MBoC | ARTICLE

CCPG1 recognizes endoplasmic reticulum luminal
proteins for selective ER-phagy
Shunsuke Ishiia,†, Haruka Chinob,†,‡, Koji L. Odec, Yoshitaka Kurikawab, Hiroki R. Uedac,d,Akira Matsuurae, Noboru Mizushimab,*, and Eisuke Itakurae,*aDepartment of Biology, Graduate School of Science and Engineering, Chiba University, Chiba 263-8522, Japan;
Department of Biochemistry and Molecular Biology, Graduate School of Medicine, Tokyo 113-0033, Japan;c
Department of Systems Pharmacology, Graduate School of Medicine, University of Tokyo, Tokyo 113-0033, Japan;dLaboratory for Synthetic Biology, RIKEN Center for Biosystems Dynamics Research, Osaka 565-0871, Japan;eDepartment of Biology, Graduate School of Science, Chiba University, Chiba, 263-8522, Japanb

ABSTRACT The endoplasmic reticulum (ER) is a major cell compartment where protein synthesis, folding, and posttranslational modifications occur with assistance from a wide varietyof chaperones and enzymes. Quality control systems selectively eliminate abnormal proteinsthat accumulate inside the ER due to cellular stresses. ER-phagy, that is, selective autophagyof the ER, is a mechanism that maintains or reestablishes cellular and ER-specific homeostasis
through removal of abnormal proteins. However, how ER luminal proteins are recognized by
the ER-phagy machinery remains unclear. Here, we applied the aggregation-prone protein,
six-repeated islet amyloid polypeptide (6xIAPP), as a model ER-phagy substrate and foundthat cell cycle progression 1 (CCPG1), which is an ER-phagy receptor, efficiently mediates its
degradation via ER-phagy. We also identified prolyl 3-hydroxylase family member 4 (P3H4) asan endogenous cargo of CCPG1-dependent ER-phagy. The ER luminal region of CCPG1 contains several highly conserved regions that we refer to as cargo-interacting regions (CIRs);
these interact directly with specific luminal cargos for ER-phagy. Notably, 6xIAPP and P3H4
interact directly with different CIRs. These findings indicate that CCPG1 is a bispecific ERphagy receptor for ER luminal proteins and the autophagosomal membrane that contributes
to the efficient removal of aberrant ER-resident proteins through ER-phagy.

This article was published online ahead of print in MBoC in Press (http://www.molbiolcell.org/cgi/doi/10.1091/mbc.E22-09-0432) on February 3, 2023.
†
These authors contributed equally to this work.
‡
Present address: Department of Cell Biology, Harvard Medical School, Boston,
MA 02115.
Author contributions: S.I. and H.C. performed the experiments; K.L.O. performed
mass spectrometry under the supervision of H.R.U.; Y.K. provided experimental
support; A.M. contributed to the writing and data interpretation; S.I., H. C., N. M.,
and E.I. conceptualized the project and wrote the manuscript.
Conflict of interest: The authors declare no conflicts of interest.
*
Address correspondence to: Eisuke Itakura (eitakura@chiba-u.jp); Noboru
Mizushima (nmizu@m.u-tokyo.ac.jp).
Abbreviations used: ATG, autophagy-related; CCPG1, cell cycle progression 1;
CIR, cargo-interacting region; FIR, FIP200-interacting region; IAPP, islet amyloid
polypeptide; LIR, LC3-interacting region; P3H4, prolyl 3-hydroxylase family member 4; RG, RFP-GFP; WT, wild type.
© 2023 Ishii et al. This article is distributed by The American Society for Cell Biology under license from the author(s). Two months after publication it is available
to the public under an Attribution–Noncommercial-Share Alike 4.0 International
Creative Commons License (http://creativecommons.org/licenses/by-nc-sa/4.0).
“ASCB®,” “The American Society for Cell Biology®,” and “Molecular Biology of
the Cell®” are registered trademarks of The American Society for Cell Biology.

Molecular Biology of the Cell • 34:ar29, 1–18, April 1, 2023

Monitoring Editor
Sharon Tooze
The Francis Crick InstituteReceived: Sep 26, 2022
Revised: Dec 23, 2022
Accepted: Jan 27, 2023

INTRODUCTION
The endoplasmic reticulum (ER) is the largest organelle and synthesizes approximately 35% of the proteins in a cell (Uhlén et al., 2015;
Juszkiewicz and Hegde, 2018). These proteins mature through processes occurring in the ER, including cleavage of signal sequences,folding, disulfide bond formation, and glycosylation. Then the proteins are selectively transported to the endomembrane system, theplasma membrane, or the exterior of the cell (Ellgaard and Helenius,
2003; Bukau et al., 2006; Ni and Lee, 2007). However, external
stresses and genetic mutations can lead to the accumulation of unfolded proteins in the ER, causing ER storage diseases (e.g., hereditary emphysema; Callea et al., 1992; Rutishauser and Spiess, 2002;
Hebert and Molinari, 2007). Unfolded proteins accumulated in the
ER are recognized by unfolded protein response (UPR) receptors(IRE1, PERK, and ATF6), which induce molecular chaperones to refold or degrade the proteins through ER-associated degradation(ERAD); this is known as the ER stress response pathway (Chakrabarti
et al., 2011; Ruggiano et al., 2014; Hetz et al., 2020). However, not34:ar29, 1  all misfolded proteins in the ER are degraded through ERAD (Houcket al., 2014; De Leonibus et al., 2019).
Macroautophagy (hereafter referred to as autophagy) is a bulk
degradation system in which an isolation membrane engulfs a portion of the cytoplasm and delivers it to lysosomes for degradation(Mizushima and Komatsu, 2011; Mercer et al., 2018; Melia et al.,2020; Klionsky et al., 2021). Recent studies have revealed that large
cellular structures, such as organelles (Lemasters, 2005; Singh et al.,2009; Maejima et al., 2013), ribosomes (Kraft et al., 2008), protein
aggregates (Webb et al., 2003; Øverbye et al., 2007), invading
pathogens (Nakagawa et al., 2004), lysosomes (Maejima et al., 2013)and protein droplets (Wilfling et al., 2020; Yamasaki et al., 2020), areselectively degraded via autophagy. This selectivity is achieved
through autophagy receptors, which link cargos to the autophagosomal membrane. The interaction motifs between autophagy receptors and autophagy-related gene (ATG) proteins are evolutionarily
conserved, as several proteins contain the microtubule-associatedprotein light chain 3 (LC3)-interacting region (LIR). The LIR contributes to selective interaction with cargos, which are recruited to the
autophagosomal membrane (Stolz et al., 2014). The FIP200-interacting region (FIR) is another binding motif that links cargos to FIP200,
a component of the ULK1 complex that is indispensable to autophagy induction (Smith et al., 2018; Turco et al., 2019). ER-phagy (reticulophagy) is a type of autophagy that can selectively remove theER (De Duve, 1963; Bernales et al., 2006) and maintains ER homeostasis through remodeling of ER conformation via the degradation ofexcessive membrane or removal of unfolded luminal proteins (Jiaet al., 2011; Pengo et al., 2013; Molinari, 2021).
Since the identification of yeast ATG39 and ATG40, as well asmammalian FAM134B, as ER-phagy receptors (Khaminets et al.,2015; Mochida et al., 2015), several additional ER-phagy receptor
proteins have been identified, including SEC62 (Fumagalli et al.,2016), RTN3L (Grumati et al., 2017), cell cycle progression 1 (CCPG1)(Smith et al., 2018), TEX264 (An et al., 2019; Chino et al., 2019),
ATL3 (Chen et al., 2019), CALCOCO1 (Nthiga et al., 2020), C53(Stephani et al., 2020), Epr1 (Zhao et al., 2020), and RHD3 (Sunet al., 2022). All ER-phagy receptors tether the ER membrane to the
autophagosomal membrane through interaction with Atg8/LC3(Mochida and Nakatogawa, 2022). CCPG1 also interacts with FIP200(Smith et al., 2018). Each receptor is responsive to diverse intracellular stresses. The reticulon-like proteins ATG40 and FAM134B haveshort hairpin transmembrane domains and generate membrane curvature, leading to fragmentation of the ER and incorporation into
the autophagosome under starvation conditions (Bhaskara et al.,2019; Jiang et al., 2020; Mochida et al., 2020). SEC62 causes ER
degradation during recovery from ER stress (Fumagalli et al., 2016),
while CCPG1 promotes peripheral ER degradation during ER stress(Smith et al., 2018). TEX264 ensures the canonical degradation of
the ER via ER-phagy and is regulated by phosphorylation of its LIR(Chino et al., 2019, 2022). SEC62, CCPG1, and TEX264 all have one
or two transmembrane domains, and only CCPG1 has a long intraluminal domain.
In the UPR pathway, luminal domains of UPR receptors on the ER
membrane sense unfolded luminal proteins in the ER and activate
UPR signaling (Schröder and Kaufman, 2005). Although bulk incorporation of the ER into autophagosomes via ER-phagy receptors
has been studied thoroughly, the process of selective protein recognition inside the ER during ER-phagy remains largely unexplored.
Recent studies have revealed that several ER-resident proteins link
ER-luminal cargos to ER-phagy receptors. Calnexin and binding immunoglobulin protein (BiP) are ER chaperones that deliver misfolded proteins to the ERAD machinery and also interact with
2 | S. Ishii et al.

FAM134B to mediate ER-phagy for misfolded procollagen (Fregno
et al., 2018; Forrester et al., 2019). PGRMC1, which is a binding
partner of ER-phagy receptor RTN3L, captures low-molecularweight misfolded proteins (Chen et al., 2021). However, whether
any of these ER-phagy receptors can directly recognize ER luminal
proteins for selective ER-phagy remains unclear.
In this study, we characterized an ER luminal protein, six-repeated
islet amyloid polypeptide (6xIAPP), as a model ER-phagy substrate
in mammalian cells and found that aggregated 6xIAPP is efficiently
degraded by ATG protein- and CCPG1-dependent ER-phagy processes. We found that the luminal region of CCPG1 is essential for
ER-phagic degradation of selective proteins and identified prolyl
3-hydroxylase family member 4 (P3H4) as an endogenous ER luminal protein recognized by CCPG1. CCPG1 possesses several highly
conserved regions among vertebrates, and 6xIAPP and P3H4 interact directly with different parts of the luminal CCPG1 domains,which contain cargo-interacting regions (CIRs). Thus, our data revealed that CCPG1 luminal regions directly recognize ER luminal
cargos for ER-phagy, and simultaneous binding of multiple cargos
to CCPG1 might drive the efficient removal of aberrant ER-resident
proteins.

RESULTS
6xIAPP-RFP-GFP-KDEL is delivered to the lysosomes
Islet amyloid polypeptide (IAPP) is a 37-amino acid (aa) protein secreted from pancreatic islet β-cells, and aggregation of IAPP is associated with type 2 diabetes (Klöppel et al., 1985; Butler et al.,2003). As 6xIAPP forms oligomers and induces severe ER stress inyeast (Kayatekin et al., 2018), we examined whether 6xIAPP is degraded through ER-phagy in mammalian cells. To monitor lysosomal
degradation of IAPP, 6xIAPP or 1xIAPP was fused with red fluorescent protein (RFP), green fluorescent protein (GFP), and an ER retention signal, KDEL (forming IAPP-RFP-GFP-KDEL, hereafter referred
to as IAPP-RG) and inserted into a lentivirus vector under a doxycycline (Dox)-dependent promoter (Figure 1A). In mammalian cells,
6xIAPP-RG showed reticular and dotlike localization patterns andincreased molecular mass, compared with 1xIAPP-RG, which was
similar in size to the 4.3-MDa ribosome complex (60S ribosome protein L19; RPL19) (Supplemental Figure S1, A and B). Glycosylation
can be an indicator of proteins translocated into the ER lumen. Because IAPP-RG does not have any glycosylation sites, its electrophoretic mobility was not altered by treatment with endoglycosidase H(Endo H; Supplemental Figure S1C). Therefore, we artificially inserted an opsin glycosylation site into the C-terminus of each IAPP
construct (IAPP-Gly-HA-KDEL and IAPP-RG-Gly-KDEL). These proteins (both 1x and 6xIAPP) underwent glycosylation in mammaliancells (Supplemental Figure S1, D–G). Membrane permeabilization
assays also showed that, while cytosol-facing RFP-GFP-cytochromeb5 (RG-Cytb5; a transmembrane domain of cytochrome b5 fusedwith RFP-GFP) was readily stained with anti-GFP antibody after digitonin treatment, 6xIAPP-RG was stained only after cells had beentreated with the stronger detergent Triton X-100 but not digitonin(Supplemental Figure S1H). These results suggest that 6xIAPP in
mammalian cells primarily resides in the ER luminal environment at
steady state.
When IAPP-RG is delivered to the lysosomes, the acidic conditions and lysosomal proteases rapidly attenuate GFP fluorescence.
In contrast, RFP, which is relatively resistant to the lysosomal environment, accumulates within the lysosomes (Katayama et al., 2008).
Therefore, an increase in the RFP signal of lysosomes detected
through fluorescence microscopy, elevation of the RFP/GFP ratio
based on flow cytometry, and detection of cleaved RFP throughMolecular Biology of the CellFIGURE 1: 6xIAPP-RFP-GFP-KDEL is delivered to lysosomes. (A) Schematic diagrams of the ER-phagy substrate 1x (or6x) IAPP-RFP-GFP-KDEL (1x or [6x] IAPP-RG). In the lysosome, GFP fluorescence is diminished, whereas RFP fluorescenceremains intact due to resistance to acidic pH and lysosomal proteases. (B–D) 6xIAPP, but not 1xIAPP, is delivered tolysosomes. Tet-On HeLa cells expressing 1x (or 6x) IAPP-RG were incubated in medium containing doxycycline (Dox) for
24 h and then cultured in medium with or without Dox for a further 24 h. After fixation, the cells were stained with
antibodies against LAMP1 and analyzed by confocal microscopy. GFP-negative and RFP-positive signals accumulated in
LAMP1-positive structures (arrowhead) are indicated. Scale bars represent 10 and 1 µm (inset) (B). The cells treated with
Dox as described above were trypsinized, and their green and red fluorescence intensities were measured using flow
cytometry. Representative dot plots of GFP versus RFP intensities and corresponding histograms are shown, C. The
fluorescence ratio was calculated in RFP-positive cells. Data represent the mean ± standard error (SE) of at least three
independent experiments. Differences were statistically analyzed by Dunnett’s multiple comparison test, D.
(E, F) Proteasomal inhibition is not related to lysosomal degradation of 6xIAPP. Tet-On HeLa cells expressing 1x (or 6x)
IAPP-RG were cultured in medium containing Dox with dimethylsulfoxide (DMSO), bafilomycin A1, or bortezomib for
24 h before immunoblotting analysis using antibodies against RFP, HSP90, and Histone H3. Cell lysates were centrifuged,
and the supernatants were collected as the soluble fraction. The pellets containing the nuclear compartment werecollected as the insoluble fraction. Each pellet was resuspended in an equal volume of lysis buffer. # indicatesdegradative products of IAPP-RG, E. The band intensities of cleaved RFP and 6xIAPP-RG were quantified, and the ratioof cleaved RFP/total amount of 6xIAPP (normalized to DMSO treatment) is shown. Data represent the mean ± SE ofthree independent experiments. Differences were statistically analyzed by Dunnett’s multiple comparison test, F.
Volume 34 April 1, 2023

CCPG1 ER-phagy| 3

FIGURE 2: ER-phagy degrades 6xIAPP in the ER. (A, B) Dot structure of 6xIAPP-RG colocalized with endogenous LC3.
Tet-On HeLa cells expressing 1x (or 6x) IAPP-RG were incubated in medium containing Dox for 48 h before fixation. The
cells were stained with antibodies against LC3 and analyzed through confocal microscopy. Scale bars represent 10 and
1 µm (inset), A. Quantification of the number of LC3 puncta per cell. Solid bars indicate the median, boxes theinterquartile range (25th–75th percentile), and whiskers the 0th–100th percentile range. Data were collected from33 cells of each cell type. Differences were analyzed using one-way analysis of variance (ANOVA) and Sidak’s multiple
comparison test, B. (C–E) 6xIAPP-RG accumulated in autophagy-KO cells. Tet-On HeLa cells stably expressing Cas9 andthe indicated sgRNA were cultured in the presence of Dox for 24 h (a mixed population of wild-type [WT] and knockout[KO] cells was used). Following the removal of Dox, the cells were cultured with or without bafilomycin A1 for 24 h
before flow cytometry. Dot plots of GFP versus RFP fluorescence intensities are shown, with black letters indicating thelysosomal fraction (%) and red letters all other fractions (%), C. The fluorescence ratio was calculated for RFP-positive
cells. Data represent the mean ± SE of three independent experiments. Differences were analyzed using one-way
4 | S. Ishii et al.

Molecular Biology of the Cell

immunoblotting can be interpreted as representing lysosomal degradation of IAPP-RG.
1xIAPP-RG was clearly colocalized with translocon-associatedprotein subunit alpha (TRAPα), which is an ER marker (Supplemental Figure S1I), but not with lysosomal-associated membrane protein 1 (LAMP1), which is a lysosome marker (Figure 1B). On theother hand, although 6xIAPP-RG showed a reticular pattern andnumerous dot structures colocalized with the TRAPα signal, GFPnegative and RFP-positive signals of 6xIAPP-RG accumulated in
LAMP1-positive lysosomes, and RFP signals associated with lysosomes remained after Dox removal (Figure 1B and Supplemental
Figure S1I). We next quantified the reduction in fluorescence afterDox removal through flow cytometry. Although 1xIAPP-RG wasstable, 6xIAPP-RG was rapidly degraded within 24 h of Dox removal (Figure 1C). Notably, the RFP/GFP ratio of 6xIAPP-RG was
efficiently increased to 2.5-fold higher than that of 1xIAPP-RG(Figure 1, C and D), and this increase was clearly suppressed by thelysosomal inhibitor bafilomycin A1 (Figure 1D; Supplemental
Figure S2A). Treatment with bafilomycin A1 enlarged lysosomes
and caused the accumulation of GFP and RFP signals of 6xIAPPRG, but not 1xIAPP-RG, inside lysosomes (Supplemental Figure
S2B). We confirmed that 6xIAPP-HA-KDEL (6xIAPP-HA), in which
HA was used instead of RFP-GFP, also accumulated upon bafilomycin A1 treatment; these findings suggested that lysosomal degradation of 6xIAPP is not mediated by the RFP-GFP tag (Supplemental Figure S2, C and D). Moreover, both 6xIAPP-RG and 6xIAPP-HA
accumulated inside lysosomes after the inactivation of lysosomal
proteases using a lysosome inhibitor cocktail that contained E64d,
pepstatin A, and leupeptin (Supplemental Figure S2E). Flow cytometric analysis also demonstrated a significant reduction in the
RFP signal for 6xIAPP-RG. A previous report revealed that 6xIAPP
on the ER is degraded by the proteasome in yeast (Kayatekin et al.,2018). Indeed, treatment not only with bafilomycin A1, but alsowith the proteasomal inhibitor bortezomib, led to 6xIAPP-RG accumulation in the insoluble fraction (Figure 1E). Importantly, whilebafilomycin A1 clearly inhibited the generation of cleaved RFP,
bortezomib neither reduced nor increased the amount of cleaved
RFP derived from 6xIAPP-RG (Figure 1, E and F), indicating that theproteasome system is not associated with lysosomal degradation
under these conditions. The 6xIAPP accumulated due to proteasome inhibition is likely located in the cytosol, where it cannot be
removed through ER-phagy. We found that both GFP and RFP signals in cells expressing low 6xIAPP-RG levels were diminished afterDox removal. In contrast, only the GFP signal was markedly reduced in cells strongly expressing 6xIAPP-RG (Supplemental
Figure S2, F and G). Although most of the 6xIAPP that accumulated under proteasome inhibitor treatment was not glycosylated,some glycosylated 6xIAPP accumulated under lysosomal inhibitor(Supplemental Figure S2H). These results indicate that 6xIAPP exists in two distinct populations in mammalian cells, undergoing lysosomal and proteasomal degradation, respectively. Cells highly
expressing 6xIAPP-RG were used to detect lysosomal degradationin subsequent experiments.

6xIAPP in the ER is degraded by ER-phagy
GFP dot structures of 6xIAPP-RG, but not 1xIAPP-RG, were colocalized with LC3, an autophagosome marker (Figure 2A). To verify
whether 6xIAPP is degraded by autophagy, HeLa cells expressing
6xIAPP-RG with core autophagy genes knocked out, FIP200 (Haraet al., 2008), ATG5 (Mizushima et al., 2001), or ATG9A (Saitoh et al.,2009), were generated using clustered regularly interspaced shortpalindromic repeats (CRISPR). Quantitative analysis via flow cytometry revealed that the reduction of GFP signals of 6xIAPP-RG afterDox removal was significantly suppressed in the autophagy-KO cells(Figure 2, C and D; Supplemental Figure S3A). Furthermore, FIP200KO cells showed diminished colocalization of 6xIAPP-RG with LC3
and fewer RFP dots with LAMP1-positive structures (Figure 2E).
These data suggest that 6xIAPP is degraded in an autophagy-dependent manner.
We observed that the number of endogenous LC3 dots was increased when 6xIAPP-RG was expressed (Figure 2, A and B). To
quantify autophagic activity, 1x or 6xIAPP-HA under the Tet-On promoter was introduced into HeLa cells stably expressing RFP-GFPLC3 as a ratiometric autophagy probe (Kimura et al., 2007). With theexpression of 6xIAPP-HA, but not 1xIAPP-HA, the RFP/GFP ratio of
RFP-GFP-LC3 was elevated 2.7-fold and increased numbers of GFP
punctate structures were observed within 48 h of Dox induction(Supplemental Figure S3, B–D). In addition, the amount of endogenous LC3-II increased with expression of the 6xIAPP construct(Supplemental Figure S3, E and F). These data suggest that autophagic activity is promoted by the expression of 6xIAPP.
If ER-phagy degraded 6xIAPP present in the ER lumen, 6xIAPP
should be degraded along with ER membrane components. Therefore, we detected the lysosomal degradation of an ER component
using RG-Cytb5, which localizes RFP-GFP on the ER (Supplemental
Figure S3G). RG-Cytb5 was stably localized to the ER in HeLa cells,
and treatment with Torin 1, an inhibitor of mammalian target of rapamycin complex 1 (mTORC1), induced GFP-negative and RFPpositive dots (Figure 2, F and G). The expression of 6xIAPP-HA increased the number of GFP-negative and RFP-positive dots. Whiletunicamycin, an ER stressor, moderately increased the amount ofcleaved RFP, Torin 1 and 6xIAPP-HA induced marked accumulationof cleaved RFP (Figure 2, H and I), suggesting that 6xIAPP is eliminated by ER-phagy alongside ER components.

ANOVA and Sidak’s multiple comparison test, D. HeLa cells stably expressing Cas9 and sgRNA for FIP200 or nontarget(control) sgRNA were treated with Dox for 24 h to induce expression of 6xIAPP-RG before fixation. Cells were stainedwith antibodies against LAMP1 or LC3 and analyzed through immunofluorescence microscopy. Scale bars represent 10
and 1 µm (inset), E. (F–I) 6xIAPP induces lysosomal degradation of an ER protein. HeLa cells stably expressing RFP-GFPcytochrome b5 (RG-Cytb5) were incubated with Dox to induce 6xIAPP-HA-KDEL (6xIAPP-HA) expression, or with Torin
1, for 24 h. After fixation, cells were stained with antibodies against HA and analyzed through immunofluorescencemicroscopy. GFP-negative and RFP-positive signals (arrowhead) are indicated. Scale bars represent 10 and 1 µm (inset),
F. Quantification of the number of GFP negative, RFP positive puncta per cell. Solid bars indicate the median, boxes theinterquartile range (25th–75th percentile), and whiskers the 0th–100th percentile range. Data were collected from45 cells of each cell type. Differences were statistically analyzed by Dunnett’s multiple comparison test, G. HeLa cells
stably expressing RG-Cytb5 were treated with the indicated compounds or Dox for induction of 6xIAPP-HA expression
for 24 h. Cell lysates were analyzed through immunoblotting using antibodies against RFP, HA, or endogenous HSP90(loading control). # indicates degradative products of RG-Cytb5, H. The cleaved RFP/RG-Cytb5 band intensity ratio(normalized to the WT) is shown. Data represent the mean ± SE of three independent experiments. Differences were
statistically analyzed by Dunnett’s multiple comparison test, I.
Volume 34 April 1, 2023

CCPG1 ER-phagy| 5

FIGURE 3: CCPG1 is an essential ER-phagy receptor for the degradation of 6 × IAPP. (A, B) 6xIAPP-RG selectively
interacts with CCPG1 in mammalian cells. Tet-On HeLa cells (WT and FIP200-KO) expressing 1x (or 6x) IAPP-RG were
cultured with Dox for 48 h before IP. Inputs (5% of total) and immunoprecipitants (80% of total) were analyzed throughimmunoblotting using the indicated antibodies, A. The IP product/input ratio is shown. Data represent the mean ± SEof three independent experiments. Differences were statistically analyzed by Dunnett’s multiple comparison test,
B. (C) The expression of 6xIAPP promotes transcription of CCPG1. Tet-On HeLa cells highly expressing 6xIAPP-RG weretreated with each of the ER stressors for 16 h, or Dox to induce 6xIAPP-RG for 16–24 h. Quantitative real-time PCR was
performed for endogenous CCPG1 and GAPDH (loading control). Data represent the mean ± SE of three independent
6 | S. Ishii et al.

Molecular Biology of the Cell

CCPG1 is the essential ER-phagy receptor associated withthe degradation of 6 × IAPP
To investigate whether autophagic degradation of 6xIAPP requires ER-phagy receptor proteins, such as FAM134B, SEC62,
RTN3, CCPG1, and TEX264, immunoprecipitation (IP) of 1xIAPPRG and 6xIAPP-RG was performed. While no ER-phagy receptor
was precipitated in the presence of 1xIAPP-RG, 6xIAPP-RG coprecipitated with endogenous CCPG1 (Figure 3, A and B). We
observed that the amount of endogenous CCPG1, but not theamount of other ER-phagy receptors, was increased upon expression of 6xIAPP-RG (Figure 3A). The increase in CCPG1 wasfurther augmented by bafilomycin A1 treatment (Figure S4A).
Consistent with the previous finding that ER stressors (e.g., tunicamycin and thapsigargin) transcriptionally activate the expression of CCPG1 (Smith et al., 2018), 6xIAPP-RG increased themRNA abundance of CCPG1 (Figure 3C), suggesting that aggregated ER-luminal proteins induce CCPG1 expression. Additionally, FIP200-KO cells accumulated CCPG1, which also interacts
with 6xIAPP. This result is consistent with the finding that CCPG1KO cells, but not other ER-phagy receptor-KO cells, suppressed
the reduction in GFP fluorescence of 6xIAPP-RG, as measured
using flow cytometry (Figure 3D; Supplemental Figure S5, A and
B). The autophagic degradation of 6xIAPP-RG was restored byexogenous expression of HA-CCPG1 (Figure 3D), suggestingthat the KO phenotype is not an off-target effect of CRISPR. We
also showed that 6xIAPP formed large complexes (Supplemental
Figure S1A) and endogenous CCPG1 migrated to high-density
fractions, which have molecular mass similar to that of 6xIAPP(Supplemental Figure S5, C and D). These results indicate that
CCPG1 plays an essential role in ER-phagy-dependent degradation of 6xIAPP aggregates.
CCPG1 is a vertebrate-specific gene and regulates ER-phagy via
cytoplasmic N-terminal LIR and FIRs (Smith et al., 2018). Notably,
among ER-phagy receptors, only CCPG1 has a large ER luminal region, which consists of >500 aa (Kostenko et al., 2006). However, thephysiological function of this luminal domain remains unclear. We
hypothesized that the ER luminal region of CCPG1 is associatedwith the recognition of ER luminal cargos. The CCPG1Δ289–757
mutant lacking the whole ER luminal region (CCPG1 luminal deletion) failed to restore the RFP/GFP ratio of 6xIAPP in CCPG1-KO
cells (Figure 3D). CCPG1-KO cells showed fewer GFP-negative andRFP-positive dots for 6xIAPP-RG; these were recovered through reexpression of full-length CCPG1, but not CCPG1 luminal deletion(Figure 3E). We then performed an RFP cleavage assay, which demonstrated that reexpression of full-length CCPG1 restored lysosomal
degradation of 6xIAPP-RG (Figure 3, F and G). On the other hand,

reexpression of CCPG1 luminal deletion had only a moderate effecton CCPG1-KO, likely due to bulk ER-phagy via the cytoplasmic domain of CCPG1. These results indicate that the luminal region of
CCPG1 is required for ER-phagy-dependent lysosome degradationof 6xIAPP.
Overexpression of CCPG1 leads to the formation of punctate
structures on the ER, which reportedly colocalize with ATG proteinsin mammalian cells (Smith et al., 2018). Although the luminal-deletion mutant of CCPG1 colocalized with LC3, the rate of puncta formation was reduced (Supplemental Figure S5E). On the other hand,
the mutant of both LIR and FIRs, which was an interaction-deficientmutant for LC3 and FIP200, showed complete abolition of colocalization of CCPG1 with LC3, instead accumulating as large amorphous structures under ER stress conditions. However, no accumulation of CCPG1 was observed when both the luminal domain and
ATG protein-interaction motifs were deleted (Supplemental Figure
S5E). These data suggest that the localization of CCPG1 to autophagosomes is dependent on LIR and FIRs, but not the luminal domain.
In addition, luminal domain–-dependent formation of aggregatelike structures by CCPG1 may result from interactions with misfolded
proteins in the ER.

P3H4 is an endogenous substrate for CCPG1-dependent
ER-phagy
To identify endogenous substrates for CCPG1-dependent ERphagy, we searched for proteins that interacted with CCPG1 in aluminal domain–dependent manner. We performed IP using
FLAG-CCPG1 and its mutants, as described below, and subjected
the immunoprecipitates to mass spectrometry (MS) analysis. MS
screening 1 used FLAG-CCPG1 full-length and FLAG-CCPG1 luminal domain deletion as a negative control, and MS screening 2
used FLAG-CCPG1 cytosolic domain deletion (Figure 4A). The
resultant immunoprecipitants were analyzed through liquid chromatography with tandem mass spectrometry (LC-MS/MS; Figure
4B). We identified numerous ER luminal proteins, including ER
chaperones, ERAD components, glycosidases, and enzymes forcollagen maturation. Among these proteins, we focused on P3H4,
which was identified in both MS screens and has not yet beenstudied in the context of autophagy. P3H4 is an ER-resident protein belonging to the Leprecan (leucine proline-enriched proteoglycans) family (Ochs et al., 1996; Gruenwald et al., 2014) andforms a complex with prolyl 3-hydroxylase to regulate lysine hydroxylation of collagen (Heard et al., 2016). P3H4 exhibited
CCPG1-luminal domain-dependent interaction with CCPG1(Figure 4C). To monitor pulse-chased lysosomal degradation ofP3H4, we employed a HaloTag-based cleavage assay for P3H4

experiments. Differences were statistically analyzed by Dunnett’s multiple comparison test. (D–F) Lysosomal
degradation of 6xIAPP-RG occurs in a manner dependent on the ER-luminal region of CCPG1. WT and CCPG1-KO HeLa
cells expressing 6xIAPP-RG, with or without exogenous HA-CCPG1 (full-length or luminal deletion mutant), wereincubated in medium containing Dox and then subjected to flow cytometry, immunofluorescence (IF), andimmunoblotting. After 24 h of incubation in Dox-containing medium, the cells were incubated with Dox-free medium fora further 24 h. The fluorescence ratio was calculated for RFP-positive cells. Data represent the mean ± SE of three
independent experiments. Differences were analyzed using one-way ANOVA and Sidak’s multiple comparison test, D.
Cells treated with Dox as described were fixed and stained with antibodies against HA-tag and then analyzed using
immunofluorescence microscopy. GFP-negative and RFP-positive signals (arrowhead) are indicated. Scale bars represent
10 and 1 µm (inset), E. Lysates of each treatment were analyzed through immunoblotting using antibodies against RFP,
HA, and β-actin (loading control). # indicates degradative products of IAPP-RG, F. (G) The cells treated with Dox as
described above were analyzed through immunoblotting. The band intensities of cleaved RFP and 6xIAPP-RG were
quantified, and the ratio of cleaved RFP to 6xIAPP-RG (normalized to the WT) is shown. Data represent the mean ± SEof three independent experiments. Differences were analyzed using one-way ANOVA and Sidak’s multiple comparison
test.
Volume 34 April 1, 2023

CCPG1 ER-phagy| 7

FIGURE 4: P3H4 is an endogenous for CCPG1-dependent ER-phagy. (A) Strategy used to identify endogenous CCPG1
luminal domain–interacting proteins. (B) Results of differential interactome screening. Two independent IP and MS
analyses were conducted for each LC-MS/MS sample. (C) HEK293T cells transiently expressing ssHalo-P3H4 and WT, ormutated FLAG-CCPG1, were subjected to IP with anti-FLAG antibody and detection with anti-Halo and anti-FLAG
antibodies. (D) Immunoblotting of endogenous P3H4 in postnuclear supernatants of the indicated organs from
Atg5+/+;NSE-Atg5 (+/+) and Atg5−/−;NSE-Atg5 (KO) mice. (E, F) P3H4 accumulated in CCPG1-KO cells. WT or
CCPG1-KO cells with or without exogenous HA-CCPG1 were treated with DMSO or bafilomycin A1 for 24 h and thenlysed with lysis buffer. Each cell lysate was analyzed through immunoblotting using antibodies against P3H4, HA-tag, or
β-actin (loading control). * indicates nonspecific band, E. Band intensities were quantified and the ratio of P3H4 toβ-actin (normalized to the WT) is shown. Data represent the mean ± SE. Differences were analyzed using one-way
ANOVA and Sidak’s multiple comparison test, F.

(HaloTag becomes resistant to lysosomal degradation after ligand
binding; Yim et al., 2022). HeLa cells stably expressing ssHaloP3H4 were transfected with siRNA against CCPG1. After 3 d, thecells were exposed to tetramethylrhodamine-conjugated ligand
for 1 h and then incubated for 24 h. Generation of the Halo cleavage band was significantly impaired in CCPG1-depleted cells(Supplemental Figure S4, A and B). Cleavage of reporters was restored through reexpression of CCPG1, but not the luminal-domain deletion mutant or LIR/FIR mutants (Supplemental Figure S6,
A and B). These data suggest that P3H4 is degraded by CCPG1 ina luminal domain– and autophagy-dependent manner. Notably,

8 | S. Ishii et al.

endogenous P3H4 accumulated markedly in the pancreas, stomach, heart, and muscle of adult brain-rescued ATG5-KO mice (Yoshii et al., 2016; Figure 4D), suggesting that some tissues constitutively degrade P3H4 via ER-phagy. Considering that CCPG1 isalso highly expressed in exocrine tissues such as the pancreas andstomach (Chino et al., 2019), these results suggest that CCPG1
has a functional relationship with P3H4. P3H4 also accumulated in
CCPG1-KO or bafilomycin A1-treated HeLa cells (Figure 4, E and
F). Additionally, overexpression of HA-CCPG1 reduced theamount of P3H4. These data indicate that P3H4 is an endogenous
substrate for CCPG1-dependent ER-phagy.

Molecular Biology of the Cell

Highly conserved ER-luminal regions of CCPG1 containfunctional domains
As the ER-luminal region of CCPG1 is essential for ER-phagy-mediated degradation of 6xIAPP and P3H4, we apportioned the luminal
region into conserved regions through sequence alignment amongvertebrates. Although the length of the CCPG1 sequence variedamong species by up to 120 aa, the ER luminal region of CCPG1
contained four highly conserved regions (region A: aa 288–401, region B: aa 411–457, region C: aa 482–550, region D: aa 621–734;
Supplemental Figure S7A). We presumed that regions A–D of
CCPG1 include the cargo-interacting region (CIR) and generate Cterminal-truncated mutants (Δ742–757, Δ621–757, Δ551–757,
Δ411–757, and Δ289–757; Figure 5A). First, we measured bulk ERphagy activity using ssRFP-GFP-KDEL, an ER-luminal fluorescencereporter for ER-phagy (Chino et al., 2019), and found that overexpression of the full-length sequence or any truncated mutant in
CCPG1-KO cells promoted lysosomal degradation of ssRFP-GFPKDEL (Supplemental Figure S7, B and C), suggesting that the cytosolic region (containing LIR and FIRs) and transmembrane domain of
CCPG1 are sufficient for the induction of bulk ER-phagy. The cleavage of ssRFP-GFP-KDEL by some truncated mutants, including
Δ551–757 and Δ411–757, compared with full-length CCPG1 wasmore efficient, probably due to the high expression level. Fromthese results, enhancement of bulk-ER-phagy activity caused byoverexpression of the CCPG1 luminal-deletion mutant might resultin partial recovery of lysosomal degradation of 6xIAPP (Figure 3E).
In contrast, lysosomal degradation of 6xIAPP-RG was not rescued
through overexpression of the Δ411–757 and Δ289–757 mutants of
CCPG1 in CCPG1-KO cells (Figure 5, B–D), indicating that highly
conserved regions within 411–551 aa, including regions B and C,
might recognize 6xIAPP for degradation via ER-phagy. We defined
the highly conserved regions B–D as CIR 1–3 (Figure 5E).
To characterize these CIRs, we generated CIR1-, CIR2-, and dual
CIR1- and CIR2-truncated mutants and analyzed the associated lysosomal degradation of 6xIAPP-RG. Contrary to our expectations,lysosomal degradation of 6xIAPP-RG was rescued by all CIR1- andCIR2-related mutants (Figure 5, F–H). Therefore, we hypothesizedthat region D, defined as CIR3, may contribute to 6xIAPP degradation along with CIR1, CIR2, or both. We found that truncated CCPG1
with both CIR1 and CIR3 deleted was unable to restore the lysosomal degradation of 6xIAPP-RG (Figure 5, G and H). Alternatively,
bulk ER-phagy detected using ssRFP-GFP-KDEL was restored withoverexpression of any CIR-related mutants of CCPG1 (Supplemental Figure S7, C and D). These results demonstrate that the ER luminal region of CCPG1 contains functional domains, that is, CIRs, required for the selective degradation of 6xIAPP.
We conducted similar experiments using another cargo, P3H4.
CCPG1 mutants lacking CIR2 did not rescue the lysosomal degradation of RG-P3H4 (Figure 5, I and J). Furthermore, CCPG1ΔCIR2 nolonger colocalized to the dot-like structures observed for RG-P3H4,
in contrast to the full-length sequence and other mutants of CCPG1(Figure 5K). These findings suggest that P3H4 undergoes CIR2-dependent ER-phagy degradation and that CCPG1 contains multiple
cargo recognition sequences in the ER luminal region.

CIR1 interacts directly with IAPP, while CIR2 interacts
with P3H4
Next, we verified whether CCPG1 interacts directly with P3H4 and
6xIAPP. All recombinant proteins containing the full length of the ER
luminal region, each CIR region alone (CIRs 1–3), CIR1+3 of CCPG1,
P3H4, and 6xIAPP combined with small tags (ALFA, HA, and FLAG;
Götzke et al., 2019), and dihydrofolate reductase (DHFR) for stabiliVolume 34 April 1, 2023

zation (Iwakura et al., 1992) were synthesized in vitro using a cell-freeprotein synthesis system (Shimizu et al., 2001, 2005). Expression levels were estimated through immunoblotting, and the synthesized
proteins were mixed at a 1:3 volume ratio (antigen:binding partner)
in lysis buffer for IP using specific antibodies. 6xIAPP interacted efficiently with the entire C-terminal region (241–757) of CCPG1(Figure 6, A and B). Although HA-6xIAPP did not coprecipitate with
CIR2 and CIR3 alone, it did coprecipitate with CIR1 alone, and with
CIR1+CIR3. In contrast, the entire C-terminal region and CIR2 alone
coimmunoprecipitated by HA-P3H4 (Figure 6, C and D). These observations suggest that the CIRs of CCPG1 directly interact with ER
luminal cargos, and that different CIRs contribute to the recognitionof different cargos.

The ER luminal region of CCPG1 simultaneously recognizes
multiple cargos
As 6xIAPP and P3H4 interacted directly with different CIRs of
CCPG1, we hypothesized that a single molecule of CCPG1 could
interact simultaneously with 6xIAPP and P3H4. Therefore, we
checked for competitive inhibition using excess levels of the same
recombinant proteins with different tags synthesized using a cellfree protein translation system. The presence of excess FLAG-P3H4
inhibited co-IP of HA-P3H4 with ALFA-CCPG1 (full-length of C-terminal; Figure 7, A and B), while the addition of excess DHFR-FLAG
had no effect on that interaction. Similarly, co-IP of HA-6xIAPP by
ALFA-CCPG1 was inhibited by excess FLAG-6xIAPP (Figure 7, C
and D), indicating that interaction of the cargo with CCPG1 cancompete with the same excess cargos under in vitro conditions.
Then we confirmed competitive inhibition of 6xIAPP and P3H4
against CCPG1. Recombinant AFLA-CCPG1 was mixed with HAP3H4 or FLAG-6xIAPP or both, and then immunoprecipitated using
anti-ALFA antibody beads. The presence of P3H4 had no inhibitory
effect on binding between 6xAIPP and CCPG1; similarly, 6xAIPP did
not inhibit the binding of CCPG1 to P3H4 (Figure 7, E and F). This
result suggests that a single CCPG1 molecule interacts simultaneously with multiple cargos.

DISCUSSION
In this study, we found that CCPG1 directly interacts with ER luminal
proteins via CIRs, which recognize different types of cargo (Figure7G). CCPG1 is a receptor not only for LC3 and FIP200, but also for
ER luminal proteins (via its long ER luminal region). Therefore, ourfindings provide new insights, indicating that CCPG1 is a bispecificreceptor for ER luminal cargos and the autophagic membrane; it
drives selective ER-phagy and the cargo selectivity thereof, thereby
promoting efficient degradation of particular cargos throughER-phagy.
6xIAPP promotes primary nucleation and increases proteotoxicity (Kayatekin et al., 2018). Excessive expression of 6xIAPP in yeast
clogs the translocon, which is then removed through proteasomal
degradation via Ste24. Upon ZMPSTE24 inhibition by Lopinavir,
6xIAPP proteotoxicity also decreases cell viability in mammalianpancreatic cells (Kayatekin et al., 2018). We employed 6xIAPP fusedwith several different tags (fluorescent proteins or HA-tags) and the
KDEL signal as a model ER-phagy substrate. Moreover, we foundthat 6xIAPP is primarily translocated within the ER at a steady state
in mammalian cells, as verified by the analysis of N-glycosylation
using a construct fused with the glycosylation site from opsin (Supplemental Figure S1D). Aggregated 6xIAPP was primarily degradedthrough autophagy under the investigated conditions (Figure 2).
On the other hand, proteasome inhibition led to the accumulationof 6xIAPP, which was detected in the insoluble fraction without
CCPG1 ER-phagy| 9

FIGURE 5: The highly conserved C-terminal region of CCPG1 contains functional domains. (A) Diagram of the truncatedmutants of CCPG1. (B–D) The ER luminal region of CCPG1 at 411–757 aa is required for lysosomal degradation of
6xIAPP. CCPG1 KO HeLa cells coexpressing the indicated CCPG1 mutants (constitutive) and 6xIAPP-RG (Tet-On) wereincubated with medium containing Dox for 48 h and subjected to IF and immunoblotting. Cells were stained with
antibodies against HA-tag and analyzed through immunofluorescence microscopy. Scale bars represent 10 and 1 µm
(inset), B. Cell lysates were analyzed through immunoblotting with antibodies against RFP, HA-tag, and β-actin (loading
control). # indicates degradative products of 6xIAPP-RG, C. The band intensities of cleaved RFP and 6xIAPP-RG were
quantified and the ratio of cleaved RFP to 6xIAPP-RG (normalized to the WT) is shown. Data represent the mean ± SE ofthree independent experiments. Differences were analyzed using one-way ANOVA and Sidak’s multiple comparison
test, D. (E) Diagram of cargo-interacting region (CIR) mutants of CCPG1. (F–H) Both CIR1 and CIR3 of CCPG1 arerequired for lysosomal degradation of 6xIAPP. Tet-On WT HeLa cells and CCPG1 KO HeLa cells coexpressing theindicated CIR mutants of CCPG1 (stably) and 6xIAPP-RG (Tet-On) were cultured with Dox for 48 h and then subjected to
IF and immunoblotting. Cells were stained with antibodies against HA-tag and analyzed through immunofluorescencemicroscopy. Scale bars represent 10 and 1 µm (inset), F. Cell lysates were analyzed through immunoblotting using
antibodies against RFP, HA-tag, and β-actin (loading control). # indicates degradative products of 6xIAPP-RG. * indicates10 | S. Ishii et al.

Molecular Biology of the CellFIGURE 6: CIR1 and CIR3 interact directly with 6xIAPP, while CIR2 interacts with P3H4. (A, B) CIR1 of CCPG1 interacts
directly with 6xIAPP in vitro. DHFR-ALFA-tagged C-terminal truncated mutants of CCPG1 and DHFR-HA-6xIAPP
synthesized using a cell-free protein synthesis system were mixed with anti-HA antibody-conjugated Sepharose andincubated at 4°C for 1 h. After washing, eluates from the Sepharose were analyzed through immunoblotting with
antibodies against ALFA-tag and HA-tag, A. The band intensities of DHFR-tag and the luminal domains of CCPG1 were
quantified; the ratio of IP to input is shown. Data represent the mean ± SE of three independent experiments.
Differences were statistically analyzed by Dunnett’s multiple comparison test, B. (C, D) CIR2 of CCPG1 interacts directlywith P3H4 in vitro. DHFR-ALFA-tagged C-terminal truncated mutants of CCPG1 and DHFR-HA-P3H4 were synthesizedusing a cell-free protein synthesis system and then mixed with anti-HA antibody-conjugated Sepharose and incubated at
4°C for 1 h. After washing, eluates from the Sepharose were analyzed through immunoblotting with antibodies against
ALFA-tag and HA-tag, C. The band intensities of DHFR-tag and luminal domains of CCPG1 were quantified; the ratio of
IP to input is shown. Data represent the mean ± SE of three independent experiments. Differences were statisticallyanalyzed by Dunnett’s multiple comparison test, D.

nonspecific bands, G. Band intensities of cleaved RFP and 6xIAPP-RG were quantified and the ratio of cleaved RFP to
6xIAPP-RG (normalized to the WT) is shown. Data represent the mean ± SE of three independent experiments.
Differences were statistically analyzed by Dunnett’s multiple comparison test, H. (I–K) CIR2 of CCPG1 is required forlysosomal degradation of P3H4. Tet-On WT HeLa cells and CCPG1 KO HeLa cells coexpressing full-length or CIRmutant CCPG1 (constitutive) and ssRFP-GFP-P3H4 (RG-P3H4) (Tet-On) were cultured with Dox for 48 h and thensubjected to IF and immunoblotting. Cell lysates were analyzed through immunoblotting using antibodies against RFP,
HA-tag, and β-actin (loading control), I. The band intensities of cleaved RFP and RG-P3H4 were quantified, and the ratioof cleaved RFP to RG-P3H4 (normalized to the WT) is shown. Data represent the mean ± SE of three independent
experiments. Differences were analyzed using one-way ANOVA and Sidak’s multiple comparison test, J. Cells were
stained with antibodies against HA-tag and analyzed through immunofluorescence microscopy. GFP and RFP-positive
signals colocalizing with HA-positive structures (arrowhead) are indicated. Scale bars represent 10 and 1 µm (inset), K.
Volume 34 April 1, 2023

CCPG1 ER-phagy| 11

FIGURE 7: The ER luminal region of CCPG1 simultaneously recognizes multiple cargos. (A–D) The same ER luminal
cargo competitively inhibits the interaction with CCPG1. DHFR-ALFA-CCPG1 (C-terminal full-length) and DHFR-HAP3H4 (or 6xIAPP) with or without DHFR-FLAG-P3H4 (or 6xIAPP) or DHFR-FLAG were synthesized using a cell-freeprotein synthesis system, mixed with anti-ALFA Sepharose, and then incubated at 4°C for 1 h. FLAG-tagged products
were added at three times the level of HA-tagged cargos (indicated as × 3). Eluted products were analyzed throughimmunoblotting with antibodies against ALFA-, FLAG-, and HA-tag, A and C. The bar graph shows the ratio between
the indicated band intensities. Data represent the mean ± SE of three independent experiments. Differences were
statistically analyzed by Dunnett’s multiple comparison test (B and D. (E, F) Different ER luminal cargos are able tointeract with CCPG1 simultaneously. Synthesized DHFR-HA-P3H4 and DHFR-FLAG-6xIAPP were mixed with DHFRALFA-CCPG1 (C-terminal), either alone or in combination. Anti-ALFA beads were added to the mixture, which wasallowed to react at 4°C for 1 h. After washing four times with 1 × lysis buffer, SDS sample buffer was added and each
eluate was analyzed through immunoblotting using antibodies against ALFA-, FLAG-, and HA-tag, E. The bar graphshows the ratio of IP to input (n = 3). The data represent mean ± SE. Differences were statistically analyzed by Student’s
t test, F. (G) A simultaneous recognition model, in which CCPG1 interacts directly with multiple ER luminal cargos.

12 | S. Ishii et al.

Molecular Biology of the Cellglycosylation (Supplemental Figure S2H); this indicated that 6xIAPP
was mislocalized to the cytoplasm because of stress-induced inhibition of protein translocation (Kang et al., 2006) or was dislocatedfrom the clogged ER translocon by ZMPSTE24, a mammalian homologue of yeast Ste24 (Ast et al., 2016; Kayatekin et al., 2018). These
results demonstrate that 6xIAPP in mammals does not always clog
the translocon, but some proportion may be successfully translocated into the ER, where aggregated 6xIAPP is targeted for ERphagy. Analysis of cells expressing low and high levels of 6xIAPP
indicated a difference in dependence (Supplemental Figure S2, F
and G). Low expression levels of 6xIAPP were mainly degradedthrough a nonlysosomal pathway. On the other hand, highly expressed 6xIAPP was largely degraded through the lysosomal pathway (Supplemental Figure S2, F and G). We speculated that 6xIAPP
aggregated in the ER is an ERAD-inefficient substrate, similar to aggregated procollagen (Ishida et al., 2009). 6xIAPP formed large
oligomers under our conditions (Supplemental Figure S1, A and B).
The conformation of aggregated 6xIAPP at high concentrations in
the ER might lead to its recognition by CCPG1.
We observed that the overexpression of 6xIAPP alone is sufficient to induce autophagy (Figure 2B; Supplemental Figure S3, B-F),
likely because the increased abundance of CCPG1 is directly recognized by both FIP200 and LC3 (Figure 3A; Supplemental Figure S4,
A and B). This finding is supported by a previous report that overexpression of WT CCPG1, but not mutant LIR or FIR, promotes ERphagy (Smith et al., 2018). This raises the question of how 6xIAPP
induces CCPG1 expression. ER stress induces CCPG1 expression(Smith et al., 2018). Moreover, the UPR factors PERK, XBP1, andMIST1 are associated with CCPG1 transcription (Tian et al., 2010;
Adamson et al., 2016). However, we did not detect activation of the
UPR pathway by 6xIAPP overexpression in HeLa cells, although ERphagy was induced (unpublished data). Future work is needed to
identify the specific transcription factor(s) responsible for CCPG1
induction by 6xIAPP. Another open question is how CCPG1 induces
autophagy. We initially assumed that a luminal cargo transduces asignal from the ER lumen to the cytoplasmic side to induce ERphagy, similarly to the induction of the UPR; if this is the case, cargobinding to CCPG1 might trigger ER-phagy. However, overexpression of a CCPG1 mutant lacking the whole ER luminal region induced autophagy (Supplemental Figure S7, B and C). Therefore, theexpression of CCPG1, which recruits FIP200 and LC3 to the ER
membrane, might be sufficient to initiate ER-phagy, even without an
ER luminal cargo. Although this mechanism may not be elegant, it isvery straightforward, and CCPG1-dependent ER-phagy does not
require any additional signal transduction pathways.
As CCPG1 has the largest ER luminal domain among known ERphagy receptors, it has been speculated that CCPG1 recognizes ER
luminal proteins (Smith and Wilkinson, 2017; Grumati et al., 2018;
Smith et al., 2018; Wilkinson, 2020). We revealed that CCPG1 interacts directly with cargos, including 6xIAPP and P3H4. P3H4 was reported to play a role in catalyzing the complex hydroxylation process of collagen chains. Physiologically, P3H4 is a prognostic factor
for lung adenocarcinoma and bladder cancer (Li et al., 2018; Jinet al., 2021); it contributes to cancer invasion and growth (Hao et al.,2020), suggesting that CCPG1 may have a protective effect against
these cancers. In addition, endogenous P3H4 accumulated in thepancreas and stomach of autophagy-deficient mice (Figure 4D),
suggested that it undergoes degradation dependent on the autophagy–lysosome pathway, particularly in exocrine organs.
Several ER chaperones, including BiP, calnexin, PDIA1, PDIA3,CALR, and TXNDC5, were identified by our MS analysis of proteinsbinding to the luminal region of CCPG1 (Figure 4, A and B). This is
Volume 34 April 1, 2023

consistent with a previous finding that ER chaperones, including
PDIA6, accumulated in the pancreas of CCPG1 hypomorphic mice
(Smith et al., 2018). As FAM134B has been reported to bind to BiP
and calnexin, which mediates the capture of luminal misfolded proteins under ER stress conditions (Fregno et al., 2018; Forrester et al.,2019; Chipurupalli et al., 2022), CCPG1 may employ a similar mechanism, in which ER chaperones mediate the degradation of luminal
proteins through ER-phagy. CCPG1 is an ER stress–responsive gene,suggesting that ER-phagy may share common substrates with
ERAD, and that cross-talk may exist, depending on the amount ofdegradation. In this scenario, although ER-phagy exhibits more extensive selective recognition of unfolded proteins, its substrate selectivity might be similar to the UPR, such that direct recognition of
a substrate by the luminal region of CCPG1 may contribute to ERphagy-specific selectivity.
Homology searches were performed with the Position-Specific
Iterative Basic Local Alignment Search Tool (PSI-BLAST) using human sequences of CIRs from CCPG1; however, we detected no proteins containing similar domains. Moreover, further searches using
the InterPro database (v90.0; Blum et al., 2021) identified no paralogs or proteins with conserved CIR-related regions. In addition, no
similarity among CIRs in CCPG1 was found through sequence alignment. The AlphaFold tool predicted that CIRs 1–3 have different
structures (Jumper et al., 2021; Varadi et al., 2022), and our data
indicated different cargo selectively among these CIRs; this indicated that each CIR recognizes the cargo using different machinery.
Therefore, we speculated that the ER luminal region of CCPG1 has
unique features related to the recognition of ER-phagy substrates.
Future work is needed to identify the specific motifs that bind ERphagy substrates and reveal the detailed mechanisms of substrateselectivity.

MATERIALS AND METHODS
Request a protocol through Bio-protocol.

Antibodies
Rabbit polyclonal anti-LAMP1 antibodies were a gift from Y. Tanaka(Kyusyu University). Rabbit polyclonal anti-TRAPα antibodies were agift from R.S. Hegde (MRC LMB). Mouse monoclonal anti-HA (clone
16B12, 901502) was purchased from BioLegend Funakoshi (Tokyo,
Japan). Mouse monoclonal anti-RFP (M204-3) and anti-LC3 (M152-3)
antibodies and rabbit polyclonal anti-LC3 (PM036), anti-p62(PM045), and anti-ATG9A (PD042MS) antibodies were purchasedfrom MBL (Tokyo, Japan). Mouse monoclonal anti-β-actin (28198721), anti-GFP (clone mFX75, 012-22541), and anti-FLAG (01822381) antibodies were purchased from Wako (Osaka, Japan).
Mouse monoclonal anti-RPL19 (WH0006143M1), anti-β-actin(A2228), and anti-FLAG (F4042) antibodies were purchased from
Sigma-Aldrich (St. Louis, MO, USA). Mouse monoclonal anti-HSP90(610419) antibodies were purchased from BD Biosciences (Tokyo,
Japan). Mouse monoclonal anti-Halo (G9211) antibody was purchased from Promega (Madison, WI). Rabbit polyclonal antiFAM134B (21537-1-AP), anti-RTN3 (12055-2-AP), anti-CCPG1(13861-1-AP), anti-TEX264 (25858-1-AP), anti-RB1CC1 (FIP200)(17250-1-AP), anti-ATG5 (10181-2-AP), and anti-Histone-H3 (171681-AP) antibodies were purchased from Proteintech (Tokyo, Japan).
Rabbit polyclonal anti-SEC62 (NBP1-84045) antibodies were purchased from Novus Biologicals (Centennial, CO). Rabbit polyclonalanti-P3H4 (A13754) antibodies were purchased from ABclonal(Woburn, MA). Rabbit polyclonal anti-ALFA antibodies were raised
in rabbits through immunization with the ALFA peptide (Eurofins).
GFP-nanobody and ALFA tag-nanobody Sepharose were generated
CCPG1 ER-phagy| 13through conjugation of GFP-nanobody protein purified from pOPINE GFP nanobody (plasmid 49172; Addgene, Watertown, MA) and
ALFA tag-nanobody protein (Götzke et al., 2019) purified from pRSET-A ALFA–His to N-hydroxy succinimide-activated Sepharose 4
Fast Flow (GE Healthcare, Chicago, IL). Mouse monoclonal antiFLAG M2 affinity gel (A2220) and anti-FLAG M2 magnetic beads(M8823) were purchased from Sigma-Aldrich. Mouse monoclonalanti-HA conjugated Sepharose (014-23081) was purchased fromWako.

Plasmids
To generate pCW_1x (or 6x) IAPP-RFP-GFP-KDEL or 1x (or 6x) IAPPRFP-GFP, IAPP-related constructs, human IAPP and 6xIAPP sequences (synthesized using gBlocks Gene Fragments; Integrated
DNA Technologies; Coralville, IA) were inserted into the pCW57.1
vector (plasmid #41393; Addgene) along with DNA encoding themCherry, super-folder GFP (sfGFP), and, if indicated, KDEL sequences. For pCW_1x (or 6x) IAPP-HA-KDEL, each IAPP sequenceand 3 × hemagglutinin (HA)-tag was used. pCW_1x (or 6x) IAPP-GlyHA-KDEL was generated through insertion of the N-glycosylationsite from opsin (Pedrazzini et al., 2000) into pCW_1x (or 6x) IAPP-HAKDEL. To generate pCW ssRFP-GFP-KDEL, the signal sequence of
prolactin and the mCherry, super-folder GFP, and KDEL sequenceswere subcloned into the pCW57.1 vector. For pCW ssRFP-GFPP3H4, cDNA of human P3H4 amplified from total cDNA of HEK293T
cells was used. To generate full-length or truncated CCPG1 constructs, cDNA of CCPG1s was amplified from HEK293FT total cDNA
and inserted into the pLenti cytomegalovirus vector GFP Puro (plasmid 17448; Addgene) or pMRX-IB (Morita et al., 2018); these plasmids were generated from pMXs (Kitamura et al., 2003) along withenhanced GFP, 3xFLAG-tag, or 3xHA-tag. Truncated constructs wereprepared through PCR-mediated site-directed mutagenesis. To generate pMRXIB_RFP-GFP-LC3 and pMRXIB_RFP-GFP-Cytb5, cDNAof rat LC3 and the transmembrane domain of Cytb5 were amplified
from pMXs-IP-EGFP-LC3 (plasmid 38195; Addgene) and pMXs-puro
GFP-b5 ER (plasmid 38274; Addgene) and inserted into the pMRXIB vector. For the ATG- or ER-phagy receptor gene-targeted CRISPR
vector, the specific sgRNA sequences were inserted into lentiCRISPR
v2 hygro (plasmid 98291; Addgene) or lentiGuide-puro (plasmid52963; Addgene). For plasmids with a cell-free protein synthesis system in vitro, DHFR (amplified from PURExpress DHFR), 3xHA-tag,
3xFLAG-tag, ALFA-tag, P3H4, and the indicated regions of CCPG1(amplified from total cDNA of HEK293FT) were inserted into a PURExpress plasmid (provided with the PURE system kit from New England Biolabs, Ipswich, MA). pCMV-VSVG (plasmid 8454; Addgene)and psPAX2 (plasmid 12260; Addgene) were used for lentivirus production. pCMV-VSVG and Gag were used for retrovirus production.

Cell culture
HeLa and HEK293FT cells were cultured in DMEM (Nacalai Tesque,
Kyoto, Japan) supplemented with 10% fetal bovine serum (FBS; MP
Biomedicals, Santa Ana, CA) and 50 mg/ml penicillin and streptomycin (regular medium) in a 5% CO2 incubator. Tetracycline-On (TetOn) cells were generated through lentiviral transduction with apCW57.1 vector containing the single-vector Tet-On component.
For compound treatment, cells were incubated at the indicatedtimes with 0.2 µM bafilomycin A1 (LC Laboratories, Woburn, MA), 1µM Torin 1 (Tocris Bioscience, Ellisville, MO), 2 µg/ml tunicamycin(Sigma-Aldrich), 0.2 µM thapsigargin (Santa Cruz Biotechnology,
Dallas, TX), 10 µg/ml E64d (Peptide Institute, Osaka, Japan), 100µM pepstatin A (Peptide Institute), 20 µg/ml leupeptin (Peptide Institute), or 1 µg/ml Dox (Clontech, Mountain View, CA).
14 | S. Ishii et al.

Generation of a KO cell line using CRISPR/Cas9 gene
editing (mixed population of WT and KO cells)
SgRNA sequences for KO cells were designed using CHOPCHOP[FIP200 (5′-GGCTGCAATCATGGCCAACC-3′): Atg5 (5′-AAG­
AGTAAGTTATTTGACGT-3′): Atg9A (5′-AGGATATTCGA­GAGAA­GAAG-3′): CCPG1(5′-ACAGTGATTCATCTTGTGGT-3′): FAM134B(5′-ACTCTTTGGCAGCAACCGTG-3′):
RTN3(5′-AAGAAGA­CTGGGTTTGTCTT-3′):
SEC62(5′-CCACCAATATGATGGGTCAC-3′): TEX264 (5′-GCTACTACTGGGCCTGATTG-3′)] and cloned
into lentiCRISPR v2 hygro (plasmid 98291; Addgene) or lentiGuidepuro (plasmid 52963; Addgene). HeLa cells were infected with thelentivirus and then cultured for 7 d under hygromycin/puromycin
selection. The hygromycin/puromycin-resistant cells were used as
KO cell lines.

Generation of a clonal KO cell line using CRISPR/Cas9
gene editing
To generate clonal HeLa CCPG1 KO cells, HeLa cells were transfected with pSpCas9(BB)-2A-Puro (PX459) V2.0 (plasmid 62988;
Addgene) containing the sgRNA sequence of CCPG1. Puromycin
was transiently applied for 24 h posttransfection. After culturing for>5 d, single cell clones were isolated through dilution into 96-wellplates and screened for successful KO of CCPG1 throughimmunoblotting.

Generation of stable cell lines through lentiviral andretroviral infection
Stable cell lines were generated using lentiviral and retroviral expression systems. HEK293FT cells were transiently cotransfected withlentiviral or retroviral vectors using PEI MAX regent (Polysciences,
Warrington, PA). After culturing for 72 h, the growth medium containing the virus was centrifuged, and the resulting supernatant wascollected. HeLa cells were incubated with this virus-containing medium for 48 h and then selected with 1 µg/ml puromycin (InvivoGen,
San Diego, CA), 5 µg/ml blasticidin S (Wako), 100 µg/ml hygromycin
B (Wako), or 500 µg/ml G418 (Nacalai Tesque).

Immunoblotting
Cells were washed with cold phosphate-buffered saline (PBS) andlysed in lysis buffer (1% Triton X-100, 50 mM Tris-HCl pH 7.5, 1 mM
EDTA [EDTA], and 150 mM NaCl) supplemented with protease inhibitor cocktail (EDTA-free; Nacalai Tesque) and 1 mM phenylmethanesulfonyl fluoride for the collection of soluble fractions, or 1xSDS
sample buffer (0.0625 mM Tris-HCl pH 6.8, 2% SDS,10% 2-mercaptoethanol, and 10% glycerol) for the collection of whole-cell lysates,for 15 min at 4°C. The soluble lysates were clarified through centrifugation at 20,630 × g for 5 min, and 6 × sodium dodecyl sulfate (SDS)
sample buffer was added. The samples were heated to 95°C for
5 min before SDS/polyacrylamide gel electrophoresis (SDS/PAGE),
in which 20 µg of protein per lane was separated and then transferred to a polyvinylidene disulfide membrane (Millipore, Burlington, MA). Immunoblot analysis was performed with the indicatedantibodies suspended in Signal Enhancer Hikari (Nacalai Tesque)and the immunoreactive proteins were visualized using ImmunoStar
Zeta (Wako).

siRNA knockdown experiments
Stealth RNAi oligonucleotides were purchased from Thermo Fisher
Scientific (Waltham, MA). The following sequences were used: siCCPG1, 5′-UUCCAAUAUAGAUACUGUCUUCGGG -3′ and siLuciferase (siLuc), 5′-AAUUAAGUCCGCUUCUAAGGUUUCC-3′. The
stealth RNAi oligonucleotides were transfected into cells using
Molecular Biology of the Cell

Lipofectamine RNAiMAX (13778150; Thermo Fisher Scientific) according to the manufacturer’s instructions.

Mice
All animal experiments were approved by the Institutional Animal
Care and Use Committee of the University of Tokyo. Atg5−/−;NSE-Atg5
mice (Yoshii et al., 2016) and Fip200flox/flox;Nestin-Cre mice (Lianget al., 2010) have been described previously. To obtain postnuclear
supernatants, tissue homogenates were centrifuged at 500 × g for
10 min and the supernatants were boiled in sample buffer.

LC-MS/MS analysis of FLAG-CCPG1 immunoprecipitates
HeLa stably expressing FLAG-CCPG1, as well as the FLAG-CCPG1
luminal-deletion mutant and FLAG and FLAG-CCPG1 cytosol-deletion mutants, were incubated and lysed with lysis buffer (50 mM
Tris-HCl, pH 7.5, 150 mM NaCl, 1% NP-40, and complete EDTA-freeprotease inhibitor [03969-21; Nacalai Tesque]). After centrifugation
at 17,700 × g for 10 min, the supernatants were incubated with antiFLAG M2 magnetic beads for 3 h at 4°C with gentle rotation. The
eluted proteins were enzymatically digested according to a phasetransfer surfactant (PTS) protocol (Rappsilber et al., 2007). Then 50 µlof each eluted sample was mixed with 85 µl PTS buffer. Samples
were reduced with 10 mM dithiothreitol at room temperature for
30 min and alkylated with 50 mM 2-iodoacetamide (804744; SigmaAldrich) at room temperature for 30 min. Next, samples were diluted
fivefold through addition of 50 mM NH4HCO3 solution followed bydigestion with 1 µg of lysyl endopeptidase (LysC; 121-05063; Wako)
at 37°C for 4 h. Samples were further digested with 1 µg trypsin at
37°C for 8 h. An equal volume of ethyl acetate acidified with 0.5%
trifluoroacetic acid (TFA) was added to the digested samples. Aftercentrifugation twice at 10,000 × g for 10 min at room temperature,
the aqueous phase containing the peptides was collected and driedusing a SpeedVac concentrator (Thermo Fisher Scientific). The dried
peptides were solubilized in 100 µl of 2% acetonitrile and 0.1% TFA,
and the peptide mixture was trapped on a handmade C18 STAGE
tip prepared as described previously (Boersema et al., 2009). The
trapped peptides were subjected to a previously reported dimethyllabeling procedure (Boersema et al., 2009). Subsequently, CH2O
and NaBH3CN (light label) were added to the FLAG-only sample.
Similarly, CD2O and NaBH3CN (heavy label) were added to the
FLAG-NEK9 sample. The dimethyl-labeled peptides remaining on
the tip were eluted with 100 µl of 80% acetonitrile and 0.1% TFA.
The light- and heavy-label eluates were mixed and dried using a
SpeedVac concentrator. Each sample was dissolved in 2% acetonitrile and 0.1% TFA and loaded into the LC-MS system with an Orbitrap Exploris 480 MS instrument (Thermo Fisher Scientific) equippedwith a nano-high-performance liquid chromatography system (Advance UHPLC; Bruker Daltonics, Billerica, MA, USA) and HTC-Pal
autosampler (CTC Analytics, Zwingen, Switzerland) with a trap
column (0.3 × 5 mm, L-column ODS; Chemicals Evaluation andResearch Institute, Tokyo, Japan). Samples were separated using a
gradient of mobile phases A (0.1% [v/v] formic acid in H2O) and B(0.1% [v/v] formic acid in acetonitrile) at a flow rate of 300 nL/min(4–32% B for 190 min, 32–95% B for 1 min, 95% B for 2 min, 95% to
4% B for 1 min, and 4% B for 6 min) with a homemade capillarycolumn (length 200 mm and inner diameter 100 µm) packed with
2-µm C18 resin (L-column2; Chemicals Evaluation and Research Institute). Then the eluted peptides were electrosprayed (2.1 kV) andintroduced into the MS instrument. Data were obtained in positive
ion mode for data-dependent MS/MS (ddMS2) acquisition. Full MS
spectra were obtained across a scan range of 350–1800 m/z with
60,000 full width at half maximum (FWHM) resolution at 200 m/z.
Volume 34 April 1, 2023

MS2 spectra were obtained with 7500 FWHM resolution at 200 m/z.
For ddMS2 acquisition, full MS spectra were obtained every 3 s andMS2 spectra were obtained during the 3 s intervals. The most abundant precursor ions (excluding isotopes of a cluster) above the 5.0 ×
103 intensity threshold with a charge state from 2+ to 7+ were selected using a 2.0-m/z isolation window. A 20-s dynamic exclusion
period was applied. The raw data obtained were used for a database search (UniProt reviewed the mouse database on September
13, 2018) with the Sequest HT algorithm running on the Proteome
Discoverer 2.5 platform (Thermo Fisher Scientific). The parameters
for database searches were as follows: peptide cleavage was set totrypsin; missed cleavage sites were allowed for up to two residues;
peptide lengths were set to 6–144 aa; and mass tolerances were set
to 10 ppm for precursor ions and 0.02 Da for fragment ions. Carbamidomethylation of cysteine and dimethylation [H(4)C(2) or D(4)C(2)] of lysin and the peptide N-terminus were set as fixed modifications. Oxidation of methionine was set as a variable modification. A
significance threshold of p < 0.05 was applied. The abundances ofprecursor ions were calculated based on the area of the precursors,
determined with Proteome Discoverer 2.5.

Insoluble fraction assay
Cells were lysed in lysis buffer with protease inhibitor cocktail and
1 mM phenylmethanesulfonyl fluoride for 15 min at 4°C. The lysate
was clarified through centrifugation at 20,630 × g for 30 min. Thenthe supernatant was collected as the soluble fraction, and the pellet(containing the insoluble fraction) was washed three times with fresh
lysis buffer. The insoluble pellet was suspended in a volume of lysis
buffer equal to the soluble fraction volume and then mixed with 6 ×
SDS sample buffer. The samples were heated to 95°C for 5 min andthen sonicated.

Glycosylation assay
Cells expressing IAPP constructs fused with the glycosylation site of
opsin (Pedrazzini et al., 2000) were lysed in 1 × lysis buffer without
protease inhibitor, incubated for 15 min on ice, and then centrifuged at 20,630 × g for 5 min. The supernatant of each lysate was
mixed with 10 × glycoprotein denaturing buffer, heated for 5 min at
95°C, and then incubated with endoglycosidase H or H2O at 37°C
for 10 h. Next, 6 × SDS sample buffer was added to each product;
the mixture was heated to 95°C for 5 min and then analyzed throughSDS–PAGE and immunoblotting.

Sucrose gradient fractionation
Cells were lysed in CHAPS (3-[(3-cholamidopropyl)dimethylammonio]1-propanesulfonate) buffer containing protease inhibitor for 30 min
on ice. Then the lysates were centrifuged at 20,630 × g for 30 min at
4°C to remove insoluble components; the resulting supernatants
were layered on top of a 10–50% discontinuous sucrose gradientand centrifuged at 259,000 × g for 90 min at 4°C (CS 150FNX; rotor:
S55S [9124280K]; Hitachi, Tokyo, Japan). Eleven 200-µl fractions
were collected and mixed with 6 × SDS sample buffer and then analyzed through SDS–PAGE and immunoblotting.

Immunofluorescence microscopy
Cells were plated on coverslips, fixed in 3.7% formaldehyde in PBS
for 15 min, permeabilized with 50 µg/ml digitonin or 0.1% Triton
X-100 in PBS for 5 min, and then blocked with 10% newborn bovine
serum (NBS) in PBS for 45 min. After blocking, each sample was incubated with the indicated primary antibodies for 1 h. After washing,each sample was incubated with Alexa-647 conjugated anti-mouseor anti-rabbit IgG secondary antibodies (Thermo Fisher Scientific) for
CCPG1 ER-phagy| 15

1 h. The stained cells were observed under a confocal laser microscope (FV1000 IX81; Olympus, Tokyo, Japan) using a 100× oil immersion objective lens with a numerical aperture of 1.40. Images
were acquired using FV10-ASW 2.1 imaging software.

Flow cytometry
Cells were trypsinized with EDTA and recovered through detachment from the dish. The cells were passed through a 70-µm cellstrainer and then resuspended in 10% FBS and 1 µg/ml 4′,6-diamidino-2-phenylindole (DAPI) in PBS for flow cytometric analysis using a
CytoFLEX S flow cytometer equipped with NUV 375-nm (DAPI),488-nm (GFP), and 561-nm (RFP) lasers (Beckman Coulter, Brea, CA,
USA). Dead cells were detected through DAPI staining. In each sample, ten thousand cells were acquired and the RFP/GFP fluorescence ratio was calculated as red fluorescence intensity divided bygreen fluorescence intensity in RFP-positive cells. The data were
processed with Kaluza software (Beckman Coulter).

Immunoprecipitation
Cells were lysed in lysis buffer with protease inhibitor cocktail and 1
mM phenylmethanesulfonyl fluoride for 15 min at 4°C. The lysate
was clarified through centrifugation at 20,630 × g for 5 min, and theresulting supernatant was collected. A portion of this supernatant
was mixed with 6 × SDS buffer, heated to 95°C for 5 min, and thenused as an input. GFP-nanobody Sepharose beads were added tothe mixture, which was incubated for 2 h at 4°C. The Sepharose
beads were washed four times with lysis buffer before elution with
SDS sample buffer. The samples were subsequently separated
through SDS–PAGE and analyzed through immunoblotting.

In vitro protein biding assay
Each DHFR- and small-tag-fusion protein was synthesized using
PURE frex2.1 and suspended in 1 × lysis buffer containing 1% Triton
X-100. Substrates for IP and target proteins were mixed at a ratio of
1:3, and the Sepharose beads conjugated to specific antibodies or
nanobodies against HA, FLAG, or ALFA were incubated at 4°C for
1 h. The beads were washed four times with 1 × lysis buffer and 2 ×
SDS sample buffer was added. The mixture was then heated to
95°C for 5 min and analyzed using SDS–PAGE and immunoblotting.
For analysis of competition for the same cargos, FLAG-tagged
products were added at three times the volume of HA-tagged cargos. For analysis of competition for different cargos, P3H4 and
6xIAPP were mixed at equal volumes. SDS–PAGE and immunoblotting were performed for analysis.

RNA extraction, reverse transcription, and quantitative
real-time PCR
Total RNA was extracted from cells using ISOGEN II (NIPPON
GENE, Tokyo, Japan). Reverse transcription was performed using
ReverTra Ace reverse transcription reagents (TOYOBO LIFESCIENCE, Osaka, Japan). The following gene-specific primer
sequences were used: human CCPG1, 5′-TTCTGTGACCCCCACTGACA-3′ (forward) and 5′-TTGGCTGCTTTCTCCTTGCT-3′ (reverse);
human GAPDH, 5′-CCACATCGCTCAGACACCA-3′ (forward)
and 5′-GGCAACAATATCCACTTTACCAGAG-3′ (reverse). Relative
quantification of gene expression was performed according to the2 (– ΔΔCT) method. The housekeeping gene GAPDH was used as aninternal control to normalize the variability in expression levels.

ACKNOWLEDGMENTS
We thank Yoshitaka Tanaka (Kyushu University) for the anti-LAMP1
anti-bodies; Takeshi Murata (Chiba University) for cell sorting
16 | S. Ishii et al.

support; and Keita Chagi, Eigo Takeda, Yutaro Hama, and membersof the Matsuura lab for valuable discussions. This work was supported by JSPS KAKENHI (Grants 19K22413, 20H03249, 22H04634
to E.I., 22J12812 to S.I., 22H04919 to N.M., 18H05270 to H.R.U.),and the JST FOREST Program (Grant JPMJFR204N to E.I.), JSTSPRING (Grant JPMJSP2109 to S.I.), JST ERATO (Grants JPMJER1702 to N.M., JPMJER1904 to H.R.U.), Human Frontier Science
Program (HFSP) Research Grant Program (Grant RGP0019/2018 to
H.R.U.), and Takeda Science Foundation (to E.I.).

REFERENCES

Adamson B, Norman TM, Jost M, Cho MY, Nuñez JK, Chen Y, Villalta JE,
Gilbert LA, Horlbeck MA, Hein MY, et al. (2016). A multiplexed singlecell CRISPR screening platform enables systematic dissection of theunfolded protein response. Cell 167, 1867–1882.e1821.
An H, Ordureau A, Paulo JA, Shoemaker CJ, Denic V, Harper JW (2019).
TEX264 is an endoplasmic reticulum-resident ATG8-interacting protein critical for ER remodeling during nutrient stress. Mol Cell 74, 891–908.e810.
Ast T, Michaelis S, Schuldiner M (2016). The protease Ste24 clears clogged
translocons. Cell 164, 103–114.
Bernales S, McDonald KL, Walter P (2006). Autophagy counterbalances
endoplasmic reticulum expansion during the unfolded protein response.
PLoS Biol 4, e423.
Bhaskara RM, Grumati P, Garcia-Pardo J, Kalayil S, Covarrubias-Pinto A,
Chen W, Kudryashev M, Dikic I, Hummer G (2019). Curvature induction
and membrane remodeling by FAM134B reticulon homology domain
assist selective ER-phagy. Nat Commun 10, 2370.
Blum M, Chang HY, Chuguransky S, Grego T, Kandasaamy S, Mitchell A,
Nuka G, Paysan-Lafosse T, Qureshi M, Raj S, et al. (2021). The InterProprotein families and domains database: 20 years on. Nucleic Acids Res
49, D344–D354.
Boersema PJ, Raijmakers R, Lemeer S, Mohammed S, Heck AJ (2009).
Multiplex peptide stable isotope dimethyl labeling for quantitative
proteomics. Nat Protoc 4, 484–494.
Bukau B, Weissman J, Horwich A (2006). Molecular chaperones and protein
quality control. Cell 125, 443–451.
Butler AE, Janson J, Bonner-Weir S, Ritzel R, Rizza RA, Butler PC (2003).
Beta-cell deficit and increased beta-cell apoptosis in humans with type 2
diabetes. Diabetes 52, 102–110.
Callea F, Brisigotti M, Fabbretti G, Bonino F, Desmet VJ (1992). Hepatic
endoplasmic reticulum storage diseases. Liver 12, 357–362.
Chakrabarti A, Chen AW, Varner JD (2011). A review of the mammalian
unfolded protein response. Biotechnol Bioeng 108, 2777–2793.
Chen Q, Xiao Y, Chai P, Zheng P, Teng J, Chen J (2019). ATL3 is a tubular
ER-phagy receptor for GABARAP-mediated selective autophagy. Curr
Biol 29, 846–855.e846.
Chino H, Hatta T, Natsume T, Mizushima N (2019). Intrinsically disordered
protein TEX264 mediates ER-phagy. Mol Cell 74, 909–921.e906.
Chino H, Yamasaki A, Ode KL, Ueda HR, Noda NN, Mizushima N (2022).
Phosphorylation by casein kinase 2 enhances the interaction between
ER-phagy receptor TEX264 and ATG8 proteins. EMBO Rep, e54801.
Chipurupalli S, Ganesan R, Martini G, Mele L, Reggio A, Esposito M,
Kannan E, Namasivayam V, Grumati P, Desiderio V, Robinson N (2022).
Cancer cells adapt FAM134B/BiP mediated ER-phagy to survive hypoxic
stress. Cell Death Dis 13, 357.
De Duve C (1963). The lysosome. Sci Am 208, 64–72.
De Leonibus C, Cinque L, Settembre C (2019). Emerging lysosomal
pathways for quality control at the endoplasmic reticulum. FEBS Lett
593, 2319–2329.
Ellgaard L, Helenius A (2003). Quality control in the endoplasmic reticulum.
Nat Rev Mol Cell Biol 4, 181–191.
Forrester A, De Leonibus C, Grumati P, Fasana E, Piemontese M, Staiano
L, Fregno I, Raimondi A, Marazza A, Bruno G, et al. (2019). A selective
ER-phagy exerts procollagen quality control via a calnexin-FAM134B
complex. EMBO J 38, e99847.
Fregno I, Fasana E, Bergmann TJ, Raimondi A, Loi M, Soldà T, Galli C,
D’Antuono R, Morone D, Danieli A, et al. (2018). ER-to-lysosomeassociated degradation of proteasome-resistant ATZ polymers occurs
via receptor-mediated vesicular transport. EMBO J 37, e99259.
Fumagalli F, Noack J, Bergmann TJ, Cebollero E, Pisoni GB, Fasana E,
Fregno I, Galli C, Loi M, Soldà T, et al. (2016). Translocon component
Sec62 acts in endoplasmic reticulum turnover during stress recovery.
Nat Cell Biol 18, 1173–1184.
Molecular Biology of the Cell

Götzke H, Kilisch M, Martínez-Carranza M, Sograte-Idrissi S, Rajavel A,
Schlichthaerle T, Engels N, Jungmann R, Stenmark P, Opazo F, Frey
S (2019). The ALFA-tag is a highly versatile tool for nanobody-based
bioscience applications. Nat Commun 10, 4403.
Gruenwald K, Castagnola P, Besio R, Dimori M, Chen Y, Akel NS, Swain FL,
Skinner RA, Eyre DR, Gaddy D, et al. (2014). Sc65 is a novel endoplasmic reticulum protein that regulates bone mass homeostasis. J Bone
Miner Res 29, 666–675.
Grumati P, Dikic I, Stolz A (2018). ER-phagy at a glance. J Cell Sci 131.
Grumati P, Morozzi G, Hölper S, Mari M, Harwardt MI, Yan R, Müller S,
Reggiori F, Heilemann M, Dikic I (2017). Full length RTN3 regulates
turnover of tubular endoplasmic reticulum via selective autophagy.
eLife 6, e25555.
Hao L, Pang K, Pang H, Zhang J, Zhang Z, He H, Zhou R, Shi Z, Han C(2020). Knockdown of P3H4 inhibits proliferation and invasion of
bladder cancer. Aging (Albany NY) 12, 2156–2168.
Hara T, Takamura A, Kishi C, Iemura S, Natsume T, Guan JL, Mizushima N(2008). FIP200, a ULK-interacting protein, is required for autophagosome formation in mammalian cells. J Cell Biol 181, 497–510.
Heard ME, Besio R, Weis M, Rai J, Hudson DM, Dimori M, Zimmerman
SM, Kamykowski JA, Hogue WR, Swain FL, et al. (2016). Sc65-null miceprovide evidence for a novel endoplasmic reticulum complex regulating
collagen lysyl hydroxylation. PLoS Genet 12, e1006002.
Hebert DN, Molinari M (2007). In and out of the ER: protein folding, quality control, degradation, and related human diseases. Physiol Rev 87,
1377–1408.
Hetz C, Zhang K, Kaufman RJ (2020). Mechanisms, regulation and functionsof the unfolded protein response. Nat Rev Mol Cell Biol 21, 421–438.
Houck SA, Ren HY, Madden VJ, Bonner JN, Conlin MP, Janovick JA, Conn
PM, Cyr DM (2014). Quality control autophagy degrades soluble ERADresistant conformers of the misfolded membrane protein GnRHR. Mol
Cell 54, 166–179.
Ishida Y, Yamamoto A, Kitamura A, Lamandé SR, Yoshimori T, Bateman
JF, Kubota H, Nagata K (2009). Autophagic elimination of misfolded
procollagen aggregates in the endoplasmic reticulum as a means of cell
protection. Mol Biol Cell 20, 2744–2754.
Iwakura M, Furusawa K, Kokubu T, Ohashi S, Tanaka Y, Shimura Y, Tsuda K(1992). Dihydrofolate reductase as a new “affinity handle.” J Biochem
111, 37–45.
Jia W, Pua HH, Li QJ, He YW (2011). Autophagy regulates endoplasmic
reticulum homeostasis and calcium mobilization in T lymphocytes. J
Immunol 186, 1564–1574.
Jiang X, Wang X, Ding X, Du M, Li B, Weng X, Zhang J, Li L, Tian R, Zhu Q,
et al. (2020). FAM134B oligomerization drives endoplasmic reticulum
membrane scission for ER-phagy. EMBO J 39, e102608.
Jin X, Zhou H, Song J, Cui H, Luo Y, Jiang H (2021). P3H4 overexpression
serves as a prognostic factor in lung adenocarcinoma. Comput Math
Methods Med 2021, 9971353.
Jumper J, Evans R, Pritzel A, Green T, Figurnov M, Ronneberger O, Tunyasuvunakool K, Bates R, Žídek A, Potapenko A, et al. (2021). Highly accurate protein structure prediction with AlphaFold. Nature 596, 583–589.
Juszkiewicz S, Hegde RS (2018). Quality control of orphaned proteins. Mol
Cell 71, 443–457.
Kang SW, Rane NS, Kim SJ, Garrison JL, Taunton J, Hegde RS (2006).
Substrate-specific translocational attenuation during ER stress defines a
pre-emptive quality control pathway. Cell 127, 999–1013.
Katayama H, Yamamoto A, Mizushima N, Yoshimori T, Miyawaki A (2008). GFPlike proteins stably accumulate in lysosomes. Cell Struct Funct 33, 1–12.
Kayatekin C, Amasino A, Gaglia G, Flannick J, Bonner JM, Fanning S,
Narayan P, Barrasa MI, Pincus D, Landgraf D, et al. (2018). Translocon
declogger Ste24 protects against IAPP oligomer-induced proteotoxicity.
Cell 173, 62–73.e69.
Khaminets A, Heinrich T, Mari M, Grumati P, Huebner AK, Akutsu M,
Liebmann L, Stolz A, Nietzsche S, Koch N, et al. (2015). Regulation of
endoplasmic reticulum turnover by selective autophagy. Nature 522,
354–358.
Kimura S, Noda T, Yoshimori T (2007). Dissection of the autophagosome
maturation process by a novel reporter protein, tandem fluorescenttagged LC3. Autophagy 3, 452–460.
Kitamura T, Koshino Y, Shibata F, Oki T, Nakajima H, Nosaka T, Kumagai H(2003). Retrovirus-mediated gene transfer and expression cloning: powerful tools in functional genomics. Exp Hematol 31, 1007–1014.
Klionsky DJ, Petroni G, Amaravadi RK, Baehrecke EH, Ballabio A, Boya P,
Bravo-San Pedro JM, Cadwell K, Cecconi F, Choi AMK, et al. (2021).
Autophagy in major human diseases. EMBO J 40, e108863.

Volume 34 April 1, 2023

Klöppel G, Löhr M, Habich K, Oberholzer M, Heitz PU (1985). Islet pathology and the pathogenesis of type 1 and type 2 diabetes mellitus
revisited. Surv Synth Pathol Res 4, 110–125.
Kostenko EV, Olabisi OO, Sahay S, Rodriguez PL, Whitehead IP (2006).
Ccpg1, a novel scaffold protein that regulates the activity of the Rho
guanine nucleotide exchange factor Dbs. Mol Cell Biol 26, 8964–
8975.
Kraft C, Deplazes A, Sohrmann M, Peter M (2008). Mature ribosomes are selectively degraded upon starvation by an autophagy pathway requiring
the Ubp3p/Bre5p ubiquitin protease. Nat Cell Biol 10, 602–610.
Lemasters JJ (2005). Selective mitochondrial autophagy, or mitophagy, asa targeted defense against oxidative stress, mitochondrial dysfunction,
and aging. Rejuvenation Res 8, 3–5.
Li W, Ye L, Chen Y, Chen P (2018). P3H4 is correlated with clinicopathologicalfeatures and prognosis in bladder cancer. World J Surg Oncol 16, 206.
Liang CC, Wang C, Peng X, Gan B, Guan JL (2010). Neural-specific deletionof FIP200 leads to cerebellar degeneration caused by increased neuronal death and axon degeneration. J Biol Chem 285, 3499–3509.
Maejima I, Takahashi A, Omori H, Kimura T, Takabatake Y, Saitoh T, Yamamoto A, Hamasaki M, Noda T, Isaka Y, Yoshimori T (2013). Autophagy
sequesters damaged lysosomes to control lysosomal biogenesis andkidney injury. EMBO J 32, 2336–2347.
Melia TJ, Lystad AH, Simonsen A (2020). Autophagosome biogenesis: frommembrane growth to closure. J Cell Biol 219, e202002085.
Mercer TJ, Gubas A, Tooze SA (2018). A molecular perspective of mammalian autophagosome biogenesis. J Biol Chem 293, 5386–5395.
Mizushima N, Komatsu M (2011). Autophagy: renovation of cells and tissues. Cell 147, 728–741.
Mizushima N, Yamamoto A, Hatano M, Kobayashi Y, Kabeya Y, Suzuki K,
Tokuhisa T, Ohsumi Y, Yoshimori T (2001). Dissection of autophagosome
formation using Apg5-deficient mouse embryonic stem cells. J Cell Biol
152, 657–668.
Mochida K, Nakatogawa H (2022). ER-phagy: selective autophagy of theendoplasmic reticulum. EMBO Rep 23, e55192.
Mochida K, Oikawa Y, Kimura Y, Kirisako H, Hirano H, Ohsumi Y,
Nakatogawa H (2015). Receptor-mediated selective autophagy degrades
the endoplasmic reticulum and the nucleus. Nature 522, 359–362.
Mochida K, Yamasaki A, Matoba K, Kirisako H, Noda NN, Nakatogawa H(2020). Super-assembly of ER-phagy receptor Atg40 induces local ER
remodeling at contacts with forming autophagosomal membranes. Nat
Commun 11, 3306.
Molinari M (2021). ER-phagy responses in yeast, plants, and mammaliancells and their crosstalk with UPR and ERAD. Dev Cell 56, 949–966.
Morita K, Hama Y, Izume T, Tamura N, Ueno T, Yamashita Y, Sakamaki Y,
Mimura K, Morishita H, Shihoya W, et al. (2018). Genome-wide CRISPR
screen identifies. J Cell Biol 217, 3817–3828.
Nakagawa I, Amano A, Mizushima N, Yamamoto A, Yamaguchi H, Kamimoto
T, Nara A, Funao J, Nakata M, Tsuda K, et al. (2004). Autophagy defends
cells against invading group A Streptococcus. Science 306, 1037–1040.
Ni M, Lee AS (2007). ER chaperones in mammalian development and human diseases. FEBS Lett 581, 3641–3651.
Nthiga TM, Kumar Shrestha B, Sjøttem E, Bruun JA, Bowitz Larsen K,
Bhujabal Z, Lamark T, Johansen T (2020). CALCOCO1 acts with VAMPassociated proteins to mediate ER-phagy. EMBO J 39, e103649.
Ochs RL, Stein TW, Chan EK, Ruutu M, Tan EM (1996). cDNA cloning andcharacterization of a novel nucleolar protein. Mol Biol Cell 7, 1015–
1024.
Øverbye A, Fengsrud M, Seglen PO (2007). Proteomic analysis of membrane-associated proteins from rat liver autophagosomes. Autophagy 3,
300–322.
Pedrazzini E, Villa A, Longhi R, Bulbarelli A, Borgese N (2000). Mechanismof residence of cytochrome b(5), a tail-anchored protein, in the endoplasmic reticulum. J Cell Biol 148, 899–914.
Pengo N, Scolari M, Oliva L, Milan E, Mainoldi F, Raimondi A, Fagioli C,
Merlini A, Mariani E, Pasqualetto E, et al. (2013). Plasma cells require
autophagy for sustainable immunoglobulin production. Nat Immunol
14, 298–305.
Rappsilber J, Mann M, Ishihama Y (2007). Protocol for micro-purification,enrichment, pre-fractionation and storage of peptides for proteomics
using StageTips. Nat Protoc 2, 1896–1906.
Ruggiano A, Foresti O, Carvalho P (2014). Quality control: ER-associateddegradation: protein quality control and beyond. J Cell Biol 204,
869–879.
Rutishauser J, Spiess M (2002). Endoplasmic reticulum storage diseases.
Swiss Med Wkly 132, 211–222.

CCPG1 ER-phagy| 17

Saitoh T, Fujita N, Hayashi T, Takahara K, Satoh T, Lee H, Matsunaga K,
Kageyama S, Omori H, Noda T, et al. (2009). Atg9a controls dsDNAdriven dynamic translocation of STING and the innate immune
response. Proc Natl Acad Sci USA 106, 20842–20846.
Schröder M, Kaufman RJ (2005). ER stress and the unfolded protein response. Mutat Res 569, 29–63.
Shimizu Y, Inoue A, Tomari Y, Suzuki T, Yokogawa T, Nishikawa K, Ueda T(2001). Cell-free translation reconstituted with purified components.
Nat Biotechnol 19, 751–755.
Shimizu Y, Kanamori T, Ueda T (2005). Protein synthesis by pure translation
systems. Methods 36, 299–304.
Singh R, Kaushik S, Wang Y, Xiang Y, Novak I, Komatsu M, Tanaka K, Cuervo
AM, Czaja MJ (2009). Autophagy regulates lipid metabolism. Nature
458, 1131–1135.
Smith M, Wilkinson S (2017). ER homeostasis and autophagy. Essays Biochem 61, 625–635.
Smith MD, Harley ME, Kemp AJ, Wills J, Lee M, Arends M, von Kriegsheim
A, Behrends C, Wilkinson S (2018). CCPG1 is a non-canonical autophagy
cargo receptor essential for ER-phagy and pancreatic ER proteostasis.
Dev Cell 44, 217–232.e211.
Stephani M, Picchianti L, Gajic A, Beveridge R, Skarwan E, Sanchez
de Medina Hernandez V, Mohseni A, Clavel M, Zeng Y, NaumannC, et al. (2020). A cross-kingdom conserved ER-phagy receptor
maintains endoplasmic reticulum homeostasis during stress. Elife 9,
e58396.
Stolz A, Ernst A, Dikic I (2014). Cargo recognition and trafficking in selective
autophagy. Nat Cell Biol 16, 495–501.
Sun J, Wang W, Zheng H (2022). ROOT HAIR DEFECTIVE3 is a receptor for
selective autophagy of the endoplasmic reticulum in Arabidopsis. Front
Plant Sci 13, 817251.
Tian X, Jin RU, Bredemeyer AJ, Oates EJ, Błazewska KM, McKenna CE,
Mills JC (2010). RAB26 and RAB3D are direct transcriptional targetsof MIST1 that regulate exocrine granule maturation. Mol Cell Biol 30,
1269–1284.

18 | S. Ishii et al.

Turco E, Witt M, Abert C, Bock-Bierbaum T, Su MY, Trapannone R, Sztacho
M, Danieli A, Shi X, Zaffagnini G, et al. (2019). FIP200 claw domain binding to p62 promotes autophagosome formation at ubiquitin condensates. Mol Cell 74, 330–346.e311.
Uhlén M, Fagerberg L, Hallström BM, Lindskog C, Oksvold P, Mardinoglu A,
Sivertsson Å, Kampf C, Sjöstedt E, Asplund A, et al. (2015). Proteomics.
Tissue-based map of the human proteome. Science 347, 1260419.
Varadi M, Anyango S, Deshpande M, Nair S, Natassia C, Yordanova G,
Yuan D, Stroe O, Wood G, Laydon A, et al. (2022). AlphaFold protein
structure database: massively expanding the structural coverage ofprotein-sequence space with high-accuracy models. Nucleic Acids Res
50, D439–D444.
Webb JL, Ravikumar B, Atkins J, Skepper JN, Rubinsztein DC (2003). Alphasynuclein is degraded by both autophagy and the proteasome. J Biol
Chem 278, 25009–25013.
Wilfling F, Lee CW, Erdmann PS, Zheng Y, Sherpa D, Jentsch S, Pfander
B, Schulman BA, Baumeister W (2020). A selective autophagy pathway for phase-separated endocytic protein deposits. Mol Cell 80,
764–778.e767.
Wilkinson S (2020). Emerging principles of selective ER autophagy.
J Molecular Biol 432, 185–205.
Yamasaki A, Alam JM, Noshiro D, Hirata E, Fujioka Y, Suzuki K, Ohsumi
Y, Noda NN (2020). Liquidity is a critical determinant for selective autophagy of protein condensates. Mol Cell 77, 1163–1175.e1169.
Yim WW, Yamamoto H, Mizushima N (2022). A pulse-chasable reporter
processing assay for mammalian autophagic flux with HaloTag. Elife 11,
e78923.
Yoshii SR, Kuma A, Akashi T, Hara T, Yamamoto A, Kurikawa Y, Itakura E,
Tsukamoto S, Shitara H, Eishi Y, Mizushima N (2016). Systemic analysis
of Atg5-null mice rescued from neonatal lethality by transgenic ATG5
expression in neurons. Dev Cell 39, 116–130.
Zhao D, Zou CX, Liu XM, Jiang ZD, Yu ZQ, Suo F, Du TY, Dong MQ, He W,
Du LL (2020). A UPR-induced soluble ER-phagy receptor acts with VAPs
to confer ER stress resistance. Mol Cell 79, 963–977.e963.

Molecular Biology of the Cell