have reported the concentration of Ape1 under physiological conditions, which is in the range of 0.2–2 mM and a mean concentration of 0.75 mM, which is over the saturated concentration of cell-free-expressed Ape1 (Ho et al., 2018). In addition, a saturation concentration determined independently of a protein’s total concentration, which was observed for

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Ape1, is known to be an important feature of liquid-liquid phase separation (Alberti et al., 2019). Considering these findings, we conclude that Ape1, which for many years has been described as a complex, is in fact capable of phase separation into a semi-liquid droplet both in vitro and in vivo. A P22L Mutation Solidifies Semi-liquid Ape1 Droplets The propeptide of Ape1 has two important roles in the Cvt pathway. The first is to link Ape1 dodecamers through trimerization, resulting in the formation of the Ape1 complex (Yamasaki et al., 2016). The second is to bind Atg19, the Ape1 receptor protein (Scott et al., 2001). Mutations in the propeptide of Ape1 that inhibit the Cvt pathway include the L11S mutation, which impairs both the trimerization of propeptides (thereby inhibiting Ape1 complex formation) and the interaction of propeptide with Atg19 completely, and the P22L mutation, which enhances the trimerization of propeptide and reduces Ape1 binding affinity for Atg19 (Oda et al., 1996). We next studied the effect of these mutations on Ape1 droplet formation in vitro (Figures 2A and S2A). Mutant Ape1L11S protein is characterized by a dispersed fluorescence signal without the formation of any assemblages, even after protease treatment. This suggests that Ape1L11S exists exclusively as a dodecamer, as previously observed by X-ray crystallography (Yamasaki et al., 2016). On the other hand, Ape1P22L mutant protein was observed to form amorphous condensates rather than spherical droplets upon protease treatment. Similar results were obtained when the in vitro translation system was employed (Figures S2B and S2C). Ape1P22L condensates were more resistant to 1,6-hexanediol treatment than wild-type Ape1 droplets (Figure 2B), as observed for the Ape1 complex in vivo (Figures 2C, 2D, and S2D). Moreover, FRAP analyses indicated that Ape1P22L condensates show almost no fluorescence recovery, even when using fresh

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Figure 3. HS-AFM Reveals Dynamic Surface Motion on Ape1 Droplets and Its Impairment by P22L Mutation (A and B) AFM images of (A) mApe1 and (B) Ape1L11S. The height profile of the line between points x and y is shown to the right. Bars, 20 nm. (C) Crystal structure of Ape1L11S dodecamer (PDB: 5JHQ). Bar, 15 nm. (D and E) AFM and fluorescence images for (D) Ape1WT droplets and (E) Ape1P22L condensates. The AFM image was obtained from the circled area in the fluorescence image. The height profile of the line between x and y is shown. The area shown in box z was magnified and processed using a FFT bandpass filter, as shown in zraw and zfiltered, respectively. (F and G) Time-lapse AFM images of (F) Ape1WT droplets and (G) Ape1P22L condensates processed with a FFT bandpass filter. Numbers indicate corresponding local points existing between the subsequent frames. Local points that were not detected in subsequent frames are indicated by an asterisk. Bars, 60 nm. (H) Graph indicates the mean of each local point movement between subsequent frames of videos. ***p < 0.001. See also Videos S2 and S3.

condensates (Figures 2E, 2F, and S2E), and no fluorescence recovery was observed for the Ape1P22L complex in vivo (Figures S2F and S2G). These data indicate that the introduction of the P22L mutation results in the solidification of the Ape1 complex, possibly by enhancing the interactions between propeptides. HS-AFM Reveals Dynamic Surface Motion on Ape1 Droplets and Its Impairment by P22L Mutation To explore morphological characteristics of the surface of Ape1 droplets in detail, we employed high-speed atomic force microscopy (HS-AFM) (Ando, 2014). The Ape1L11S mutant and mature Ape1 lacking propeptides (mApe1) were observed as tetrahedral dots with a slightly sunken interior, which is consistent with the crystal structure of Ape1 (Figures 3A–3C). HS-AFM observation of Ape1 droplets was performed using the fluorescence intensity of Ape1-mCherry to identify droplets. Unprocessed HS-AFM im-

ages of Ape1WT droplets suggested a spherical morphology with a smooth surface (Figure 3D, top right). When AFM images were processed by fast Fourier transform (FFT) bandpass filter to enhance surface asperity, honeycomblike surface structures could be resolved on the Ape1 droplet, which may represent clustering of Ape1 dodecamers (Figure 3D, bottom right). HS-AFM videos showed that the honeycomb-like patterning of Ape1 droplet surfaces was dynamic and characterized by constant reorganization (Figure 3F; Video S2). On the other hand, Ape1P22L droplets had a much more granular, undulating appearance (Figure 3E), and while the honeycomb pattern was observed following processing, the surface appeared static and not dynamic (Figure 3G; Video S3). To assess fluidity quantitatively, the movement of points between subsequent frames in videos was determined. The average movement distance of peaks observed in Ape1 droplets was 9.07 ± 5.95 nm for Ape1WT, which was reduced to 2.23 ± 1.56 nm for Ape1P22L (Figure 3H). These data further support the notion that the surface of Ape1 droplets is fluid and that this fluidity can be impaired by the P22L mutation. Condensation of Atg19 on the Surface of Ape1 Droplets Is Mediated by Atg19 ‘‘Floatability’’ Atg19 has been reported to localize preferentially to the surface of the Ape1 complex in vivo (Morales Quinones et al., 2012; Yamasaki et al., 2016). As we have now shown that Ape1 is not

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Figure 4. Condensation of Atg19 on the Surface of Ape1 Droplets Is Mediated by Atg19 Floatability (A) Overview of the Atg19 domain and the truncation mutants used in this study. (B) In vitro imaging of mClover3-Atg19 with Ape1 droplets. 10 mM mClover3 or mClover3-Atg19 variants were incubated with 10 mM Ape1WT droplets. Bars, 5 mm. (C) Line profiles for broken lines shown in (B). (D) In vivo imaging of mGFP-Atg19WT or indicated mutants in the presence of giant Ape1 complexes. Bars, 5 mm. (E) Line profiles for broken lines shown in (D). (F) Summary of Atg19 localization with respect to giant Ape1 complexes in (D) (WT, n = 134; DN, n = 165; CC, n = 87). See also Figure S3.

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Figure 5. In Vitro Reconstitution of the Sequestration Step of the Cvt Pathway (A) Interaction of Atg8-LUVs with Ape1 droplets in the presence or absence of Atg19. Green indicates Atg8- LUVs labeled with NBD-PE. Bars, 5 mm. (B–E) Incubation of Atg8 conjugated-GUV and Ape1 condensates. Ape1 droplets (WT [B–D] and P22L [E]) and mClover3-Atg19 (WT [B, C, and E] and DN [D]) were incubated with GUVs (B, D, and E) or sGUVs (C) containing mKalama1-Atg8-PE conjugates. Incubation times are indicated. Sequential images of the same GUV are shown, except for images at right in (B) and (C), which were obtained from a different experiment. Bars, 5 mm (B, D, and E) or 2 mm (C). (F) Quantification of the results shown in (B) (n = 48), (D) (n = 89), and (E) (n = 62). (G) Representative results of the Ape1 maturation assay. Cells expressing Atg19 variants were treated with or without 1 mg/mL rapamycin for 90 min. (H) Quantitation of data presented in (G). The graph shows the proportion of mature Ape1 as a percentage of total Ape1 ± SD (n = 3). p (N.S.) > 0.05, 0.01 < *p < 0.05, **p < 0.01. See also Figure S4 and Videos S4, S5, S6, and S7.

simply a complex in vivo but is in fact able to phase separate into droplets, we set out to further characterize the mechanism of Atg19 recruitment to Ape1 droplets. To this end, we first studied the interaction between Atg19 and Ape1 droplets in vitro (Figures 4A–4C). mClover3-tagged Atg19, but not mClover3 alone or an mClover3-tagged Atg193A mutant (lacking Ape1-binding ability), bound preferentially to the surface of Ape1 droplets. Interestingly, Atg19CC, which comprises the minimum Ape1-binding region, was able to penetrate into Ape1 droplets. Tag-free Atg19WT and Atg19CC labeled chemically with a fluorescent dye also showed the same binding pattern as that of mClover3-tagged proteins, confirming that these results are not an artifact of tagging (Figure S3A). A C-terminal deletion mutant (DC; lacking

residues 192–415) showed surface-specific localization similar to Atg19WT, whereas an N-terminal deletion mutant (DN; lacking residues 1–152) penetrated into the Ape1 droplets in a similar manner to Atg19CC. We also performed in vitro translation of tag-free Ape1 in the presence of a physiological concentration (0.1 mM) of mClover-Atg19 (Ho et al., 2018), which revealed that Ape1 droplet formation was not affected by the pre-existence of Atg19 (Figure S3B). In addition, we found that Atg19 showed similar binding patterns; Atg19WT bound to the surface of Ape1 droplets, whereas Atg19CC penetrated into them (Figure S3C). The concentration of Atg19 did not affect the surface-specific binding to Ape1 droplets, and the thickness of the Atg19 layer was constant irrespective of Atg19 concentration (Figures S3D and S3E), suggesting that Atg19 layer thickness is structurally determined. In FRAP experiments, both Atg19WT and Atg19DN showed higher fluorescence recovery than Ape1, suggesting that Atg19 moves freely within the Ape1 droplet (Figures 1G, S3F, and S3G). It should be noted that the molecular size of Atg19DN is larger than that of Atg19DC, indicating that the ability to penetrate into the Ape1 droplets is not determined simply by size. Thus, the N-terminal region of Atg19 functions as a ‘‘float’’ that keeps Atg19 on the surface of Ape1 droplets by inhibiting penetration. In order to determine the intracellular localization of Atg19 mutants, we next examined cells overexpressing Ape1, which leads

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to the formation of giant Ape1 complexes (Suzuki et al., 2013) (Figures 4D–4F). Hereafter, we refer to the Ape1 complex in vivo as the Ape1 droplet, since we have demonstrated that it is not simply a complex in cells. Consistent with previous reports, Atg19WT localized to the surface of Ape1 droplets (observed in 64.6% droplets) (Morales Quinones et al., 2012; Yamasaki et al., 2016). Surface-specific localization was partially impaired in Atg19DN (30.3%) and could not be detected for Atg19CC. We therefore conclude that Atg19 floats within Ape1 droplets, a property most likely afforded by its N-terminal region, and that this floatability retains Atg19 at the surface of Ape1 droplets. In Vitro Reconstitution of the Sequestration Step of the Cvt Pathway We next attempted to reconstitute the sequestration of Ape1 droplets by autophagosomal precursor membranes (IMs). To this end, we adopted an in vitro approach using purified proteins and liposomes, as employed in previous reconstitution experiments using an artificial cargo (Sawa-Makarska et al., 2014). IMs contain abundant phosphatidylethanolamine-conjugated Atg8 (Atg8-PE). The various selective autophagy receptors, including Atg19, directly bind to Atg8-PE as a critical part of the selective autophagy mechanism. In this experiment, large unilamellar vesicles (LUVs) and giant unilamellar vesicles (GUVs) comprising a similar phospholipid composition to endomembranes in yeast (van Meer et al., 2008) were mixed with Atg8, Atg8 conjugating enzymes, and ATP. This results in the production of Atg8-PE containing LUVs (Atg8-LUV) and GUVs (Atg8-GUV), which we used as a simplified model of precursor membranes of the IM and expanded IMs, respectively. When Atg8-LUVs and Atg19 were introduced to Ape1 droplets, Atg8LUVs tethered to the surface of Ape1 droplets and covered their surface as discontinuous membranes (Figure 5A, Atg19+). In contrast, the absence of Atg19 abolished this tethering (Figure 5A, Atg19
), suggesting that Ape1 droplets are able to recruit IM precursor membranes through an interaction between Atg19 and Atg8. When a mixture of Ape1 droplets and Atg19 was introduced to Atg8-GUVs, Ape1 droplets tethered to the surface of Atg8-GUVs in an Atg19-dependent manner (Figures 5B and S4A; Video S4). After tethering, Ape1 droplets were gradually internalized by invaginating membranes of Atg8-GUVs at the contact region (Video S5). Tight droplet-IM contact was observed throughout this process, and complementary contact interfaces were maintained. Tethering was associated first with spherical Ape1 droplet flattening along the membrane surface to expand the contact region. Following this, invagination into the GUV lumen commenced, together with the contacting membranes of the GUV, while the Ape1 droplets resumed a spherical shape (Figure S4B). In some cases, Ape1 droplets were completely sequestered into the lumen of Atg8-GUVs after a long incubation period (Figures 5B and 5F). However, the GUV-derived inner vesicles surrounding droplets were still connected to the outer membrane of Atg8-GUV by a narrow tubule (Figure S4C). These observations suggest that Atg8-PE and Atg19 are sufficient for the selective sequestration of Ape1 droplets by membranes, although other factors are likely required for complete membrane closure.

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In yeast, the size ratio of Ape1 droplets to the Cvt vesicle is typically ~0.4–0.6 (as determined by electron microscopy) (Baba et al., 1997; Scott et al., 1997). In order to increase the physiological relevance of the reconstitution experiment, we next prepared smaller Atg8-GUVs that better approximate the natural size ratio and mixed these with Ape1 droplets and Atg19. Upon tethering, Ape1 droplets induced a drastic topological change in Atg8-GUVs, from spherical to cup-shaped, where contacts are formed with Ape1 droplet surfaces (Figure 5C). This topological transition is strikingly similar to the cup shape of IMs isolating Ape1 droplets in yeast (Baba et al., 1994; Suzuki et al., 2013). In some cases, Ape1 droplets were sequestered into the lumen of Atg8-GUVs within 5 h (Figure 5C, 300 min). Sequestration of Ape1 droplets in vitro using exogenous membranes can therefore be faithfully reconstituted, so long as the size ratio of membrane vesicles to Ape1 droplets is physiologically relevant. The observed Ape1 droplet sequestration was much slower in comparison with that in vivo, where the half-time of Ape1 maturation is ~40 min (Klionsky et al., 1992), which can be attributed to the lack of IM expansion step and the requirement for dramatic spatial remodeling of preexisting membranes in our in vitro system. In summary, our in vitro reconstitution assay reveals that a cargo (Ape1 droplet), a receptor (Atg19), and Atg8-containing membranes alone are sufficient for the highly selective sequestration of a typical selective autophagy substrate. Atg19 Floatability Is Crucial for the Sequestration Step of the Cvt Pathway Using our in vitro reconstitution system, we next studied how the floatability of Atg19 affects cargo sequestration. Atg19DN penetrated into the Ape1 droplet; however, some fractions of Atg19DN were observed to concentrate with Atg8-PE at the contact site between GUVs and Ape1 droplets (Figures 5D and 5F). Since Atg19DN moves freely within droplets (Figures S3F and S3G), Atg8-GUVs may be able to recruit Atg19DN from the droplet interior to the contact site. Time-lapse microscopy confirmed that Ape1 droplets are tightly tethered to Atg8-GUVs by Atg19DN (Video S6). However, almost no shape changes were observed in Atg8-GUVs or Ape1 droplets, even after prolonged incubation (7 h) (Figures 5D and 5F). This result is consistent with a previous report that Atg19DN is defective in Ape1 maturation under nutrient-rich conditions in vivo (Shintani et al., 2002) and suggests that in order for sequestration of Ape1 droplets by IMs to proceed, Atg19 must be densely arranged on the droplet surface in advance, which is mediated by the floatability of Atg19 within droplets. While Ape1 is selectively transported to the vacuole by the Cvt pathway under growing conditions, delivery by bulk autophagy also occurs under starvation conditions or rapamycin treatment. It is known that Cvt vesicle formation is absolutely dependent on Ape1 droplets, whereas bulk autophagosome formation does not require any specific cargo. We therefore set out to determine the Ape1 maturation efficiency of Atg19DN-expressing cells under growing or rapamycin-treated conditions (Figures 5G and 5H). Under both conditions, Ape1 maturation was severely inhibited in atg19D and Atg193A cells but was observed in cells expressing Atg19WT, with 21.7% and 57.3% Ape1 maturing in

Please cite this article in press as: Yamasaki et al., Liquidity Is a Critical Determinant for Selective Autophagy of Protein Condensates, Molecular Cell (2019), https://doi.org/10.1016/j.molcel.2019.12.026

to occur in parallel with membrane shaping. We therefore monitored the localization of Atg5 and found that the recruitment of Atg5 to Ape1 droplets was severely impaired in atg19D cells expressing Atg19DN (Figure S4D). These observations suggest that the surface-specific localization of Atg19 is important not only in the formation of tight and extensive contacts between cargo and the IM that help to shape the IM but also in the promotion of Atg8-PE conjugation on Ape1-droplet-sequestering membranes. Both of these factors contribute to IM expansion on the surface of Ape1 droplets.

Figure 6. Cargo Liquidity Is a Critical Determinant of Cvt Pathway Progression (A) Fluorescence microscopy images of Ape1 droplets and IMs. Ape1 droplets and IMs were visualized by detecting fluorescence from GFP and 2 3 mCherry-Atg8, respectively. WT and ape1P22L cells were treated with rapamycin for 1 h before observation. Bar, 5 mm. (B–D) Area and circularity of Ape1 complexes and IM lengths were analyzed, respectively, for data presented in (A) (n > 300 for B and C, n > 40 for D). *p < 0.05, **p < 0.01 (Mann-Whitney U test). (E) Frequency of appearance of IMs in (A) were analyzed. At least 60 cells were counted for each determination (n > 6). *p < 0.05 (two-tailed Student’s t test). See also Figure S5.

Atg19WT cells under growing and rapamycin-treated conditions, respectively. Compared with Atg19WT, cells harboring Atg19DN showed defects in Ape1 maturation, with increased severity in growing cells (2.0-fold reduction) in comparison to rapamycintreated cells (1.4-fold reduction). These data suggest that the surface-specific localization of Atg19 at Ape1 droplets, which plays a critical role in membrane deformation, is particularly important for Ape1 transport through the Cvt pathway. Atg19 mediates not only the interaction between Atg8 and Ape1 but also the recruitment of Atg5, a component of the E3 enzyme required for Atg8-PE conjugation (Fracchiolla et al., 2016). Furthermore, during the sequestration of Ape1 droplets by Cvt vesicles in yeast cells, membrane expansion is thought

Cargo Liquidity Is a Critical Determinant of Cvt Pathway Progression Previous studies have reported a severe defect in Ape1P22L transport under nutrient-rich conditions, which was partially recovered by rapamycin treatment (Figures S5A–S5C) (Oda et al., 1996; Suzuki et al., 2002). To investigate why the P22L mutation had a relatively greater impact on the Cvt pathway, we studied the effect of the P22L mutation using our in vitro reconstitution system. In the presence of Atg19, Ape1P22L condensates contacting Atg8-GUVs moved with Atg8-GUVs when the latter were manipulated by micropipette, demonstrating association between the two (Video S7). However, deformation of Ape1P22L-associated Atg8-GUVs was never observed, even after long incubation times (Figures 5E and 5F). These observations suggest that the increased solidity of Ape1P22L condensates affects the ability of the condensate to induce shape changes in Atg8-GUV membranes, even though they could be tethered to the membrane. We then studied the effect of the P22L mutation on the Cvt pathway in more detail using giant droplets. As shown in Figure 6A, Ape1P22L formed large Ape1 droplets resembling those of Ape1WT. We confirmed that the expression level of Ape1P22L is similar to that of Ape1WT (Figure S5D). However, quantification of Ape1 droplets revealed that Ape1P22L droplets are significantly smaller than Ape1WT droplets, whereas the number of droplets per cell is similar (Figures 6B and S5E). Moreover, Ape1P22L droplets are less circular than Ape1WT droplets (Figure 6C), indicating that the liquidity of Ape1P22L droplets is reduced in comparison to Ape1WT droplets. Importantly, IMs observed in giant Ape1P22L cells (mCherry-Atg8) were significantly shorter and fewer in number than those in giant Ape1WT cells (Figures 6D and 6E). These in vivo observations reproduce our in vitro findings and provide further evidence that cargo liquidity is a key factor determining the ability of the Cvt pathway to sequester Ape1 droplets for delivery to the vacuole. The P22L mutation reduces the liquidity of Ape1 droplets while also partially impairing the interaction between Ape1 and Atg19 (Shintani et al., 2002), which we confirmed by in vitro pull-down assays using full-length proteins (Figures S5F–S5I). Therefore, the impairment of the Cvt pathway by this mutation could simply be due to the reduced affinity of Ape1 for Atg19. We first confirmed that Atg19 is recruited to Ape1P22L condensates by observation of mGFP-Atg19 puncta (Figures 7A and S6A). In order to ensure a similar affinity of Atg19 for Ape1WT and Ape1P22L, we prepared a strain where GFP-binding protein (GBP)-fused Ape1 is co-expressed with mGFP-Atg19, enabling a strong Ape1-Atg19 interaction (Figure 7B) (Araki et al., 2013). Due to

Molecular Cell 77, 1–13, March 19, 2020 9

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Figure 7. Cargo-Dependent Selective Autophagy Requires Tight Interactions between the IM and Phase-Separated Cargos (A) The average ratio of cells containing Atg19 puncta ± SD (n = 5, cells in each field >50). (B) Schematic diagram of the interactions between Ape1-GBP and mGFP/mClover3-Atg19. (C) mGFP-Atg19 puncta in Ape1-GBP expressing cells. Graph shows the average ratio of cells containing Atg19 puncta ± SD (n = 5, cells in each field >50). Bars, 5 mm. (D) In vitro observation of mClover3-Atg19- and Ape1WT-GBP-mediated condensates. Bar, 5 mm. (E) Schematic diagram of propeptide-mediated condensates and Atg19-mediated condensates. (F) Representative data for the Ape1-GBP maturation assay. Samples shown in top, middle, and bottom panels were immunoblotted with antiApe1, anti-Atg19, and anti-PGK1 antibodies, respectively. An asterisk indicates nonspecific bands. (G) Model of the tight interaction between Ape1 droplets with floating Atg19 and cargo-specific IM during strictly selective autophagy. Both the liquidity of Ape1WTdroplets and the floatability of Atg19 are important for selective sequestration by the IM. (H) Model of cargo incorporation with mild selectivity during bulk autophagy. In this case, the liquidity of Ape1WT droplets and the floatability of Atg19 are not important for selective sequestration by the IM. See also Figure S6.

the robust GFP-GBP interaction, mGFP-Atg19 binds strongly to Ape1-GBP, irrespective of the propeptide (wild type [WT], L11S, and P22L) (Figure S6B). Atg19-mGFP showed similar puncta for both Ape1WT-GBP- and Ape1P22L-GBP-expressing cells (Figure 7C), suggesting that Atg19 is recruited to Ape1P22L puncta in a similar manner to Ape1WT puncta. We noticed that Atg19-GFP also formed comparable puncta, even in Ape1L11SGBP-expressing cells, and also found that recombinant Ape1WT-GBP and Ape1L11S-GBP proteins formed amorphous condensates with mClover3-Atg19 (Figures 7D and S6C). These observations, together with a previous report that Atg19 forms a trimer (Bertipaglia et al., 2016), suggest that Ape1-GBP forms condensates with mGFP/mClover3-Atg19 via the GBP-mGFP/ mClover3 interaction rather than the inter-propeptide interaction; Ape1-GBP dodecamers are linked to each other via Atg19 trimerization in Ape1-GBP condensates, just as Ape1 dodeca-

10 Molecular Cell 77, 1–13, March 19, 2020

mers are linked to each other by propeptide trimerization in Ape1 condensates (Figure 7E). Ape1-GBP condensates do not appear to be liquid-like, which we confirmed by FRAP experiments on Atg19-GFP, as well as 1,6-hexanediol treatment of these condensates (Figures S6D–S6G). We then monitored maturation of Ape1-GBP and found that no Ape1-GBP maturation was observed for any of the propeptide types under growing conditions (Figure 7F, left 6 lanes). These observations suggest that Ape1GBP condensates are not delivered to the vacuole via the Cvt pathway, even when sufficient amounts of Atg19 are available. This is very likely due to the lack of liquidity of these condensates. In contrast, upon rapamycin treatment, maturation was observed for Ape1WT-GBP and Ape1L11S-GBP (Figure 7F, right 6 lanes), indicating that bulk autophagy can isolate Ape1GBP condensates with little liquidity. Meanwhile, maturation of Ape1P22L-GBP was not observed, even upon rapamycin treatment. This is probably due to the formation of exceedingly solid condensates that cannot be enclosed, even by bulk autophagosomes. These data lend further support to our conclusion that liquidity is important for selective autophagy of condensates, but not for bulk autophagy, as summarized in Figures 7G and 7H.

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DISCUSSION In this study, we find that the Ape1 complex in yeast is in fact a liquid droplet formed by phase separation and that Ape1 fluidity and the floatability of Atg19 determine the transport of Ape1 droplets to the vacuole by the Cvt pathway. We demonstrate that a clear distinction can be drawn between the Cvt pathway and bulk autophagy in terms of this cargo dependency; a template (Ape1 droplets) is required for the expansion of Cvt vesicle IMs, whereas bulk autophagy has no such requirement (Shintani et al., 2002; Suzuki et al., 2002, 2010). Using a new in vitro reconstitution approach, we conclude that cargo-receptor-Atg8 interactions are the main mechanism by which cargo sequestration occurs during selective autophagy and propose a model of selective autophagy of Ape1 (Figure 7G; note that the process of membrane expansion is precluded from the present discussion). During the Cvt pathway, an Ape1-specific IM expands along the surface of the Ape1 droplet with a high curvature, which is mediated by the tight and extensive contact between the IM and the droplet through Atg19. In this scenario, a sufficient density of exposed Atg19 at the cargo surface is critical, which relies on the floatability of Atg19 as it prevents its penetration into Ape1 droplets (Figure S6H). The liquidity of Ape1 droplets further strengthens the contact made with the IM by affording a degree of flexibility to droplets, allowing their flattening along the IM and thereby formation of a complementary contact interface (Figures 5B and S4B) (Suzuki et al., 2013). On the other hand, simple tethering of a cargo to the IM would be sufficient for sequestration of the cargo with mild selectivity during bulk autophagy, because the autophagosome is formed independently without using cargo as a template (Figure 7H). Based on the findings presented in this study, we propose a ‘‘strict’’ form of selective autophagy that utilizes cargo as a template for membrane formation and thereby excludes most of the cytoplasm from the lumen of autophagosomes. Strict selective autophagy in general requires a high density of receptors on the cargo surface in order to form tight and extensive contacts between the cargo and the IM. The ability of a receptor to float appears to greatly facilitate receptor concentration on the surface of liquid-like protein condensates, a strategy most likely adopted by other selective autophagy receptors for various liquid-like biomolecular condensates. This study also raises the prospect that liquid-like protein droplets would be more suitable as a cargo for strict selective autophagy than solid-like protein aggregates and amyloids due to the difference in their ability to deform IMs. In the case of PGL granules in C. elegans embryos, gel-like states with low mobility, rather than liquid-like states with high mobility, are associated with increased PGL granule degradation by selective autophagy (Zhang et al., 2018). Considering that Ape1 droplets are semi-liquid-like, it is likely that there is an optimal liquidity for biomolecular condensates to be an ideal cargo for selective autophagy; low liquidity renders the cargo resistant to deformation, thereby impairing formation of a complementary contact interface between cargo and the IM, whereas high liquidity results in an unstable cargo that cannot function as a template for IM expansion. The p62 body, a comparable selective cargo in

mammals, has also been reported to be liquid-like in its droplet state, albeit with low mobility (Sun et al., 2018; Zaffagnini et al., 2018). Future analysis on various states of the same cargo is important for fully understanding the optimal liquidity for selective autophagy. This principle may also be applicable to the selective autophagy of organelles, which are bound by a lipid bilayer membrane that can be thought of as a two-dimensional liquid that is both deformable but also structurally stable. The lipid bilayer may function as a template for IM expansion if the organelle surface contains sufficient receptors. In fact, receptors specific to degraded organelles are exposed at the organellar surface during organellophagy (Okamoto, 2014). These findings provide strong initial evidence that selective autophagy is able to make a substantial contribution to the degradation of biomolecular condensates characterized by semi-liquidity but is not able to degrade solid-like aggregates and amyloids by this mechanism. These findings have implications for the development of therapeutic compounds aiming to treat the progression of neurodegenerative diseases by using selective autophagy to eliminate pathological condensates. STAR+METHODS Detailed methods are provided in the online version of this paper and include the following: d d d d

KEY RESOURCES TABLE LEAD CONTACT AND MATERIALS AVAILABILITY EXPERIMENTAL MODEL AND SUBJECT DETAILS B Yeast strains METHOD DETAILS B Plasmids for recombinant protein expression in bacteria B Protein expression and purification B Plasmids for yeast experiments B C28H28N8O4 labeling B MBP pull down assay B Preparation and observation of Ape1 droplets B Microscope observation of Atg5 B In vitro translation of Ape1 using the PURE system B FRAP experiments B Immunoprecipitation B Western Blotting B Sample preparation for HS-AFM observation B HS-AFM imaging B HS-AFM image processing and analysis B Preparation of GUVs B Observation of GUVs B The single GUV method for observation of GUV-protein-droplet interactions B Micropipette manipulation for GUV-droplet tethering measurement B Interaction of Atg8-LUV with Ape1 droplets in the presence and absence of Atg19 B Visualization of IMs and Ape1 complexes by fluorescence microscopy

Molecular Cell 77, 1–13, March 19, 2020 11

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B

d

Morphological analysis of autophagy-related structures and Ape1 complexes B Statistical Analysis DATA AND CODE AVAILABILITY

Ando, T. (2014). High-speed AFM imaging. Curr. Opin. Struct. Biol. 28, 63–68. Araki, Y., Ku, W.-C., Akioka, M., May, A.I., Hayashi, Y., Arisaka, F., Ishihama, Y., and Ohsumi, Y. (2013). Atg38 is required for autophagy-specific phosphatidylinositol 3-kinase complex integrity. J. Cell Biol. 203, 299–313.

SUPPLEMENTAL INFORMATION

Baba, M., Takeshige, K., Baba, N., and Ohsumi, Y. (1994). Ultrastructural analysis of the autophagic process in yeast: detection of autophagosomes and their characterization. J. Cell Biol. 124, 903–913.

Supplemental Information can be found online at https://doi.org/10.1016/j. molcel.2019.12.026.

Baba, M., Osumi, M., Scott, S.V., Klionsky, D.J., and Ohsumi, Y. (1997). Two distinct pathways for targeting proteins from the cytoplasm to the vacuole/ lysosome. J. Cell Biol. 139, 1687–1695.

ACKNOWLEDGMENTS

Bajar, B.T., Wang, E.S., Lam, A.J., Kim, B.B., Jacobs, C.L., Howe, E.S., Davidson, M.W., Lin, M.Z., and Chu, J. (2016). Improving brightness and photostability of green and red fluorescent proteins for live cell imaging and FRET reporting. Sci. Rep. 6, 20889.

We thank Yuki Ishii for assistance with protein preparation, Shukun Hotta for assistance with yeast experiments, Yasuhiro Araki for providing plasmids containing the GBP sequence, Naonobu Fujita for assistance with microscope experiments, Tatsuya Kawaoka for advice on data analysis, Hidetoshi Noda and Naoya Kumagai for providing C4N4 compounds, and Alexander I. May for proofreading the manuscript. This work was supported in part by JSPS KAKENHI grants 25111004, 18H03989, 19H05707 (to N.N.N.), 17K18339 (to A.Y.), 19K16344 (to D.N.), 17H05894, 17K07319 (to Y.F.), 16H06375 (to Y.O.), 16H06280, 18H04853 (to K.S.), and 18J13429 (to E.H.); CREST, Japan Science and Technology Agency grant JPMJCR13M7 (to N.N.N.); the Takeda Science Foundation (N.N.N., Y.F., and A.Y.); the Mochida Memorial Foundation for Medical and Pharmaceutical Research (N.N.N.); the Tokyo Biochemical Research Foundation (N.N.N. and J.M.A.); and the Naito Foundation (N.N.N. and Y.F.). AUTHOR CONTRIBUTIONS A.Y. and N.N.N. conceived the project. A.Y. purified recombinant proteins and performed most of the in vitro experiments. Y.F. performed cell-free expression experiments. J.M.A. performed GUV experiments. D.N. performed HSAFM analyses. A.Y., E.H., K.S., and Y.O. performed yeast experiments. A.Y. and N.N.N. wrote the manuscript. N.N.N. supervised the work. DECLARATION OF INTERESTS The authors declare no competing interests. Received: July 18, 2019 Revised: November 22, 2019 Accepted: December 20, 2019 Published: January 28, 2020 REFERENCES Adachi, A., Koizumi, M., and Ohsumi, Y. (2017). Autophagy induction under carbon starvation conditions is negatively regulated by carbon catabolite repression. J. Biol. Chem. 292, 19905–19918. Aguzzi, A., and Altmeyer, M. (2016). Phase separation: linking cellular compartmentalization to disease. Trends Cell Biol. 26, 547–558. Ai, H.W., Shaner, N.C., Cheng, Z., Tsien, R.Y., and Campbell, R.E. (2007). Exploration of new chromophore structures leads to the identification of improved blue fluorescent proteins. Biochemistry 46, 5904–5910. Alam, J.M., and Yamazaki, M. (2011). Spontaneous insertion of lipopolysaccharide into lipid membranes from aqueous solution. Chem. Phys. Lipids 164, 166–174. Alam, J.M., Kobayashi, T., and Yamazaki, M. (2012). The single-giant unilamellar vesicle method reveals lysenin-induced pore formation in lipid membranes containing sphingomyelin. Biochemistry 51, 5160–5172. Alberti, S., Gladfelter, A., and Mittag, T. (2019). Considerations and challenges in studying liquid-liquid phase separation and biomolecular condensates. Cell 176, 419–434. Anding, A.L., and Baehrecke, E.H. (2017). Cleaning house: selective autophagy of organelles. Dev. Cell 41, 10–22.

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Noda, H., Asada, Y., Shibasaki, M., and Kumagai, N. (2019a). A fluorogenic C4N4 probe for azide-based labelling. Org. Biomol. Chem. 17, 1813–1816. Noda, H., Asada, Y., Maruyama, T., Takizawa, N., Noda, N.N., Shibasaki, M., and Kumagai, N. (2019b). A C4N4 diaminopyrimidine fluorophore. Chemistry 25, 4299–4304. Oda, M.N., Scott, S.V., Hefner-Gravink, A., Caffarelli, A.D., and Klionsky, D.J. (1996). Identification of a cytoplasm to vacuole targeting determinant in aminopeptidase I. J. Cell Biol. 132, 999–1010. Okamoto, K. (2014). Organellophagy: eliminating cellular building blocks via selective autophagy. J. Cell Biol. 205, 435–445. Parvez, F., Alam, J.M., Dohra, H., and Yamazaki, M. (2018). Elementary processes of antimicrobial peptide PGLa-induced pore formation in lipid bilayers. Biochim. Biophys. Acta Biomembr. 1860, 2262–2271. Rawicz, W., Olbrich, K.C., McIntosh, T., Needham, D., and Evans, E. (2000). Effect of chain length and unsaturation on elasticity of lipid bilayers. Biophys. J. 79, 328–339. Robinson, J.S., Klionsky, D.J., Banta, L.M., and Emr, S.D. (1988). Protein sorting in Saccharomyces cerevisiae: isolation of mutants defective in the delivery and processing of multiple vacuolar hydrolases. Mol. Cell. Biol. 8, 4936–4948. Rogov, V., Do¨tsch, V., Johansen, T., and Kirkin, V. (2014). Interactions between autophagy receptors and ubiquitin-like proteins form the molecular basis for selective autophagy. Mol. Cell 53, 167–178. Sawa-Makarska, J., Abert, C., Romanov, J., Zens, B., Ibiricu, I., and Martens, S. (2014). Cargo binding to Atg19 unmasks additional Atg8 binding sites to mediate membrane-cargo apposition during selective autophagy. Nat. Cell Biol. 16, 425–433. Schneider, C.A., Rasband, W.S., and Eliceiri, K.W. (2012). NIH Image to ImageJ: 25 years of image analysis. Nat. Methods 9, 671–675. Scott, S.V., Baba, M., Ohsumi, Y., and Klionsky, D.J. (1997). Aminopeptidase I is targeted to the vacuole by a nonclassical vesicular mechanism. J. Cell Biol. 138, 37–44. Scott, S.V., Guan, J., Hutchins, M.U., Kim, J., and Klionsky, D.J. (2001). Cvt19 is a receptor for the cytoplasm-to-vacuole targeting pathway. Mol. Cell 7, 1131–1141. Shimizu, Y., Inoue, A., Tomari, Y., Suzuki, T., Yokogawa, T., Nishikawa, K., and Ueda, T. (2001). Cell-free translation reconstituted with purified components. Nat. Biotechnol. 19, 751–755. Shimizu, Y., Kanamori, T., and Ueda, T. (2005). Protein synthesis by pure translation systems. Methods 36, 299–304.

Suzuki, K., Morimoto, M., Kondo, C., and Ohsumi, Y. (2011). Selective Autophagy Regulates Insertional Mutagenesis by the Ty1 Retrotransposon in Saccharomyces cerevisiae. Dev. Cell 21, 358–365. Suzuki, K., Akioka, M., Kondo-Kakuta, C., Yamamoto, H., and Ohsumi, Y. (2013). Fine mapping of autophagy-related proteins during autophagosome formation in Saccharomyces cerevisiae. J. Cell Sci. 126, 2534–2544. Tamba, Y., and Yamazaki, M. (2005). Single giant unilamellar vesicle method reveals effect of antimicrobial peptide magainin 2 on membrane permeability. Biochemistry 44, 15823–15833. Tamba, Y., Terashima, H., and Yamazaki, M. (2011). A membrane filtering method for the purification of giant unilamellar vesicles. Chem. Phys. Lipids 164, 351–358. Taylor, J.P., Brown, R.H., Jr., and Cleveland, D.W. (2016). Decoding ALS: from genes to mechanism. Nature 539, 197–206. van Meer, G., Voelker, D.R., and Feigenson, G.W. (2008). Membrane lipids: where they are and how they behave. Nat. Rev. Mol. Cell Biol. 9, 112–124. Wang, Z., and Zhang, H. (2019). Phase separation, transition, and autophagic degradation of proteins in development and pathogenesis. Trends Cell Biol. 29, 417–427. Watanabe, Y., Noda, N.N., Kumeta, H., Suzuki, K., Ohsumi, Y., and Inagaki, F. (2010). Selective transport of a-mannosidase by autophagic pathways: structural basis for cargo recognition by Atg19 and Atg34. J. Biol. Chem. 285, 30026–30033. Yamasaki, A., and Noda, N.N. (2017). Structural biology of the Cvt pathway. J. Mol. Biol. 429, 531–542. Yamasaki, A., Watanabe, Y., Adachi, W., Suzuki, K., Matoba, K., Kirisako, H., Kumeta, H., Nakatogawa, H., Ohsumi, Y., Inagaki, F., and Noda, N.N. (2016). Structural basis for receptor-mediated selective autophagy of aminopeptidase I aggregates. Cell Rep. 16, 19–27. Yamazaki, M. (2008). Chapter 5. The single Guv method to reveal elementary processes of leakage of internal contents from liposomes induced by antimicrobial substances. Adv. Planar Lipid Bilayers Liposomes 7, 121–142. Zaffagnini, G., Savova, A., Danieli, A., Romanov, J., Tremel, S., Ebner, M., Peterbauer, T., Sztacho, M., Trapannone, R., Tarafder, A.K., et al. (2018). p62 filaments capture and present ubiquitinated cargos for autophagy. EMBO J. 37, e98308. Zhang, G., Wang, Z., Du, Z., and Zhang, H. (2018). mTOR regulates phase separation of PGL granules to modulate their autophagic degradation. Cell 174, 1492–1506.e22.

Molecular Cell 77, 1–13, March 19, 2020 13

Please cite this article in press as: Yamasaki et al., Liquidity Is a Critical Determinant for Selective Autophagy of Protein Condensates, Molecular Cell (2019), https://doi.org/10.1016/j.molcel.2019.12.026

STAR+METHODS KEY RESOURCES TABLE

REAGENT or RESOURCE

SOURCE

IDENTIFIER

Antibodies Rabbit anti-Ape1 antibody

Suzuki et al., 2002

N/A

Rabbit anti-Atg19 antibody

Watanabe et al., 2010

N/A

Mouse anti-PGK1 antibody (clone 22C5D8)

Thermo Scientific

#459250, RRID: AB_2532235

Mouse anti-GFP antibody (clones 7.1 and 13.1)

Roche

#11814460001, RRID:AB_390913

Goat anti-mouse IgG-peroxidase labeled antibody

Sigma-Aldrich

#A9044

Goat anti-rabbit IgG-peroxidase labeled antibody

Sigma-Aldrich

#A0545

Anti-Rabbit IgG HRP Conjugate

Promega

#W4011

Novagen

#69450-3CN

Rapamycin

LC Laboratories

#R-5000

1,6-hexanediol

Nacalai tesque

#17913-72

Polyvinyl alcohol

Merck-Millipore

#8.14894.0101

One-step CBB

Biocraft

#CBB-1000

Bacterial and Virus Strains E. coli BL21 (DE3) Chemicals, Peptides, and Recombinant Proteins

C4N4

Noda et al., 2019a, 2019b

N/A

Protease inhibitor cocktail

Nacalai tesque

#04080-11

Phenylmethylsulfonyl fluoride

Sigma-Aldrich

#7626

ECL Prime Western Blotting Detection Reagent

GE Healthcare

#RPN2232

Immobilon Western chemiluminescent HRP substrate

Merck-Millipore

# WBKLS0500

Isopropyl b-D-thiogalactopyranoside

Nacalai tesque

#19742-94

1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine

Avanti Polar Lipids

#860320

1-palmitoyl-2-oleoyl-sn-glycero-3phosphoethanolamine

Avanti Polar Lipids

#850757

L-a-phosphatidylinositol

Avanti Polar Lipids

#840042

1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamineN-[methoxy(polyethylene glycol)-2000

Avanti Polar Lipids

#880160

Ampicillin

Nacalai tesque

#02739-32

Immersion oil Type-F For Microscopy

Olympus

#IMMOIL-F30CC

Digitonin

Nacalai tesque

#12333-51

New England Biolabs

#E2621

S. cerevisiae: AYY0094, Strain background: BY4741, Genotype: atg1D::kanMX4 ape1D::natNT2 his3::APE1mcherry-HIS3

Yamasaki et al., 2016

N/A

S. cerevisiae: AYY0070, Strain background: BY4741, Genotype: ape1D::natNT2 his3::APE1-mcherry-HIS3

Yamasaki et al., 2016

N/A

S. cerevisiae: AYY0073, Strain background: BY4741, Genotype: ape1D::natNT2 his3::ape1 P22L-mcherry-HIS3

Yamasaki et al., 2016

N/A

S. cerevisiae: AYY0141, Strain background: SEY6261, Genotype: atg19D::kanMX4 leu2::mRFP-APE1-LEU2 pRS424-CUP1P-APE1 pRS316

This study

N/A

Critical Commercial Assays NEBuilder HiFi DNA Assembly Experimental Models: Organisms/Strains

(Continued on next page)

e1 Molecular Cell 77, 1–13.e1–e9, March 19, 2020

Please cite this article in press as: Yamasaki et al., Liquidity Is a Critical Determinant for Selective Autophagy of Protein Condensates, Molecular Cell (2019), https://doi.org/10.1016/j.molcel.2019.12.026

Continued REAGENT or RESOURCE

SOURCE

IDENTIFIER

S. cerevisiae: AYY0142, Strain background: SEY6261, Genotype: atg19D::kanMX4 leu2::mRFP-APE1-LEU2 pRS424-CUP1P-APE1 pRS316-mGFP-ATG19

This study

N/A

S. cerevisiae: AYY0143, Strain background: SEY6261, Genotype: atg19D::kanMX4 leu2::mRFP-APE1-LEU2 pRS424-CUP1P-APE1 pRS316-mGFP-atg19 3A

This study

N/A

S. cerevisiae: AYY0144, Strain background: SEY6261, Genotype: atg19D::kanMX4 leu2::mRFP-APE1-LEU2 pRS424-CUP1P-APE1 pRS316-mGFP-atg19153-191

This study

N/A

S. cerevisiae: AYY0146, Strain background: SEY6261, Genotype: atg19D::kanMX4 leu2::mRFP-APE1-LEU2 pRS424-CUP1P-APE1 pRS316-mGFP-atg19D2–152

This study

N/A

S. cerevisiae: AYY0147, Strain background: BY4741, Genotype: atg19D::natNT2 pRS316

This study

N/A

S. cerevisiae: AYY0148, Strain background: BY4741, Genotype: atg19D::natNT2 pRS316-ATG19

This study

N/A

S. cerevisiae: AYY0149, Strain background: BY4741, Genotype: atg19D::natNT2 pRS316-atg19 3A

This study

N/A

S. cerevisiae: AYY0150, Strain background: BY4741, Genotype: atg19D::natNT2 pRS316-atg19D2–152

This study

N/A

S. cerevisiae: AYY0013, Strain background: BY4741, Genotype: ape1D::natNT2 his3::APE1-HIS3

Yamasaki et al., 2016

N/A

S. cerevisiae: AYY0022, Strain background: BY4741, Genotype: ape1D::natNT2 his3::ape1 P22L-HIS3

This study

N/A

S. cerevisiae: AYY0231, Strain background: SEY6261, Genotype: leu2::mRFP-Ape1-LEU2 atg19D::kan Atg5-mNeonGreen::natNT2 pRS316-Atg19 WT

This study

N/A

S. cerevisiae: AYY0270, Strain background: SEY6261, Genotype: leu2::mRFP-Ape1-LEU2 atg19D::kan Atg5-mNeonGreen::natNT2 pRS316-Atg19 153-415

This study

N/A

S. cerevisiae: AYY0237, Strain background: BY4741, Genotype: ape1D::natNT2 atg19D::hphNT1 his3::APE1 WT-GBP-HIS3 pRS316-Atg19 WT

This study

N/A

S. cerevisiae: AYY0238, Strain background: BY4741, Genotype: ape1D::natNT2 atg19D::hphNT1 his3::APE1 L11S-GBP-HIS3 pRS316-Atg19 WT

This study

N/A

S. cerevisiae: AYY0239, Strain background: BY4741, Genotype: ape1D::natNT2 atg19D::hphNT1 his3::APE1 P22L-GBP-HIS3 pRS316-Atg19 WT

This study

N/A

S. cerevisiae: AYY0240, Strain background: BY4741, Genotype: ape1D::natNT2 atg19D::hphNT1 his3::APE1 WT-GBP-HIS3 pRS316-mGFP-Atg19

This study

N/A

S. cerevisiae: AYY0241, Strain background: BY4741, Genotype: ape1D::natNT2 atg19D::hphNT1 his3::APE1 L11S-GBP-HIS3 pRS316-mGFP-Atg19

This study

N/A

S. cerevisiae: AYY0242, Strain background: BY4741, Genotype: ape1D::natNT2 atg19D::hphNT1 his3::APE1 P22L-GBP-HIS3 pRS316-mGFP-Atg19

This study

N/A

S. cerevisiae: AYY0243, Strain background: BY4741, Genotype: ape1D::natNT2 his3::APE1 WT-GBP-HIS3 pRS316-mGFP-Atg19

This study

N/A

S. cerevisiae: AYY0244, Strain background: BY4741, Genotype: ape1D::natNT2 his3::APE1 L11S-GBP-HIS3 pRS316-mGFP-Atg19

This study

N/A

(Continued on next page)

Molecular Cell 77, 1–13.e1–e9, March 19, 2020 e2

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Continued REAGENT or RESOURCE

SOURCE

IDENTIFIER

S. cerevisiae: AYY0245, Strain background: BY4741, Genotype: ape1D::natNT2 his3::APE1 P22L-GBP-HIS3 pRS316-mGFP-Atg19

This study

N/A

S. cerevisiae: AYY0267, Strain background: BY4741, Genotype: ape1D::natNT2 atg19D::hphNT1 his3::APE1 WT-HIS3 pRS316-mGFP-Atg19

This study

N/A

S. cerevisiae: AYY0268, Strain background: BY4741, Genotype: ape1D::natNT2 atg19D::hphNT1 his3::APE1 L11S-HIS3 pRS316-mGFP-Atg19

This study

N/A

S. cerevisiae: AYY0269, Strain background: BY4741, Genotype: ape1D::natNT2 atg19D::hphNT1 his3::APE1 P22L-HIS3 pRS316-mGFP-Atg19

This study

N/A

S. cerevisiae: AYY0271, Strain background: BY4741, Genotype: ape1D::natNT2 his3::GAL1p-APE1-mCherryHIS3 pRS426-GAL1p-Ape1

This study

N/A

S. cerevisiae: BY4741, Genotype: MATa his3D1 leu2D0 met15D0 ura3D0

Brachmann et al., 1998

N/A

S. cerevisiae: SEY6210, Genotype: MATa lys2 suc2 his3 leu2 trp1 ura3

Robinson et al., 1988

N/A

S. cerevisiae: GYS244, Strain background: SEY6210, Genotype: ape1D::APE1-GFP:kanMX

Suzuki et al., 2002

N/A

S. cerevisiae: GYS312, Strain background: SEY6210, Genotype: ape1D::APE1P22L-GFP:kanMX

Suzuki et al., 2002

N/A

S. cerevisiae: YMM192, Strain background: SEY6210, Genotype: mRFP-APE1::LEU2 atg19D::kanMX

Suzuki et al., 2011

N/A

S. cerevisiae: YOC5598, Strain background: GYS244, Genotype: ape1D::APE1-GFP:kanMX pYEX-BX-APE1 pRS314-mCherry-mCherry-atg8

This study

N/A

S. cerevisiae: YOC5599, Strain background: GYS312, Genotype: ape1D::APE1P22L-GFP:kanMX pYEX-BXAPE1P22L pRS314-mCherry-mCherry-atg8

This study

N/A

pGEX6P-1

GE healthcare

#28954648

pGEX6P-mCherry

This study

N/A

pGEX6P-mClover3

This study

N/A

pGEX6P-prApe1 WT-mCherry

This study

N/A

pGEX6P-prApe1 WT-CS-mCherry

This study

N/A

pGEX6P-prApe1 L11S-mCherry

This study

N/A

pGEX6P-prApe1 L11S-CS-mCherry

This study

N/A

pGEX6P-prApe1 P22L-mCherry

This study

N/A

Recombinant DNA

pGEX6P-prApe1 P22L-CS-mCherry

This study

N/A

pGEX6P-mApe1

Yamasaki et al., 2016

N/A

pGEX6P-prApe1 WT-GBP

This study

N/A

pGEX6P-prApe1 L11S-GBP

This study

N/A

pGEX6P-mKalama1-Atg8 K26P

This study

N/A

pACYC184-Atg5 Atg12

Hanada et al., 2007

N/A

pET11a-Atg7 Atg10

Hanada et al., 2007

N/A

pNCS-mClover3

Bajar et al., 2016

Addgene plasmid #74236

pBAD-mKalama1

Ai et al., 2007

Addgene plasmid #14892

pET15b

Merck-Millipore

# 69661

pET15b-MBP-Atg19 WT

Yamasaki et al., 2016

N/A (Continued on next page)

e3 Molecular Cell 77, 1–13.e1–e9, March 19, 2020

Please cite this article in press as: Yamasaki et al., Liquidity Is a Critical Determinant for Selective Autophagy of Protein Condensates, Molecular Cell (2019), https://doi.org/10.1016/j.molcel.2019.12.026

Continued REAGENT or RESOURCE

SOURCE

IDENTIFIER

pET15b-MBP-Atg19 3A

Yamasaki et al., 2016

N/A

pET15b-MBP-mClover3-Atg19 WT

This study

N/A

pET15b-MBP-mClover3-Atg19 3A

This study

N/A

pET15b-MBP-mClover3-Atg19 153-191

This study

N/A

pET15b-MBP-mClover3-Atg19 1-191

This study

N/A

pET15b-MBP-mClover3-Atg19 153-415

This study

N/A

pYA562-1

Araki et al., 2013

N/A

pYM-N22

Janke et al., 2004

N/A

pRS316

Sikorski and Hieter, 1989

N/A N/A

pRS316-Atg19 WT

Yamasaki et al., 2016

pRS316-Atg19 153-415

This study

N/A

pRS316-mGFP-Atg19 WT

This study

N/A

pRS316-mGFP-Atg19 3A

This study

N/A

pRS316-mGFP-Atg19 153-191

This study

N/A

pRS316-mGFP-Atg19 153-415

This study

N/A

pRS424-CUP1P-APE1

Suzuki et al., 2013

N/A

pRS426-GAL1p-prApe1

This study

N/A

pYEX-BX[prApe1]

Suzuki et al., 2013

N/A

pYEX-BX[prApe1P22L]

This study

N/A

pRS314[2 3 mCherry-Atg8]

Kamada et al., 2010

N/A

Image Lab software

Bio-Rad

N/A

Software and Algorithms ImageJ version 1.52a and 1.48v

https://imagej.nih.gov/ij/

N/A

cellSens

Olympus

N/A

Adobe Photoshop CC

Adobe Systems

N/A

Adobe Illustrator CC

Adobe Systems

N/A

FV31S-SW

Olympus

N/A

R software, version 3.1.3

https://www.r-project.org

N/A

RandomForest library, version 4.6–10

https://cran.r-project.org/web/ packages/randomForest/ randomForest.pdf

N/A

MetaVue imaging software

Molecular Devices

N/A

GST accept resin

Nacalai tesque

# 09277-14

Amylose Resin High Flow

New England Biolabs

# E8022L

HisPur Ni-NTA Superflow Agarose

Thermo scientific

#25216

SupraBead-protein G

Recenttec

#R9-MF-PRG-3000

Zirconia silica beads 0.6 mm

Bio Medical Science

#ZS06-001

Superdex 200 26/60 prep grade

GE Healthcare

# 28989336

Superdex 75 increase 10/300 GL

GE Healthcare

#29148721

Bio-Gel P-6 Desalting Cartridge

Bio-Rad

# 7325312

Hi Trap Q HP column

GE Healthcare

# 17115401

Sp Sepharose Fast flow

GE Healthcare

# 17072901

Coverslip 24x32 mm2, 0.13-0.17 mm thick

Matsunami Glass Ind.

# C024321

Glass bottom dish

MatTek

# P35G-1.5-14-C/H

Other

Cover glass 24x24 0.12-0.17 mm thickness

Muto Pure Chemical

#24242

Uneven slide glass

Matsunami Glass Ind.

# TF0410

76 3 26 mm slide glass

Matsunami Glass Ind.

#S1225

18 3 18 mm micro cover glass

Matsunami Glass Ind.

#No. 1-S

Molecular Cell 77, 1–13.e1–e9, March 19, 2020 e4

Please cite this article in press as: Yamasaki et al., Liquidity Is a Critical Determinant for Selective Autophagy of Protein Condensates, Molecular Cell (2019), https://doi.org/10.1016/j.molcel.2019.12.026

LEAD CONTACT AND MATERIALS AVAILABILITY Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact Nobuo N. Noda ([email protected]). All reagents generated in this study are available from the Lead Contact without restriction. EXPERIMENTAL MODEL AND SUBJECT DETAILS Yeast strains Yeast strains used in this study are listed in the Key Resources Table. For yeast strain handling, standard methods were used, as described previously (Yamasaki et al., 2016). Briefly, yeast cells were cultured until OD ~3.0 (except for Figure 2C, where culture was to OD ~8.5) in YPD (1% Bacto yeast extract, 2% Bacto peptone, and 2% glucose), SDCA with Ade and Trp (0.17% yeast nitrogen base without amino acids and ammonium sulfate, 0.5% ammonium sulfate, 0.5% casamino acid, 0.002% adenine sulfate, 0.002% tryptophan, and 2% glucose) for pRS316 carrying strains, or SDCA with Ade for pR316 and pRS424 carrying strains. To drive gene expression under the CUP1 promotor, 0.25 mM CuSO4 was added to media. To induce expression from the GAL1 promoter, log phase cells were washed twice and incubated in media containing 2% galactose for 6 h. Rapamycin (LC Laboratories) was added to media at 1 mg/ml for 90 min or 3 h. METHOD DETAILS Plasmids for recombinant protein expression in bacteria For construction of pGEX6P-Ape1-mCherry and Ape1-CS-mCherry variants, pGEX6P-Ape1 WT, L11S, and P22L were modified (Yamasaki et al., 2016). mCherry or CS (HRV 3C protease recognition site; amino-acid sequence LEVLFQGP)-mCherry was inserted into the downstream region of Ape1 without a linker. For mClover3-Atg19 expression, the pET15b-MBPmClover3 vector with CS and mClover3 insertions (amplified from pBAD-mClover3 (Addgene) at the downstream region of the MBP gene) was used. Atg19 WT, 3A, DC (residues 1-191), CC (residues 153-191), and DN (residues 153-415) were amplified from pGEX6P-Atg19WT (Yamasaki et al., 2016) by PCR and inserted into the downstream region of mClover3 without a linker. For mKalama1-Atg8K26P expression, the mKalama1 gene amplified from pBAD-mKalama1 (Addgene) was inserted into the upstream region of the ATG8 gene of pGEX6P-Atg8 K26P (Liu et al., 2018). For construction of pGEX6P-Ape1GBP, the GFP binding protein sequence, which was amplified from pYA562-1 (Araki et al., 2013), was inserted into the downstream region of Ape1 L514 in pGEX6P-Ape1. All gene insertions were performed by NEBuilder HiFi DNA Assembly (NEB). The expression vectors for Atg8 conjugation enzymes (Atg3, Atg7, Atg16, and Atg5-Atg12) were used as described previously (Hanada et al., 2007). Protein expression and purification All proteins used for in vitro experiments were expressed in BL21 (DE3). For the expression of Atg5-Atg12 conjugates, the transformant of BL21 (DE3) with pACYC 184-Atg5 Atg12 and pET11a-Atg7 Atg10 was used (Hanada et al., 2007). Bacteria were grown in 2x YT medium. Isopropyl b-D-thiogalactopyranoside was added to the culture when OD600 reached 0.8-1.0 and was further incubated overnight at 16 C. After centrifugation, the pellets were resuspended in PBS or MBP buffer (20 mM Tris pH 8.0, 150 mM sodium chloride), supplemented with 5 mM EDTA, 2 mM DTT, 1 mM PMSF, and 1x protease inhibitor (Nacalai tesque) for GST-fused proteins or MBP-fused protein, respectively, or in sodium phosphate buffer (50 mM sodium phosphate pH 8.0, 300 mM sodium chloride, 10 mM imidazole) for Atg8 conjugation enzymes. The samples were sonicated for 10 min and centrifuged at 18000 rpm for 40 min. The supernatants were recovered and applied to GST accept resin (Nacalai tesque), amylose resin (NEB), or Ni-NTA resin (Thermo scientific) for GST-, MBP-, or 6xHis-fused proteins, respectively. The resins were washed with each resuspension buffer 3 times and GST-, MBP-, or 6xHis-fused proteins were eluted with glutathione buffer (50 mM Tris pH 8.0, 10 mM glutathione), maltose buffer (10 mM Maltose, 20 mM Tris pH 8.0, and 200 mM sodium chloride), or imidazole buffer (50 mM sodium phosphate pH 8.0, 150 mM sodium chloride, 500 mM imidazole), respectively. The eluted samples were desalted by Bio-Gel P-6 Desalting Cartridge (Bio-rad) and digested with HRV 3C protease at 4 C overnight, except for Atg5-Atg12 conjugates, which were incubated with TEV protease at 4 C overnight. After removal of digested affinity tags by each affinity resin, the samples were purified by size-exclusion chromatography using Superdex 200 columns (GE healthcare) for GST-Ape1 and Atg19 or a Superdex 75 column (GE healthcare) for Atg8 with 20 mM HEPES pH 7.0 and 150 mM sodium chloride. For Atg3, Atg7, and Atg5-Atg12 proteins, the samples were purified by ion-exchange chromatography with Hi Trap Q HP column (GE healthcare) for Atg3 and Atg7 and Sp Sepharose Fast flow column (GE healthcare) for Atg5-Atg12 with 20 mM Tris pH 8.0 as buffer A and 20 mM Tris pH 8.0, 1 M sodium chloride as buffer B. Atg5-Atg12 was subsequently mixed with Atg16. The samples were further purified by size-exclusion chromatography using a Superdex 200 column with 20 mM HEPES pH 7.0 and 150 mM sodium chloride. Purified proteins were stored at
80 C until use.

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Plasmids for yeast experiments mGFP, which harbors a monomerizing A206K mutation, was inserted into the upstream region of the first methionine of Atg19 in pRS316-Atg19WT (Yamasaki et al., 2016). Atg19 variants and pYEX-BX[Ape1P22L] were generated by site-directed mutagenesis. To construct pRS426-GAL1p-Ape1, a GAL1 promoter amplified from pYM-N22 (Janke et al., 2004) and Ape1 gene were inserted into pRS426 using HindIII and BamHI. C28H28N8O4 labeling C28H28N8O4 (C4N4) labeling was performed as previously described (Noda et al., 2019b, 2019a). Briefly, protein was mixed with an equivalent amount of C4N4 for 30 min in the dark. The same volume of glutathione buffer was then added to inactivate unconjugated C4N4. The samples were purified by size-exclusion chromatography or dialyzed with 20 mM HEPES pH 7.0, 150 mM sodium chloride for buffer exchange. Purified proteins were stored at
80 C until use. MBP pull down assay 0.66 nM MBP-fused proteins were incubated with amylose resin for 1 h at 4 C. GST-Ape1 variants were pre-treated with HRV 3C protease for 1 h at 4 C. After removal of supernatant from the resin, 25 mg of protein was added to the resin and incubated for 1 h at 4 C. Resins were washed with 500 mL of PBS 3 times and elution performed using 40 mL of maltose buffer. Boiled samples with sample buffer were analyzed by SDS-PAGE and stained with One-step CBB (Biocraft). Gel images were captured by Gel Doc EZ (Bio-Rad) and quantitated by Image Lab software (Bio-Rad). Preparation and observation of Ape1 droplets Mixtures of Ape1 (GST-Ape1) were prepared at a 9:1 ratio of GST-CS-Ape1-CS-mCherry and GST-CS-Ape1-mCherry. 10 mM GSTApe1 was incubated in 20 mM HEPES pH 7.0, 150 mM sodium chloride, and 5 mM DTT for 5 min and supplemented with 1 mL of HRV 3C protease (7.33 mg/ml). GST-Ape1 with the protease was transferred to a glass-bottom dish (MatTek) or cover glass (Muto Pure Chemical) coated with 2% polyvinyl alcohol. To prevent sample evaporation, moist paper was placed in the dish, or the cover glass was sealed with uneven slide glass (Matsunami Glass). The samples were observed using a FV3000RS confocal laser scanning microscope (Olympus) equipped with 20x, 40x, and 60x objective lens (Olympus). To detect mKalama1 or C4N4, a 405 nm laser was used for excitation and 402 to 437 nm fluorescence signals were detected. To detect mClover3 and mCherry, 488 and 561 nm lasers were used and 488 to 521 and 565 to 595 nm signals were detected, respectively. To record droplet number and size, images were taken every 106 s. Analyses and quantitation of droplet number and size were performed in ImageJ, as described in a previous study (Schneider et al., 2012; Zaffagnini et al., 2018). For line profile measurements, the images were set to the correct scale and converted to an 8-bit image in ImageJ. The plot profiles of drawn lines were obtained using the ‘‘Plot Profile’’ tool. To observe the effect of 1,6-hexanediol in vitro, Ape1 condensates (1 h after generation) were incubated with 10% 1,6-hexanediol. To observe the effect of 1,6-hexanediol in vivo, cells were incubated in the presence of 20 mg/ml digitonin for 5 min before further supplementation of 5% 1,6-hexanediol. Microscope observation of Atg5 To observe Atg5-mNeonGreen, we used an inverted fluorescence microscope (IX81; Olympus) equipped with an electron-multiplying CCD camera (ImagEM C9100-13) and a 150x objective lens (UAPON 150x OTIRF, NA/1.45; Olympus). The further details were described previously (Adachi et al., 2017). In vitro translation of Ape1 using the PURE system Reconstituted cell-free protein synthesis was performed using the PUREfrex2.1 system according to the manufacture’s instructions (GeneFrontier). First, to construct expression plasmids encoding Ape1, the Ape1 gene was amplified by PCR and cloned into the pET23a (+) vector (Novagen). To enhance protein expression efficiency, AT-rich codons were used where possible at the second to sixth N-terminal codons of Ape1. NEBuilder HiFi DNA Assembly Cloning Kit (New England Biolabs) was used for cloning. Next, a protein synthesis reaction mixture was prepared with solution I, solution II, solution III, 0.5mM Cysteine, 4mM GSH, and a template plasmid (12ng per 1mL of reaction solution). The reaction solution was incubated at 37 C for indicated times. For the determination of the critical concentration for phase separation of Ape1, the reaction solution was centrifuged at 15,000 x g for 5 min at each time point. The samples (total and supernatant) were separated by SDS-PAGE and detected by One Step CBB. Four different concentrations of recombinant full-length Ape1 L11S were separated on the same SDS-PAGE gel to prepare a standard curve for calculating molar concentration of Ape1. Quantitative analysis was carried out with BioRad Gel Doc EZ Imager and Image Lab Software. FRAP experiments The same microscope system employed in Ape1 droplet observation was used for both in vitro and in vivo FRAP experiments. Fluorescence of Ape1WT droplets, Ape1P22L condensates or mClover3-Atg19 variants was bleached using 561 nm or 488 nm lasers set to 100% intensity for 1 s, with images subsequently recorded every 1.6 s or 1.2 s, respectively. The region of interest (ROI) was set to the bleached area. 100% and 0% fluorescence intensities were recorded within the ROI immediately before and after photobleaching. All fluorescence intensity readings were below the saturation threshold of the image capture system. Ape1 complex

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fluorescence in yeast cells was bleached using a 561 nm excitation wavelength set to 20% intensity for 0.2 s with subsequent recordings made every 1.6 s. To normalize ROI fluorescence, the background fluorescence value was subtracted. Immunoprecipitation Immunoprecipitation was performed as previously described (Yamasaki et al., 2016). Briefly, 25 OD600 units of yeast cells were resuspended in 200 mL buffer A (20 mM Tris pH 8.0, 150 mM sodium chloride, 10% glycerol), 5 mM EDTA, protease inhibitor cocktail, and 1 mM phenylmethylsulfonyl fluoride (Sigma-Aldrich), to which we added zirconia beads. After cell disruption by beads beater (MP BIOMEDICALS) for 2 min, 50 mL buffer B (20 mM Tris pH 8.0, 150 mM sodium chloride, 10% glycerol, 0.5% Triton X-100) were added to samples, followed by a 1 h incubation at 4 C. The supernatants were recovered after centrifugation at 15000 g for 15 min. 200 mL of these lysates (20 OD600 volume) were used for immpunoprecipitation and mixed with 1 mg anti-GFP antibody (Roche) and 10 mL Protein G resin (Recenttec) for 2 h at 4 C. The resin was washed with the same lysate buffer 3 times and then boiled in 25 mL sample buffer containing 20 mM DTT at 98 C for 5 min. The samples were subsequently analyzed by western blotting. Western Blotting Western blotting was performed as previously described (Yamasaki et al., 2016). Briefly, yeast cells were pelleted and resuspended in 20% TCA for 10 min on ice. After centrifugation, the supernatant was removed. The pellet was washed with ice-cold acetone, dried at room temperature, and then supplemented with sample buffer (100 mM Tris pH 7.4, 2% SDS, 10% glycerol, 20 mM DTT, 0.05% Bromophenol blue, 1% protease inhibitor cocktail (Nacalai tesque), 0.1 mg/mL PMSF). Samples were disrupted with 0.6 mm zirconia silica beads (Bio Medical Science) for 2 min using a cell disruptor (Scientific Industries). The sample lysates were recovered after 2 min centrifugation at 15,000 g. Samples boiled with sample buffer were applied to 10% or 12% acrylamide gels. Rabbit antiApe1 serum (1/1000 or 1/10000 dilution, (Suzuki et al., 2002)), rabbit anti-Atg19 serum (1/1000 dilution, (Watanabe et al., 2010)) and mouse anti-PGK1 antibody (0.5 mg/mL, Invitrogen) were used as primary antibodies. Goat anti-mouse IgG-peroxidase labeled antibody and goat anti-rabbit IgG-peroxidase labeled antibody (Sigma-Aldrich) or horseradish peroxidase-labeled anti-rabbit secondary antibody (Promega) were used at 1/10000 dilution or 1/5000 as secondary antibodies, respectively. The membrane was soaked in Immobilon Western chemiluminescent HRP substrate (Merck-Millipore) or enhanced chemiluminescence reagent (GE Healthcare) and then analyzed by ImageQuant LAS 4000 mini (GE healthcare), IR-LAS 1000 imaging system (FUJIFILM), or FUSION FX7 system (Vilber-Lourmat). Sample preparation for HS-AFM observation For HS-AFM imaging, either coverslips (24 3 32 mm2, 0.13-0.17 mm thick) (Matsunami Glass Ind., Ltd., Osaka, Japan) or mica (~3 3 ~3 mm2) attached to coverslips were used as a solid support. Coverslips were immersed in 5 M KOH solution for 1 h, followed by three washes in Milli-Q water. Subsequently, the cleaned coverslips were sonicated in Milli-Q water for 20 min and stored in ethanol at 4 C until use. Ethanol was completely removed before each experiment. For HS-AFM observation of Ape 1 droplets, PreScission protease was applied to GST-Ape1WT or GST-Ape1P22L (2 mM) in 10 mL of reaction buffer (150 mM NaCl, 20 mM HEPES-NaOH pH 7.0, 5 mM DTT) to remove the GST-tag. After incubation at 23 C for 20 min, 10 mL of reaction buffer was added to the reaction mixture. Next, 20 mL of sample was deposited onto cleaned coverslips. After a 5 min incubation, excess proteins were washed out with observation buffer (600 mM NaCl, 20 mM HEPES-NaOH pH 7.0). 0.01% glutaraldehyde diluted by observation buffer was applied onto the coverslip to fix Ape1 droplets loosely. After incubation for 5 min, the coverslip was rinsed in observation buffer. For HS-AFM observation of Ape1L11S dodecamer, PreScission protease was applied to GST-Ape1L11S (2 mM) in 10 mL of reaction buffer (150 mM NaCl, 20 mM HEPES-NaOH pH 7.0, 5 mM DTT) to remove the GST-tag. After incubation at 23 C for 20 min, the reaction mixture was diluted to 1/20 with reaction buffer. Then 20 mL of the sample was deposited onto cleaved mica. After a 5 min incubation, excess protein was washed away using observation buffer. For HS-AFM observation of the mApe1 dodecamer, mApe1 (0.1 mM) in 20 mL of 20 mM HEPES-NaOH pH 7.0 containing 150 mM NaCl and 5 mM DTT was deposited onto cleaved mica. After a 5 min incubation, excess protein was washed away using observation buffer. HS-AFM imaging HS-AFM images were acquired in tapping mode using a tip-scan type HS-AFM instrument (Fukuda et al., 2013) (Nano Explorer PSNEX, Research Institute of Biomolecule Metrology Co., Ltd., Ibaraki, Japan) equipped with a fluorescence microscope and 100x objective lens (Olympus). We used cantilevers measuring ~9 mm long, ~2 mm wide and ~0.13 mm thick with a resonant frequency of ~1.5 MHz and a spring constant of 0.1–0.2 N/m (BL-AC10DS, Olympus). Ape1 droplets were selected for observation by mCherry fluorescence and located in the HS-AFM scanning area (~4 3 ~6 mm2) prior to nanoscale imaging by HS-AFM. HS-AFM imaging conditions were as follows: scan size and pixel size, 1000 3 500 nm2 (120 3 60 pixels) or 240 3 96 nm2 (120 3 48 pixels) for Ape1 droplets and 120 3 48 nm2 (120 3 48 pixels) for Ape1L11S and mature Ape1; imaging rate, ~1.18 frames/sec (fps) or ~3.13 fps for Ape1 droplets and 2 fps for Ape1L11S and mature Ape1; cantilever free oscillation amplitude, 5-10 nm. The set-point amplitude was 90% of the free amplitude. All imaging was performed at 23 C.

e7 Molecular Cell 77, 1–13.e1–e9, March 19, 2020

Please cite this article in press as: Yamasaki et al., Liquidity Is a Critical Determinant for Selective Autophagy of Protein Condensates, Molecular Cell (2019), https://doi.org/10.1016/j.molcel.2019.12.026

HS-AFM image processing and analysis The free software ImageJ was used to process and analyze images. To enhance the surface contrast of droplets, an FFT bandpass filter was applied to images (Figures 3D and 3E). The migration distance (Figure 3H) was calculated using filtered images. The local maxima in each image (Figures 3F and 3G) were detected by the ‘‘Find Maxima’’ function, excluding points at the edges of droplets. Preparation of GUVs A natural swelling method is used to prepare GUVs containing PE in buffer from a dry lipid film (Alam et al., 2012). To prepare GUVs using electrically neutral lipids in a buffer at physiological ion concentration (~150 mM NaCl), we used a very small fraction of the PEGlipid (i.e., PEG-lipid method) (Alam and Yamazaki, 2011; Alam et al., 2012). We prepared 200 mL of a 1 mM phospholipid mixture consisting of 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC), 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine (POPE), L-a-phosphatidylinositol (Liver, PI), and 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-2000 (DPPE-PEG2000) (all from Avanti Polar Lipids) at a molar ratio of 59:30:10:1 in chloroform in a 5 mL glass vial. We then produced a homogeneous thin film of lipid mixture by evaporation of chloroform by the gentle application of nitrogen gas. For the complete removal of chloroform, we then put the glass vial in a vacuum desiccator connected to a rotary pump overnight. Next, we prehydrated thin lipid film in the bottom of the glass vial with 20 mL of water at 60 C for 7 min. Thereafter 1 mL of HEPES buffer (20 mM HEPES, pH 7.0, 150 mM NaCl, 1 mM EGTA) containing 0.1 M sucrose was added and incubated for 2-3 h at 60 C. Observation of GUVs 100 mL of GUV solution (containing 0.1 M sucrose solution as the internal solution) was diluted into 900 mL of HEPES containing 0.1 M glucose solution (external solution) into a hand-made microchamber that was formed on a glass slide (Matsunami Glass) by depositing (in parallel) 2 bar-shaped silicon-rubber (3mm silicon sheet) spacers between a coverslip (Micro cover glass, Muto Pure Chemicals) and the glass slide (Parvez et al., 2018). The microchamber was coated with 0.10% (w/v) BSA in the same buffer as that used for the experiments to avoid strong contact between GUVs and the glass surface. 0.1M sucrose solution and 0.1M glucose solution were used at the inside and outside solutions of GUVs in these experiments to enhance the contrast of GUVs for differential interference contrast (DIC) observation. Observation of GUVs was performed using an FV3000RS at room temperature (~23 C), as described above. The single GUV method for observation of GUV-protein-droplet interactions The single-giant unilamellar vesicle (GUV) method, which can add different proteins one by one in the vicinity of a GUV precisely, was used for observation of GUV-protein interactions (Yamazaki, 2008). Purified proteins for Atg8 lipidation and either Ape1 droplets, Atg19WT/Atg19DN-coated Ape1 droplets or Ape1P22L condensates in HEPES buffer containing 0.1 M glucose were added slowly one by one into the vicinity of a single GUV through a 15-20 mm diameter glass micropipette, the position of which was controlled by a micromanipulator (Narishige) at room temperature (Parvez et al., 2018). The distance between the GUV and the tip of the micropipette was maintained at ~50 mm. A glass micropipette was prepared as follows; first we pulled a glass tube of 1.0 mm diameter (Narishige) to a needle point using a puller (Narishige), and then broke it by micro-forage (Narishige) at the desired tip diameter. Proteins and droplets in the external solution of GUVs were filled in the micropipette by aspiration using a vacuum pump (ULVAC KIKO). The micropipette was held by a micromanipulator, allowing us fine control over the position of the tip of the micropipette in relation to the GUV. The application pressure in the vicinity of the GUV was controlled by changing the height of vertical column of water to which the micropipette was hydraulically connected (Yamazaki, 2008). The application pressure was measured using a differential pressure transducer (Validyne), pressure amplifier (Karone), and a digital multimeter. For the constant application of proteins or droplets in the vicinity of GUVs using the micropipette, we first determined the equilibrium pressure by approaching the tip of micropipette to a small vesicle and adjusting the pressure to keep the vesicle at the tip of the micropipette. After fixing equilibrium pressures, an additional ~300 mV was applied to constantly supply the sample solution in the vicinity of GUVs. We then observed interactions at a range of time points. For the lipidation of Atg8 on the GUV membrane we introduced a mixture of 100 mM mKalama1-Atg8, 10 mM Atg3, 10 mM Atg7, 2 mM Atg12-5-16, 2 mM ATP and 0.5 mM MgCl2 in the vicinity of GUVs from the micropipette. Preparation methods for Ape1 droplets and Ape1P22L condensates were based on the above method, where concentrations of 20 mM (Ape1WT), 2.5 mM (Ape1P22L), and 1 mM of Atg19WT/Atg19DN was used for subsequent reactions. Thereafter, protein-coated droplets or condensates were introduced to GUVs/Atg8-lipidated GUVs as a mixture from the micropipette. Micropipette manipulation for GUV-droplet tethering measurement A micropipette aspiration method was used to hold the GUV or Atg8 lipidated-GUV on the tip of a micropipette (Parvez et al., 2018; Rawicz et al., 2000). A ~10mm-diameter micropipette, which was coated with 0.50% (w/v) BSA in HEPES buffer containing 0.10 M glucose, was used to hold the GUV by applying aspiration pressure, DP ( = Pout
Pin), where Pout and Pin are pressures on the outside and the inside of the micropipette, respectively (Parvez et al., 2018). The lateral tension of the GUV membrane, s, was maintained at 0.50 mN/m by adjusting DP. s can be calculated by the following equation (Parvez et al., 2018): s=

DPdP 4ð1
dP =DV Þ

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Please cite this article in press as: Yamasaki et al., Liquidity Is a Critical Determinant for Selective Autophagy of Protein Condensates, Molecular Cell (2019), https://doi.org/10.1016/j.molcel.2019.12.026

where dp is the internal diameter of the micropipette and Dv is the diameter of the spherical cap segment (on the outside of the micropipette) of the aspirated GUV. This small aspirated GUV or Atg8 lipidated-GUV was delivered to the Ape1 droplets/condensates by micropipette manipulation for tethering measurement. After contact of GUV with droplets/condensates, tethering was confirmed by the application of forward and backward force using micropipette manipulator. Interaction of Atg8-LUV with Ape1 droplets in the presence and absence of Atg19 LUVs were prepared by the extrusion method (Tamba and Yamazaki, 2005). Briefly, we first prepared multilamellar vesicles (MLVs) by adding 1 mL of HEPES buffer (20 mM HEPES, pH 7.0, 150 mM NaCl, 1mM EGTA) to dry lipid film composed of POPC/POPE/PI/ NBD-PE/DPPE-PEG2000 (59:29:10:1:1). Samples were then vortexed 5 times at 20 s intervals at room temperature. Next, the MLV suspension was subjected to five cycles of freezing in liquid N2 for ~30 s, followed by warming at room temperature (freeze
thawing). The resulting solution was extruded through a 400 nm pore size Nuclepore track-etched membrane using a Mini Extruder (Avanti Polar Lipids) until the solution became transparent. In order to remove liposomes smaller than 400 nm, we further purified LUVs using a membrane filtration method (400 nm filter size) (Tamba et al., 2011). Following this, samples were subjected to interaction analysis with Ape1 droplets using the single GUV method described above. To this end, we first prepared Ape1 droplets in HEPES buffer and transferred them into a homemade silicone micro-chamber (coated using 0.10% (w/v) BSA) with or without Atg19. We then used a micropipette to apply LUVs and other components of the Atg8 conjugation machinery into the vicinity of Ape1 or Atg19 blanketed-Ape1 droplets and observed liposome tethering on the surface of Ape1 droplets. Visualization of IMs and Ape1 complexes by fluorescence microscopy Ape1WT-GFP cells harboring the pYEX-BX[APE1] plasmid and Ape1P22L-GFP cells carrying the pYEX-BX[APE1P22L] plasmid were cultured in the presence of 250 mM CuSO4 for > 1 day to express Ape1 from the CUP1 promoter. After dilution, cells were cultured again until log phase (~2 3 107 cells per ml; OD600 = ~1) in SDCA medium containing 250 mM CuSO4. To induce autophagy, 200 ng/ml rapamycin (R-5000, LC Laboratories, USA) was added. An IX83 inverted fluorescence microscope (Olympus) equipped with an UPlanSApo100 3 /1.40 Oil (Olympus) and a CoolSNAP HQ CCD camera (Nippon Roper) was used for fluorescence imaging. UFGFP and U-FMCHE filter sets (Olympus) were used for GFP and mCherry visualization, respectively. Images were acquired using the MetaVue imaging software (Molecular Devices). Samples were prepared using 76 3 26 mm slide glass (S1225, Matsunami) and 18 3 18 mm micro cover glass (No. 1-S, Matsunami). Morphological analysis of autophagy-related structures and Ape1 complexes Morphological features of autophagy-related structures were analyzed with the Qautas (Quantitative autophagy-related structures analysis system) (Kawaoka et al., 2017) based on image processing using ImageJ and a machine learning discriminator using R software (version 3.1.3, R Core Team (2018). R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria. https://www.R-project.org/) with the randomForest library (version 4.6–10, https://cran.r-project.org/ web/packages/randomForest/randomForest.pdf). In brief, fluorescence microscopic images were normalized and processed with a bandpass filter. Autophagy-related structures were extracted with binarized using the ‘RenyiEntropy’ algorithm. Nine morphological parameters (area, major axis length, minor axis length, perimeter, angle, circularity, aspect ratio, roundness, and solidity) of each structure were extracted by particle analysis. Extracted structures were discriminated into dot-shaped and elongated structures by the random forest algorithm. Lengths of IMs were calculated as a half of the ‘perimeter’ value. Morphological features of Ape1 complexes were analyzed using ImageJ and R software. Fluorescence microscopy images were normalized and processed with a bandpass filter. Ape1 complexes were extracted by binarization using the ‘Li’ algorithm. Six morphological parameters (area, perimeter, circularity, aspect ratio, roundness, and solidity) of each structure were extracted by particle analysis. Statistical Analysis Statistical analyses were performed by Mann-Whitney U test for Figures 6B–6D and by unpaired Student’s t test for all the other data; p > 0.05, not significant; *p < 0.05, **p < 0.01 and ***p < 0.001. DATA AND CODE AVAILABILITY Original microscope images and blots are available at Mendeley data (https://doi.org/10.17632/k4dsc87ym3.1).

e9 Molecular Cell 77, 1–13.e1–e9, March 19, 2020