Inhibition of Endoplasmic Reticulum Associated Degradation Reduces Endoplasmic Reticulum Stress and Alters Lysosomal Morphology and Distribution

Abstract

Disturbances in proteostasis are observed in many neurodegenerative diseases. This leads to activation of protein quality control to restore proteostasis, with a key role for the removal of aberrant proteins by proteolysis. The unfolded protein response (UPR) is a protein quality control mechanism of the endoplasmic reticulum (ER) that is activated in several neurodegenerative diseases. Recently we showed that the major proteolytic pathway during UPR activation is via the autophagy/lysosomal system. Here we investigate UPR induction if the other major proteolytic pathway of the ER -ER associated degradation (ERAD)-is inhibited. Surprisingly, impairment of ERAD results in decreased UPR activation and protects against ER stress toxicity. Autophagy induction is not affected under these conditions, however, a striking relocalization of the lysosomes is observed. Our data suggest that a protective UPR-modulating mechanism is activated if ERAD is inhibited, which involves lysosomes. Our data provide insight in the cross-talk between proteolytic pathways involved in ER proteostasis. This has implications for neurodegenerative diseases like Alzheimer’s disease where disturbed ER proteostasis and proteolytic impairment are early phenomena in the pathology.

INTRODUCTION

The maintenance of proteostasis is of major importance for the function of cells. The underlying cause of several human diseases may be disturbance of proteostasis (Balch et al., 2008). This is illustrated in neurodegenerative disorders that are characterized by accumulation of aberrant proteins and -apparently ineffective- activation of protein quality control mechanisms (Scheper and Hoozemans, 2009). The unfolded protein response (UPR) of the endoplasmic reticulum (ER) is such a mechanism. We reported previously that the UPR is activated early in Alzheimer’s disease (AD) and other tauopathies (Hoozemans et al., 2005; 2009; Nijholt et al., 2012).

The UPR activates upon disturbances (ER stress) in the ER homeostasis that lead to increased protein misfolding. In resting state, the ER chaperone binding immunoglobulin (heavy chain) protein/glucose-related protein 78 (BiP/GRP78) binds three sensor proteins in the ER membrane. In case of ER stress, BiP releases from the sensors and the three signalling pathways of the UPR are activated (Ron and Walter, 2007). These pathways are involved in translational regulation and the induction of a transcriptional response. This results in an overall decreased protein load in the ER and selective upregulation of proteins that assist in protein folding (e.g. BiP) and degradation. Activation of the UPR is initiated to restore homeostasis in the ER. If ER homeostasis is restored, the UPR will be switched off, but prolonged activation will result in cell death (Szegezdi et al., 2006).

An important mechanism in the restoration of ER proteostasis is the removal of misfolded proteins. Misfolded ER proteins can be degraded by the ubiquitin proteasome system (UPS) in the cytosol after translocation from the ER to the cytosol, a process called ER associated degradation (ERAD), which requires factors in both the ER and the cytosol (Vembar and Brodsky, 2008). Newly synthesized ER polypeptides are modified with oligosaccharide precursors and subsequently trimmed by ER glucosidases to allow recognition by the lectin chaperones calnexin and calreticulin (Hebert et al., 2005). Removal of the last glucose residue marks properly folded proteins that can exit the ER through the secretory pathway.

In contrast, misfolded intermediates are reglucosylated and re-enter the glucosylation/deglucosylation lectin cycle which may be repeated until the proteins are recognized as irreversibly misfolded leading to removal of mannose residues by ER mannosidase I (Fagioli and Sitia, 2001; Nakatsukasa and Brodsky, 2008). This facilitates recognition by the ER degradation-enhancing α-mannosidase-like lectins [EDEMs; (Molinari et al., 2003)], ultimately resulting in retrotranslocation and degradation by the UPS (Fagioli and Sitia, 2001; Hosokawa et al., 2003). An alternative pathway for degradation of ER proteins is via autophagy. Autophagy involves the sequestering of material that needs to be degraded by a double membrane structure, followed by fusion with a lysosome and degradation by lysosomal enzymes.

The activity of both degradational pathways is regulated by the UPR. ERAD is directly affected by the UPR, for example activation of the UPR increases the levels of EDEM1, which changes the recognition of aberrant proteins (Ron et al., 2011). Activation of the UPR also triggers autophagy (Bernales et al., 2006; Ding et al., 2007; Ogata et al., 2006) and our group previously showed that during UPR activation autophagy is the major degradational pathway (Nijholt et al., 2011a; Scheper et al., 2011). It is possible that, under these stress conditions, parts of the ER are directly targeted for clearance by the autophagy/lysosomal system (Bernales et al., 2007).

Because ER stress is persistent in AD and other tauopathies, mechanisms to restore ER proteostasis may provide potential targets for intervention. In addition, both the proteasome and the autophagy/lysosomal systems are impaired in neurodegenerative diseases (Nijholt et al., 2011b). Therefore, further understanding of the regulation and interplay of the different proteostatic pathways is essential to design a therapeutic strategy based on proteostatic regulation. Here we investigate the behaviour of the UPR under conditions where ERAD is impaired. Counter intuitively, inhibition of ERAD attenuates UPR activity and protects against ER stress toxicity. Concomitantly we observe relocalization of lysosomes. Our data suggest that a protective pathway is activated if ERAD is impaired which does not involve induction of classical autophagy, but is related to the autophagy/lysosomal system.

MATERIALS AND METHODS

Materials

Cell culture media and reagents were obtained from Gibco/Invitrogen (USA) and other chemicals were from Sigma (USA), unless indicated otherwise.

Cell culture and treatment

SK-N-SH, HeLa cells and inducible MEF Atg5^−/− cells were cultured in Dulbecco’s modified Eagle medium with GlutaMAX supplemented with 10% (v/v) fetal calf serum (Lonza, Switzerland), 100 U/ml penicillin and 100 μg/ml streptomycin. Inducible MEF Atg5^−/− cells were a kind gift of Dr. N. Mizushima (Hosokawa et al., 2006). MEF Atg5^−/− cells were maintained in the presence of doxycylin (20 ng/ml) and cultured in the absence or presence of doxycylin for the Atg5^+/+ or Atg5^−/− genotype, respectively. Cells were incubated at 37°C, 5% CO₂ and 95% humidity. SK-N-SH cells were plated in a desired wells format at a density of ∼50.10³ cells/cm² in complete culture medium supplemented with 20 μM retinoic acid. Preconditioning with 1 μg/ml kifunensine (Calbiochem, EMD Millipore, USA) lasted 72 h (or as indicated), and was followed by tunicamycin and kifunensine treatment (as indicated) for 20 h. Treatment with Earle’s balanced salt solution (EBSS; Sigma) lasted 2 h, also, while maintaining kifunensine pressure.

RNA isolation and cDNA synthesis

The procedures for RNA isolation and cDNA synthesis are described previously (Elfrink et al., 2012). Briefly, RNA was isolated using TRIzol Reagent according to the manufacturer’s protocol (Invitrogen). cDNA synthesis was performed using a SuperScript II Reverse Transcriptase Kit (Invitrogen) on equal quantities of RNA. Oligo(dT)_12-VN primers (125 pmol) were used to prime mRNA poly-A tails.

Real-Time qPCR

The procedures for qPCR are described elsewhere (Elfrink et al., 2012). Briefly, equal quantities of triplicate cDNA samples were dried in a 384 wells plate. qPCR reactions were performed in a LightCycler 480 system (Roche, Germany). Probe and primer combinations are listed in Table 1. Results were analyzed using the LightCycler 480 software (version 1.5.0.39). Data are presented as mean± SD (n = 3) from a representative experiment of three.

Table 1.

Primer and probe combinations for qPCR

Amplicon	Primers (5′-3′)	Probea
BiP/Grp78	Fw: CATCAAGTTCTTGCCGTTCA	#10
	Rv: TCTTCAGGAGCAAATGTCTTTGT
CHOP/GADD153	Fw: AAGGCACTGAGCGTATCATGT	#21
	Rv: TGAAGATACACTTCCTTCTTGAACA
GAPDH	Fw: TCCACTGGCGTCTTCACC	#45
	Rv: GGCAGAGATGATGACCCTTTT

Primers were prepared by Sigma.

^aReferring to Universal ProbeLibrary for Human probes (Roche).

MTT viability assay

After treatment, cells were incubated with 250 μg/ml 3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) in complete culture medium (2 h, 37°C). The formazan salts were dissolved in DMSO and the OD^{570 nm} was measured using a BMG FLUOstar Omega Microplate Reader (BMG Labtech, Germany). Normalized data are presented as mean ± SEM.

Statistical analysis

Statistical analyses were performed using the unpaired two-tailed Student’s t-test and differences were accepted as statistically significant at a level of P ≤ 0.05.

SDS-PAGE and Western blotting

Cells were harvested by scraping with a rubber policeman in 1% (v/v) Triton X-100 PBS lysis buffer supplemented with protease inhibitors (Leupeptin and PMSF). Cell lysates were vigorously mixed, incubated on ice (5 min), and centrifuged (20, 000 × g, 5 min, 4°C). Supernatant protein content was determined by Bio-Rad Protein Assay (Bio-Rad, USA). Samples were analyzed for LC3 and CD147 on 18% and 10% polyacrylamide gels, respectively. Equal amounts of protein were loaded in each lane on a gel. Antibodies and incubation conditions are specified in Table 2. Visualization was performed with Lumi-Light Western Blotting Substrate (Roche) on a LAS-3000 imaging system (Fujifilm, Japan).

Table 2.

Antibodies used for western blot and immunofluorescence

Antibody	Species	Clonality	Company	Cat. no.
Western blottinga
LC3	Rabbit	Polyclonal	Novus Biologicals	NB100-2220
CD147	Goat	Polyclonal (C-19)	Santa Cruz	Sc-9754
GAPDH	Mouse	Monoclonal (6C5)	EMD Millipore	MAB374
Immunofluorescenceb
LC3c	Mouse	Monoclonal (4E12)	MBL	M152-3
LAMP1	Mouse	Monoclonal (H5G11)	Santa Cruz	sc-18821
Cathepsin D	Goat	Polyclonal	R&D Systems	AF1014

^aWestern blot dilutions were 1:1000 in 5% (w/v) fat-free dried milk in (0.05%, v/v) PBS-T, secondary antibodies were from Dako (Glostrup, Denmark).

^bImmunofluorescence dilutions were 1:100 in 5% (v/v) fish skin gelatin in PBS, secondary antibodies were from Jackson ImmunoResearch (West Grove, PA, USA).

^cCoverslips were not air dried after demi water rinse, but immediately mounted on glass slides with antifade reagent.

Immunofluorescence and confocal microscopy

For immunofluorescence stainings, cells were plated on glass coverslips. All following procedures were performed in/with PBS and at room temperature. After treatment, cells were fixed (4% [w/v] paraformaldehyde, 4% [w/v] sucrose [Merck], 15 min) and permeabilized (0.5% [v/v] Triton X-100, 5 min). Washes after formalin fixation included 187 μM glycine. Coverslips were transferred to a dark moist chamber, blocked (5% [v/v] fish skin gelatin, 30 min) and incubated with primary antibody in block buffer for 2 h. Cells were incubated with secondary antibodies in block buffer for 1 h. Washes after antibody incubations were performed with 0.05% (v/v) Tween-20. Cells were counter stained with DAPI (1.3 μM, 5 min). Coverslips were rinsed (demi water), air dried, and mounted on glass slides with Prolong Gold (Invitrogen). Antibodies and incubation conditions are listed in Table 2. Confocal imaging was performed on a Leica TCS-SP2 confocal scanner mounted on an inverted microscope (Leica Microsystems, Germany). Images were acquired with Leica Confocal Software (version 2.61, build 1537). Confocal images were deconvoluted using a theoretical point spread function by Huygens Essential Software (compute engine 4.1.1p1 64b; Scientific Volume Imaging, Hilversum, the Netherlands). Cathepsin D punctae were quantified on three fields of view, using ImageJ software (version 1.46r, National Institutes of Health, USA), with the following parameters: punctae per cell and area, perimeter and circularity of the punctae.

RESULTS

In this study, kifunensine was used to inhibit ERAD. Kifunensine inhibits ER α-1, 2-mannosidase I (including EDEM) which is required for the early steps of ERAD substrate recognition (Wang et al., 2011). Differentiated SK-N-SH cells were preconditioned with 1 μg/ml kifunensine for 24 and 72 h to mimic chronic impairment as expected in neurodegenerative diseases. The endogenous ERAD substrate CD147 (Tyler et al., 2012) accumulates in SK-N-SH cells treated with kifunensine for 72 hours, confirming the inhibition of ERAD by kifunensine (Supplementary Fig. 1). The UPR was induced by the ER stressor tunicamycin at the indicated concentrations for 20 h while kifunensine pressure was maintained. After 20 h of tunicamycin treatment, the UPR is fully activated beyond the initial signaling events, furthermore the induction of the UPR follows a dose dependent response at 0.2 and 0.5 μg/ml. Lower concentrations do not elicit a reproducible UPR response and at higher concentrations the increased toxicity interferes. The dose dependancy is demonstrated by the increased mRNA levels of the established UPR targets BiP and CHOP [apoptotic factor cAMP response element-binding (C/EPB) homologous protein; Figs. 1A and 1B]. The UPR markers are slightly increased at basal level by treatment with kifunensine, which could suggest sensitization of the UPR. However, this effect does not persist when the UPR is stimulated with tunicamycin; treatment with kifunensine strongly reduces the induction of the UPR targets. BiP mRNA is reduced approximately 3-fold for all treatment conditions (Fig. 1A) and CHOP mRNA is reduced approximately 8-fold (0.2 μg/ml tunicamycin) and 4-fold (0.5 μg/ml tunicamycin; Fig. 1B).

Fig. 1.

Kifunensine reduces UPR activation and UPR mediated cytotoxicity. SK-N-SH cells were preconditioned for 24 and 72 h with 1 μg/ml kifunensine. Treatment was performed with tunicamycin and kifunensine for 20 h at the indicated concentrations. The effect of kifunensine on UPR induction was assessed by qPCR analysis of BiP (A) and CHOP (B) mRNA. GAPDH was used as a reference gene. Kif: Kifunensine. Shown are mean and SD in arbitrary units (AU; n = 3, statistical differences are indicated by ^*, ^** and ^*** for P ≤ 0.05, P ≤ 0.01 and P ≤ 0.001, respectively). (C) MTT viability assay was performed to assess tunicamycin induced cytotoxicity in the absence or presence of kifunensine. Shown are mean and SEM values normalized to control of two normalized experiments (n = 12, statistical differences at a significance level of P ≤ 0.001 are indicated by ^***).

The viability of the cells was determined to test whether kifunensine also reduces the toxicity of the ER stress stimulus. Cellular survival is reduced, as expected, with tunicamycin treatment (Fig. 1C: control). On the other hand, survival increases 28% (0.2 and 0.5 μg/ml tunicamycin) and 16% (1.0 μg/ml tunicamycin) upon treatment with kifunensine (Fig. 1C: kifunensine). This protective effect of kifunensine on UPR activation and ER stress toxicity is not cell type specific, because it is also observed in the non-neuronal HeLa cells (Supplementary Fig. 2). Our data show that inhibition of ERAD using kifunensine renders cells less susceptible to ER stress, demonstrated by a reduction in UPR activation accompanied by a reduction in ER stress toxicity. Preconditioning with kifunensine is required for this protective effect (Supplementary Fig. 3), because addition of the kifunensine at the start of the tunicamycin treatment does not result in protection. This suggests that in response to ERAD inhibition a mechanism is activated that reduces the negative effects of ER stress.

Since ER α-1, 2-mannosidase I inhibition by kifunensine compromises ERAD, an alternative system must reduce ER stress. Autophagy is activated by the UPR, and we therefore investigated if autophagy is enhanced by ER α-1, 2-mannosidase I inhibition. Kifunensin does not affect the quantity and quality of the LC3 positive punctate structures (Fig. 2A). In addition, induction of autophagy was assessed by LC3-I to LC3-II processing on Western blot. Kifunensine preconditioning does not alter the processing of LC3 on basal level, by tunicamycin induced autophagy, and by amino acid induced autophagy starvation (EBSS; Fig. 2B) Furthermore, the functional requirement for autophagy was tested in inducible autophagy-defective Atg5^−/− MEF cells with inducible autophagy competence (Hosokawa et al., 2006). As in SK-N-SH and HeLa cells, the UPR mediated toxicity and the rescue by kifunensine preconditioning was observed in this cell system. In the presence of doxycycline the cells become autophagy-defective, however, the rescue of UPR mediated toxicity by kifunensine was still observed (Fig. 2C). These data demonstrate that kifunensine does not activate autophagy and moreover that its effects are not dependent on functional autophagy; therefore it must lead to activation of a different protective mechanism.

Fig. 2.

Kifunensine mediated ER stress protection does not involve autophagy. SK-N-SH cells were preconditioned with kifunensine for 72 h. Cells were treated with tunicamycin (1 μg/ml) for 20 h or EBSS for 2 h (both with co-treatment of kifunensine). (A) LC3 punctate structures were visualized by immunofluorescent staining. Nuclei were counterstained with DAPI. Con, control; Tm, tunicamycin; EBSS, Earle’s balanced salt solution. Scale bar indicates 10 μm. (B) Western blot analysis was performed for LC3-I to LC3-II conversion. Equal amounts of protein were loaded in each lane, and GAPDH was used as a loading control. (C) Inducible MEF Atg5^−/− cells were preconditioned with kifunensine for 72 h and treated with tunicamycin and kifunensine for 20 h. The MTT viability assay was performed in the absence or presence of doxycyclin, inducing Atg5^+/+ wt and Atg5^−/− autophagy-defective cells, respectively. UPR toxicity was analyzed by MTT. Shown are mean and SEM values normalized to control (n = 6, statistical differences are indicated by ^* and ^** for P ≤ 0.05 and P ≤ 0.01, respectively). Kif, Kifunensine.

The autophagic flux finalizes with the fusion of autophagosomes with degradative bodies, i.e. the lysosomes. Kifunensine does not affect the induction of autophagy (Fig. 2), but this does not exclude effects on the lysosomes directly. Therefore we used cathepsin D (Fig. 3A) and lysosome associated membrane protein 1 (LAMP1; Fig. 3B) to visualize the lysosomes in SK-N-SH cells. In the untreated cells both cathepsin D and LAMP1 staining show a punctate pattern, distributed throughout the cell (Figs. 3A and 3B). Kifunensine treatment increases the number, size and perimeter but not the circularity of the Cathepsin D positive structures (Table 3). ER stress induction by tunicamycin treatment relocalizes the LAMP1 positive structures to a juxtanuclear site. Positioning of lysosomes to a juxtanuclear site also occurs during nutrient stress (Korolchuk et al., 2011). The tunicamycin induced repositioning of the LAMP1 positive structures is strongly inhibited in the presence of kifunensine, and is similar to the pattern of LAMP1 distribution in untreated cells (Fig. 3B). These results demonstrate that lysosomal morphology and distribution is changed by ERAD inhibition by kifunensine, which may be indicative for altered lysosomal function.

Fig. 3.

Kifunensine alters the size and number of lysosomal punctae. SK-N-SH cells were preconditioned with kifunensine for 72 h. Treatment was performed with tunicamycin (1 μg/ml) and kifunensine treatment for 20 h. Immunofluorescent staining was performed for Cathepsin D (A) and LAMP1 (B). Nuclear counterstain was performed with DAPI. Scale bar indicates 10 μm. Quantifications of numbers, area, perimeter and circularity were performed as described in materials and methods. CathD, Cathepsin D.

Table 3.

Quantification of Cathepsin D positive punctea

	Control		Kifunensine
	Average	SD	Average	SD	p-value
Punctae per cell	4.07	0.40	4.94	0.29	0.037
Puncta area (μm2)	0.51	0.04	0.68	0.04	0.035
Puncta perimeter (μm)	2.49	0.07	2.99	0.12	0.023
Puncta circularity	0.88	0.01	0.88	0.01	0.640

DISCUSSION

In this paper we provide evidence that chronic inhibition of ERAD using kifunensine activates a pathway that protects against ER stress. Kifunensine inhibits ER α-1, 2-mannosidase I (Avezov et al., 2008; Fagioli and Sitia, 2001; Wang et al., 2011) and therefore interferes with the early recognition of ERAD substrates and is expected to decrease the retrotranslocation of ERAD substrates. To mimick chronic impairment of ERAD in a cell model we perfomed kifunensine treatment for 24 and 72 h which may be considered as persistent. ERAD is inhibited under these conditions, demonstrated by accumulation of the endogenous ERAD substrate CD147. Surprisingly this prolonged inhibition of ERAD is well tolerated by the cells. Chronic impairment of ER α-1, 2-mannosidase I is also found in patients with mutations in the MAN1B1 gene. This leads to autosomal-recessive intellectual disability, but apart from the developmental problem that underlies the clinical presentation, chronically low levels of the ER α-1, 2-mannosidase I are well tolerated (Rafiq et al., 2011), suggesting activation of compensatory mechanism for the disturbed ERAD function. This suggestion is supported by several in vitro studies. Inhibition of ERAD using kifunensine did not lead to strong UPR activation or to UPR mediated apoptosis (Wang et al., 2011), despite the increased retention of aberrant proteins in the ER. Also inhibition of ERAD via reduction of Herp (an ERAD associated ubiquitin-like protein) is shown to improve cell viability when the proteasomal function is impaired (Miura et al., 2010). In fact, hyperactivation of ERAD via overexpression of the translocon component Sec61α increases neural proteotoxicity in Drosophila. This is probably caused by accumulation of aberrant proteins in the cytosol by the inability of the UPS to deal with the increased translocation (Kanuka et al., 2003; 2005). Interestingly, inhibition of ERAD at the level of the cytosolic ERAD factor p97/VCP did lead to UPR activation and ER stress mediated apoptosis. This indicates that not inhibition of ERAD per se, but inhibition at specific steps can provide protection from ER stress (Wang et al., 2011).

The protective effect of ERAD inhibition against ER stress appears paradoxical, but may actually indicate the presence of tight regulation and solid back-up systems to maintain proteostasis. The observation that preconditioning with kifunensine is required to achieve protection also indicates that it induces a functional adaptation to decreased ERAD capacity. A likely candidate pathway that deals with the elevated ER stress if ERAD is impaired is the autophagy/lysosomal pathway. There is an intricate but not fully elucidated interplay between ERAD and autophagy via the UPR. Deficiency for the transcription factor XBP-1, that is activated via the Ire1 pathway of the UPR, reduces ERAD and induces autophagy (Hetz et al., 2009). Furthermore, knockdown of the ERAD factor EDEM induces LC3 positive punctae (Hetz et al., 2009).

We have shown that autophagy induction is not affected by kifunensine treatment, moreover protection against ER stress is also observed in autophagy deficient cells. Interestingly, we find that the number of lysosomal punctae, but also the size and perimeter of lysosomal punctae are increased. In addition, the positioning of lysosomes is affected by kifunensine treatment. Under nutrient stress conditions lysosomes reposition to a juxtanuclear site (Korolchuk et al., 2011), which we also observe under ER stress in our experiments. In cells preconditioned with kifunensine, an ER stress stimulus does not result in this typical ER stress induced lysosomal repositioning. Instead the lysosomal position is similar to that in untreated cells. Responses to ER stress and nutrient stress share common pathways in the integrated stress response (Harding et al., 2003) and our data suggest that the lysosomes may play a role in different stress responses.

There are data emerging about degradational pathways that resemble autophagy or use components of the autophagy/lysosomal machinery, but are different from classical macroautophagy. A neuron-specific alternative endo-lysosomal degradation pathway was recently reported, which protects against neurodegeneration (Haberman et al., 2012). An interesting example is provided by “EDEMosomes”, degradational compartments where ERAD factors are segregated and subjected to rapid turn-over (Cali et al., 2008). This “ERAD tuning” is thought to limit the activity of ERAD only to proteins that are really terminally misfolded. The degradation involves LC3 positive structures and an autophagy-like process, but not classical macroautophagy.

We have previously shown activation of the UPR early in AD pathology (Hoozemans et al., 2005; 2009). Lysosomal dysfunction is also an early phenomenon in AD (Nixon and Yang, 2011) and may therefore contribute to a vicious pathological cycle where ER stress cannot be resolved because of diminished lysosomal capacity. Our data suggest that inhibition of ERAD positively affects the ER stress toxicity via the lysosomes. Further studies are required to further address the regulation of the protective mechanisms against ER stress, and whether this may be employed to intervene early in the pathogenesis of neurodegenerative diseases.

Note:

Supplementary information is available on the Molecules and Cells website (www.molcells.org).