N-glycosylation-negative catalase: a useful tool for exploring the role of hydrogen peroxide in the endoplasmic reticulum

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N-glycosylation-negative catalase: a useful tool for exploring the role of hydrogen peroxide in the endoplasmic reticulum S. Lortz, S. Lenzen, I. Mehmeti

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Free Radical Biology and Medicine

Received date: 30 July 2014 Revised date: 7 November 2014 Accepted date: 27 November 2014 Cite this article as: S. Lortz, S. Lenzen, I. Mehmeti, N-glycosylation-negative catalase: a useful tool for exploring the role of hydrogen peroxide in the endoplasmic reticulum, Free Radical Biology and Medicine, http://dx.doi.org/ 10.1016/j.freeradbiomed.2014.11.024 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting galley proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

N-glycosylation-negative catalase: a useful tool for exploring the role of hydrogen peroxide in the endoplasmic reticulum

S. Lortz*, S. Lenzen, I. Mehmeti

Institute of Clinical Biochemistry, Hannover Medical School, 30623 Hannover, Germany *Address correspondence to: Dr. Stephan Lortz Institute of Clinical Biochemistry Hannover Medical School 30623 Hannover Germany Telephone: + 49/511/5323765 Fax: + 49/511/5323584 [email protected]

Keywords: H2O2; catalase; endoplasmic reticulum; disulfide bond formation; protein folding.

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Abstract Disulfide bond formation during protein folding of nascent proteins is associated with the generation of H2O2 in the endoplasmic reticulum (ER). Approaches to quantify H2O2 directly within the ER failed due to the oxidative environment in the ER lumen and ERspecific catalase expression to detoxify high H2O2 concentrations resulted in an inactive protein due to N-glycosylation. Therefore, the N-glycosylation motives at asparagine-244 and -439 of the human catalase protein were deleted by side-directed mutagenesis. The ERtargeted expression of these variants revealed that only the deletion of the N-glycosylation motive at asparagine-244 (N244) was associated with the maintenance of full enzymatic activity in the ER. Expression of catalase N244 in the ER (ER-Catalase N244) was ERspecific and protected the cells significantly against exogenously added H2O2. With the expression of ER-Catalase N244, a highly effective H2O2 inactivation within the ER was achieved for the first time. Catalase has a high H2O2 inactivation capacity without the need of reducing cofactors, which might interfere with the ER redox homeostasis, and is not involved in protein folding. With these characteristics ER-Catalase N244 is an ideal tool to explore the impact of ER-generated H2O2 for the generation of disulfide bonds or to study the induction of ER-stress pathways through protein folding overload and accumulation of H2O2.

Research Highlights: •

Human catalase, expressed in the ER, is N-glycosylated at asparagine-244

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N-glycosylation at asparagine-244 is responsible for loss of enzymatic activity

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Prevention of N-glycosylation at asparagine-244 restored enzyme activity

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ER-Catalase N244 is fully active and protected cells against H2O2 toxicity

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ER-Catalase N244 is a powerful tool to explore the role of H2O2 within the ER

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Introduction The folding of most nascent proteins within the ER is accompanied by the introduction of disulfide bonds mediated by specialized ER-oxidoreductases [1]. This family comprises, as most prominent examples, the protein disulfide isomerase (PDI) and the sulfhydryl oxidase endoplasmic reticulum oxidoreductin 1 (ERO-1) [2]. PDI is responsible for the oxidation of cysteine residues in newly synthesized proteins, whereas ERO-1 catalyzes the re-oxidation of PDI through the delivery of the electrons to molecular oxygen, generating thereby stoichiometric amounts of H2O2 for each disulfide bond produced (for review see [3]). For highly secretory active cells such as endo- and exocrine cells of the pancreas, plasma cells or hepatocytes, the proportion of these secreted proteins could account for up to 70% of the overall cellular protein synthesis capacity [4]. Consequently also high amounts of H2O2 are coincidentally released by this enzymatic folding pathway which should result in extremely high concentrations of ER-located H2O2. Whether this high H2O2 production is necessary or deleterious for protein folding and ER redox homeostasis is quite controversially discussed at present. On the one hand it seems that luminal H2O2 is reutilized through ER-specific peroxidases like glutathione peroxidase 7 and 8 (GPx7and GPx8) and probably also peroxiredoxin 4 (PRDXIV) to maintain or even booster oxidative protein folding [3, 5]. In contrast to these findings an excessive ER-derived H2O2 production due to overwhelming protein processing is associated with the activation of ERstress pathways leading to apoptotic cell death [6-8]. These mechanisms have been considered to be of great importance for the development of nonalcoholic steatohepatitis or type 2 diabetes mellitus [9, 10]. However, most of the conclusions made were achieved indirectly through inhibition/knock down or overexpression of elements of the protein folding machinery, since the oxidizing environment hampers the direct quantification of H2O2 in the ER through the H2O2-sensitive HyPer sensor protein [11, 12]. Also, effective and protein folding-independent 3

inactivation of H2O2 in the ER was not possible due to inactivation of the catalase enzyme through N-glycosylation [12]. Therefore, it was our aim to construct an N-glycosylation-negative catalase mutant for ER-specific expression with an enzyme activity comparable to that of catalase overexpressed in peroxisomes or mitochondria [13]. We present here an ER-active catalase variant as a powerful experimental tool with which the role of H2O2 for proper ER-function or ERdestruction can be investigated in detail.

Materials and methods Catalase mutagenesis and generation of expression plasmids For the deletion of the two N-glycosylation motives in the wild type catalase cDNA, the pCAT10 plasmid (ATCC, Manassas, VA, USA), which contains full-length catalase cDNA in the pSP65 vector, was used as template for the site-specific mutagenesis through the Q5 Site-Directed Mutagenesis Kit according to the manufacturer’s recommendation (New England Biolabs, Ipswich, MA, USA). The primer sequences used for mutagenesis are listed in Table 1. For the double deletion variant (N244/N439), mutations were introduced subsequently. For peroxisomal expression the mutated catalase cDNAs were subcloned into the EcoRV restriction site of the pLenti6.3/V5-MCS plasmid by standard PCR techniques. To add the ER-targeting and ER-retention signal elements necessary for ER-specific expression, the mutated catalase cDNAs were subcloned into the PstI/NotI restriction sites of the pCMV/myc/ER plasmid using PCR composite primers with PstI (frw primer) and NotI (rev primer) restriction enzyme recognition sites. Thereafter the whole expression cassette was transferred also into the EcoRV restriction site of the pLenti6.3/V5-MCS plasmid by PCRmediated subcloning. All plasmids were verified by sequencing. Cell culture and lentiviral transduction Insulin-producing RINm5F tissue culture cells were cultured in RPMI-1640 medium supplemented with 10 mM glucose, 10% (v/v) fetal calf serum (FCS), penicillin, and 4

streptomycin in a humidified atmosphere at 37°C and 5% CO2 as described previously [14]. To express one of the three mutated different catalase cDNAs in RINm5F cells, lentiviral particles were prepared as described before in detail [15]: 4 x 106 293T cells were transfected with the packaging plasmid pPAX2 (11.25 μg), the envelope plasmid pcDNA-MDG (3.75 μg), and the transfer plasmids pLenti6.3/V5-MCS-catalase N244, N439, or N244/439 for peroxisomal expression or ER-Catalase N244, N439 or N244/439 for ER-specific expression (15 μg) by calcium phosphate precipitation. After 48 h the virus containing culture medium was collected and centrifuged for 5 min at 700 x g to remove detached cells and cell debris; then the supernatant was filtered through 0.22 µm filters (Millipore Products, Schwalbach, Germany). The RINm5F cells were infected with the purified viral supernatant for 5-6 h and thereafter the viral supernatant was replaced by fresh medium. Transduced cells were selected for catalase expression by 1 μM blasticidin (Life Technologies, Darmstadt, Germany). Quantification of catalase activity Catalase activity was quantified as previously described [16]. Briefly, whole cell extracts were prepared in 50 mmol/l potassium phosphate buffer (pH 7.8) through sonication on ice with a Braun-Sonic 125 sonifier (Braun, Melsungen, Germany). The homogenates were then centrifuged at 10,000 g and 4°C for 10 min and the protein content of the supernatant was determined by the BCA assay (Thermo Fisher Scientific, Rockford, IL, USA). For the quantification of catalase activity in cell culture medium 2 x 106 cells were seeded in 1.5 ml fully supplemented RPMI-1640 medium on a 6 cm dish. 24 h thereafter the medium was collected and centrifuged at 700 g to remove detached cells and the supernatant was used for the measurement. The catalase activity was measured by ultraviolet spectroscopy, monitoring the decomposition of H2O2 at 240 nm. Peptide-N-glycosidase F digestion, N-ethyl-maleimide treatment and Western blot analysis N-linked glycans were removed from ER-Catalase by peptide-N-glycosidase F treatment of 25 µg cellular total protein with 5 U (PNGase F, Roche Diagnostics, Mannheim, 5

Germany) for 4 h at 37°C in a thermomixer at 1300 rpm. Thereafter digested proteins were subjected to Western blot analysis. For the detection of disulfides in the ER-localized catalase expressing RINm5F cells, the cells were treated for 2 h with or without 20 mM N-ethyl-maleimide (NEM) in PBS at 4°C. For control purposes RINm5F cells expressing ER-Catalase N244 were incubated in the presence of 50 mM DTT for 0.5 h or for 2 h followed by a 1.5 h incubation with 20 mM NEM. Whole cell extracts were prepared in RIPA buffer (Sigma, St. Louis, MO, USA) supplemented with complete protease inhibitor cocktail (Roche Diagnostics, Manheim, Germany). Protein content was determined by the BCA assay. RINm5F cell proteins, 25 µg per lane, were separated by reducing (Peptide-N-glycosidase F digestion) or non-reducing (Nethyl-maleimide and DTT treatment) 10% SDS-PAGE and transferred to polyvinylidene fluoride membranes. Nonspecific binding sites of the membranes were blocked with 5% nonfat dry milk for 1 h at room temperature. Then, the membranes were incubated with anticatalase antibody (sc-34285, Santa Cruz Biotechnology, Santa Cruz, CA, USA; 1:500 dilution) overnight at 4°C. An excess of primary antibody was removed by three washes with washing buffer (PBS, 0.1% Tween 20, 0.1% BSA). Subsequently, the membranes were incubated with a peroxidase-labeled anti-goat antibody (Dianova, Hamburg, Germany; 1:20,000 dilution) at room temperature for 1 h. The protein bands were visualized by chemiluminescence using the ECL detection system (GE Healthcare, Freiburg, Germany). Immunofluorescence staining 24 h after seeding RINm5F cells expressing one of the ER-Catalase variants were on 4 well LabTek chamber slides (Nunc, Roskilde, Denmark), the cells were washed twice with PBS and subsequently fixed with 4% paraformaldehyde at room temperature for 60 min. After washing, the cells were permeabilized and blocked with PBS containing 0.2% Triton X100 and 1% BSA. The cells were incubated with primary antibodies (anti-PDI, ab5484, 6

Abcam, Cambridge, UK, 1:100 dilution; anti-catalase, 100-4151, Rockland Immunochemicals Inc., Limerick, PA, USA, 1:750 dilution) diluted in PBS containing 0.1% Triton X-100 and 0.1% BSA at room temperature for 60 min. Then, the cells were washed with PBS and incubated with specific secondary antibodies (anti-mouse-Alexa Fluor 647, or anti-rabbitAlexa Fluor 488, Jackson ImmunoResearch Laboratories, West Grove, PA, USA; 1:200 dilutions) for 60 min in the dark. Afterwards the cells were washed and the nuclei were counterstained with 300 nM DAPI for 5 min at room temperature. Finally, the cells were washed and mounted with Mowiol/DABCO anti-photobleaching mounting media (Sigma, St. Louis, MO, USA). Stained cells were examined with an Olympus IX81 inverted microscope (Olympus, Hamburg, Germany) and microscopic images were post-processed using AutoDeblur and AutoVisualize (Autoquant Imaging, New York, USA). H2O2 treatment and quantification of cell viability 24 h after seeding of 25,000 cells per well of a 96-well plate, cells were incubated with 50, 100, and 500 µM H2O2 for 2 h in HEPES (20 mmol/l)-supplemented Krebs–Ringer bicarbonate medium with 5 mmol/l glucose. After removal of the H2O2 containing medium the cells were incubated for another 22 h in fresh RPMI 1640 medium and thereafter cell viability was determined by a microplate-based MTT assay (3-(4,5-dimethylthiazol-2-yl)-2,5diphenyl tetrazolium bromide) (Sigma, St. Louis, MO, USA). Real-time quantitative RT-PCR for the quantification of ER stress Total RNA was isolated as previously described [17]. For cDNA synthesis, random hexamers were used to prime the reaction of the RevertAid H- M-MuLV reverse transcriptase (Fermentas, St. Leon-Rot, Germany). QuantiTect SYBR Green technology (Qiagen, Hilden, Germany), which uses a fluorescent dye that only binds double-stranded DNA, was employed. The reactions were performed by using ViiA 7 real-time PCR system (Life Technologies, Karlsruhe, Germany). Samples were denatured at 94°C for 3 min followed by 40 PCR cycles. Each cycle comprised a melting step at 94°C for 30 s, an annealing step at 7

60°C for 30 s, and an extension step at 72°C for 30 s. Optimal parameters for the PCR reactions were empirically defined and the purity and specificity of the amplified PCR product in each experiment was verified by melting curve analysis. All transcripts showed Ctvalues, which were at least 10 Ct-values lower than the blank values. Each PCR amplification was performed in triplicate. Data are expressed as relative gene expression after normalization against the geometric mean of the housekeeping genes Tuba1a, Actb and Ppi with qbasePLUS (Biogazelle, Zulte, Belgium). The primer sequences are listed in Table 2. Statistical analysis Data are presented as mean ± SEM. Statistical analyses were performed using the unpaired two-tailed Student's t test (Graphpad Prism 5.0, Graphpad, San Diego, CA, USA).

Results Catalase mutagenesis and deletion of N-glycosylation motives Our analysis of potential N-glycosylation sites in the human catalase protein using the NetNGlyc 1.0 Server (http://www.cbs.dtu.dk/services/NetNGlyc/) revealed four possible Nglycosylation sites, two of which have a N-glycosylation potential that is greater than the preset 0.5 threshold, strongly indicating that these two sites at position N244 and N439 are utilized for N-glycosylation (Fig. 1A). Using the HomoloGene service of NCBI a very high homology of the core glycosylation motive (N244 and N439) between all analyzed species was observed, indicating an essential function of these amino acids for the catalase protein. Therefore we did not directly substitute N244 and N439, but S246 and T441. For this purpose a homologous region of a species without such a motive was chosen for substitution, namely the Mus musculus sequence for the N244 motive and the Drosophila melanogaster sequence for N439. To avoid a destruction of the catalase structure by a single isolated amino acid exchange, the following seven amino acids behind N244 were also replaced by the Mus musculus sequence. The typical Asn-X-Ser/Thr-motives for N-glycosylation of the human

8

catalase were successfully replaced through Site-Directed Mutagenesis (Fig. 1B and 1C) and verified by sequencing. Quantification of catalase activity after mutagenesis and peroxisomal and ER-specific expression To check whether catalase enzyme activity was affected by the introduced changes in the sequence, all three mutated catalase cDNAs (N244 and N439 single mutants and N244/N439 double mutant) were individually expressed in peroxisomes of RINm5F cells by using a lentiviral vector system. Quantification of the specific enzyme activity after transduction revealed that cells transduced with the N244- or with the N439-catalase cDNA showed an extraordinary high catalase activity of around 950 U/mg protein comparable to that of wild type catalase (1126 U/mg protein) and significant higher than untransfected control cells (16 ± 4 U/mg protein, Fig. 2A). Characterization of the double mutant catalase variant N244/N439 showed that the detectable enzyme activity was less than one third of that of the single mutants (278 ± 17 U/mg protein), indicating that the simultaneous amino acid exchange at both positions resulted in an enzyme activity reduction. However, compared with untransfected control cells the increase of the catalase activity is still highly significant (p<0.001, Fig. 2A). Since all constructed N-glycosylation-negative catalase variants retained enzyme activity these mutated cDNAs were prepared for targeted expression in the ER, using the ERtargeting and ER-retention signal elements of the pCMV/myc/ER vector [12]. Again, RINm5F cells were individually transfected with one of the three catalase constructs using lentiviruses for targeted expression in the ER. Measurement of the enzyme activity of ERspecifically expressed catalase variants showed that only the activity of ER-Catalase N244 (935 ± 51 U/mg protein) was comparable with that quantified in peroxisomes (Fig. 2B). In contrast, the activity of ER-Catalase N439 was reduced to only one fifth of that detectable at peroxisomal expression (Fig. 2B). This activity, however, was comparable to the activity of wild type catalase specifically expressed in the ER (51.5 ± 5.8 U/mg protein). Measurement 9

of catalase activity in the tissue culture medium of ER-Catalase N244 expressing RINm5F cells showed only minimal activity (< 5 U/50 µl medium) and which was not different from untransfected RINm5F cells and RINm5F cells overexpressing catalase in peroxisomes. Thus, RINm5F cells expressing ER-Catalase N244 showed no unspecific secretion of ER-Catalase. This documents clearly the efficiency of the retention signal in the ER-Catalase construct to restrict the expression of catalase to the ER. These findings indicate that the single N-glycosylation at position N244 is responsible for the inactivation of catalase after targeted expression within the ER. Double mutations at position N244 and N439 of the ER-Catalase protein showed an enzyme activity which was also very much lower than that of ER-Catalase N244 (Fig. 2B), most probably due to disturbance of important structures of the catalase molecule leading to reduced enzyme activity or lower molecule stability. N-glycosylation of mutated catalase and disulfide bond formation Treatment of cell lysates with Peptide-N-Glycosidase F (PNGase F) and subsequent Western blot analyses revealed that the molecular weight of ER-Catalase N439 was comparable to that of the wild type ER-Catalase, suggesting the same N-glycosylation degree in both proteins. Indeed, removal of N-glycans through PNGase F treatment reduced the molecular weight of ER-Catalase N439 to that of the size of non-glycosylated catalase protein (Mito-Catalase, Fig. 3A). However, ER-Catalase N244 and N244/439 showed a molecular weight equal to non-glycosylated Mito-Catalase or PNGase F treated wild type ER-Catalase. PNGase F treatment of ER-Catalase N244 and N244/439 resulted in no detectable change of their molecular weight (Fig. 3B), indicating that both catalase variants were non-glycosylated. Besides N-glycosylation the oxidation of normally reduced cysteine residues or the disulfide bond formation through the ER-localized folding machinery could also be responsible for the inactivation of wild type ER-Catalase. However, the analysis of the redox state of all ERCatalase variants by N-ethyl-maleimide treatment and non-reducing SDS polyacrylamide gel 10

electrophoresis showed no obvious and serious changes in the mobility of the investigated ER-Catalase variants (Fig. 3C and Supplementary Fig. S1). Determination of molecular weight and quantification of catalase activities showed that the catalase protein is N-glycosylated at position N244 and that this N-glycosylation is responsible for blunted catalase enzyme activity upon expression in the ER. Furthermore, the redox state of cysteines present in the catalase protein was not substantially altered by the ERspecific folding machinery. ER-specific expression of ER-Catalase N244 in RINm5F cells The subcellular localization of the three different catalase variants was analyzed by immunofluorescent staining. As shown in Fig. 4, co-staining approaches with PDI, an ERspecific expressed protein, and nuclear counterstaining with DAPI proved that the ERtargeted catalase variant N244 was exclusively localized in the ER. This observation indicates that the higher enzyme activity and the non-glycosylated state of ER-Catalase N244 is not the outcome of an incorrect organelle-targeting or expression in another cellular compartment such as cytosol or peroxisomes. Protection of ER-Catalase N244 expressing RINm5F cells against H2O2 induced toxicity To determine whether expression of ER-Catalase N244 protects against H2O2-induced toxicity, we compared the cell viability of control cells with cells expressing ER-Catalase N244 after H2O2 treatment. Even at the lowest concentration of 50 µM H2O2 the viability of the control cells was less than 20% compared with untreated cells, whereas the cells expressing ER-Catalase N244 showed at all used concentrations up to 500 µM H2O2 a viability of 73% or higher (Fig. 5A). The qRT-PCR analysis of the three ER stress markers Atf4, Atf6 and Chop showed in addition that ER-Catalase N244 expressing cells were completely protected against the toxicity of ER stress by 25 or 50 µM H2O2 added exogenously. A 6 or 24 h exposure of ERCatalase N244 expressing cells to H2O2 revealed no significant changes in the gene 11

expression level of all three investigated ER stress genes (Fig. 5B and C). However, in contrast to ER-Catalase N244 expressing cells, the gene expression of Atf4 (5-6 fold increase after 6 h and 3-4,5 fold increase after 24 h) and Chop (12-17 fold increase after 6 h and 6-25 fold increase after 24 h) in untransfected control cells was significantly induced by 25 and 50 µM H2O2, respectively (Fig. 5B and C). Hence, expression of ER-Catalase N244 can detoxify H2O2 effectively and protect cells against H2O2-mediated toxicity. Moreover, through effective inactivation of H2O2 within the ER, ER-Catalase N244 was able to completely prevent the induction of ER stress in these cells.

Discussion With the revealment of the oxidative folding mechanisms in the ER and the understanding of the signaling events initiating the unfolded protein response (UPR), culminating in ER stress, the generation of H2O2 within these processes moved more and more into the research focus. Under physiological conditions H2O2 is produced as a byproduct of the endoplasmic reticulum oxidoreductin 1 (Ero1) during the reoxidation of the protein disulfide isomerase (PDI) in an equimolar manner with each disulfide bond formed by the collaborative action of PDI and ERO-1 [18]. On the other side, an overwhelming protein folding demand and an overstrained protein folding capacity with accumulation of misfolded proteins in the ER lumen was often accompanied with elevated reactive oxygen species (ROS) levels [19, 20]. Therefore, a precise characterization of the ER redox status, of luminal ROS concentrations, especially of H2O2, and effective tools for the manipulation of ER generated ROS levels are essential to pave the way for better understanding of ROS-handling and consequences of inadequate ROS coping by the ER. Detailed characterization of the ER specific GSH/GSSG status has been facilitated in the last years through the establishment of glutaredoxin-fused redox-sensitive green 12

fluorescent protein (roGFP) sensors [21, 22]. However, similar approaches to quantify luminal H2O2 concentrations by use of the H2O2-sensitive biosensor HyPer failed. Most probably the fluorescence signal of HyPer in the ER was generated H2O2 independently by disulfide bond formation in the OxyR domain of the HyPer protein through the ER folding machinery [11, 12]. The other approach to obtain deeper insight into the importance of H2O2 for the ER redox homeostasis is the direct manipulation of luminal H2O2 concentrations. Since the ER resident and H2O2-inactivating enzymes glutathione peroxidase 7 and 8 (GPx7and GPx8) and peroxiredoxin 4 (PRDXIV) participate in oxidative protein folding [23], we expressed in an earlier study catalase in the ER [12]. However, ER-specifically expressed catalase showed only moderate activity due to N-glycosylation. With the generated ERCatalase N244 presented in this study we can offer for the first time a fully ER-located catalase protein with an enzyme activity comparable to that obtained after catalase overexpression in other cellular compartments [13]. Interestingly, the second investigated Nglycosylation motive N439 of the catalase protein was not utilized for N-glycosylation. In an acatalasemic mouse model an asparagine to serine substitution at this position was associated with reduced catalase activity and an unstable tetrameric structure of catalase in the affected animals [24]. N439 is located on the tenth α-helix within the fourth domain of the catalase subunit and therefore N-glycosylation at this position could have steric effects on the formation of the substrate channel leading to the heme group in the tetrameric catalase molecule [25]. In contrast to other enzymes involved in H2O2 metabolism in the ER, like PRDXIV, GPx7 or GPx8 [23], ER-Catalase N244 is definitely not involved physiologically in the formation of disulfide bonds and protein folding. Through its unique reaction mechanism catalase works independently from reducing cofactors and therefore with increasing activity no disturbances of the GSH/GSSG or NADP+/NADPH balances should become evident. In addition, catalase possesses a virtually non-saturable H2O2-inactivation capacity, suited for

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inactivation of high H2O2 concentrations, which have been postulated for cells like pancreatic beta cells with their high secretory activity [4]. With these characteristics the ER-Catalase N244 is a new valuable and unique tool for future experiments to explore the impact of ER-generated H2O2 for the redox-dependent synthesis of disulfide bonds [3], for the deleterious effects of misfolded protein aggregation or for the induction of ER-stress signaling leading to cellular dysfunction and death [8].

Acknowledgments This work has been supported by a grant from the European Union (BetaBAT, Grant Agreement 277713) in the Framework Programme 7.

References [1] Tu, B. P.; Weissman, J. S. Oxidative protein folding in eukaryotes: mechanisms and consequences. J. Cell Biol. 164:341-346; 2004. [2] Ramming, T.; Appenzeller-Herzog, C. The physiological functions of mammalian endoplasmic oxidoreductin 1: on disulfides and more. Antioxid. Redox Signaling 16:11091118; 2012. [3] Ramming, T.; Appenzeller-Herzog, C. Destroy and exploit: catalyzed removal of hydroperoxides from the endoplasmic reticulum. Int. J. Cell Biol. 2013:180906; 2013. [4] Shimizu, Y.; Hendershot, L. M. Oxidative folding: cellular strategies for dealing with the resultant equimolar production of reactive oxygen species. Antioxid. Redox Signaling 11:2317-2331; 2009. [5] Margittai, E.; Low, P.; Stiller, I.; Greco, A.; Garcia-Manteiga, J. M.; Pengo, N.; Benedetti, A.; Sitia, R.; Banhegyi, G. Production of H(2)O(2) in the endoplasmic reticulum promotes in vivo disulfide bond formation. Antioxid. Redox Signaling 16:1088-1099; 2012. [6] Bhandary, B.; Marahatta, A.; Kim, H. R.; Chae, H. J. An involvement of oxidative stress in endoplasmic reticulum stress and its associated diseases. Int. J. Mol. Sci. 14:434-456; 2012. [7] Malhotra, J. D.; Kaufman, R. J. Endoplasmic reticulum stress and oxidative stress: a vicious cycle or a double-edged sword? Antioxid. Redox Signaling 9:2277-2293; 2007. [8] Malhotra, J. D.; Miao, H.; Zhang, K.; Wolfson, A.; Pennathur, S.; Pipe, S. W.; Kaufman, R. J. Antioxidants reduce endoplasmic reticulum stress and improve protein secretion. Proc. Natl. Acad. Sci. USA 105:18525-18530; 2008. [9] Cao, S. S.; Kaufman, R. J. Endoplasmic Reticulum Stress and Oxidative Stress in Cell Fate Decision and Human Disease. Antioxid. Redox Signaling; 2014. [10] Nabeshima, A.; Yamada, S.; Guo, X.; Tanimoto, A.; Wang, K. Y.; Shimajiri, S.; Kimura, S.; Tasaki, T.; Noguchi, H.; Kitada, S.; Watanabe, T.; Fujii, J.; Kohno, K.; Sasaguri, Y. Peroxiredoxin 4 protects against nonalcoholic steatohepatitis and type 2 diabetes in a nongenetic mouse model. Antioxid. Redox Signaling 19:1983-1998; 2013. 14

[11] Malinouski, M.; Zhou, Y.; Belousov, V. V.; Hatfield, D. L.; Gladyshev, V. N. Hydrogen peroxide probes directed to different cellular compartments. PLoS One 6:e14564; 2011. [12] Mehmeti, I.; Lortz, S.; Lenzen, S. The H(2)O(2)-sensitive HyPer protein targeted to the endoplasmic reticulum as a mirror of the oxidizing thiol-disulfide milieu. Free Radic. Biol. Med. 53:1451-1458; 2012. [13] Gurgul, E.; Lortz, S.; Tiedge, M.; Jörns, A.; Lenzen, S. Mitochondrial catalase overexpression protects insulin-producing cells against toxicity of reactive oxygen species and proinflammatory cytokines. Diabetes 53:2271-2280; 2004. [14] Lortz, S.; Tiedge, M.; Nachtwey, T.; Karlsen, A. E.; Nerup, J.; Lenzen, S. Protection of insulin-producing RINm5F cells against cytokine-mediated toxicity through overexpression of antioxidant enzymes. Diabetes 49:1123-1130; 2000. [15] Zufferey, R.; Dull, T.; Mandel, R. J.; Bukovsky, A.; Quiroz, D.; Naldini, L.; Trono, D. Self-inactivating lentivirus vector for safe and efficient in vivo gene delivery. J. Virol. 72:9873-9880; 1998. [16] Tiedge, M.; Lortz, S.; Munday, R.; Lenzen, S. Complementary action of antioxidant enzymes in the protection of bioengineered insulin-producing RINm5F cells against the toxicity of reactive oxygen species. Diabetes 47:1578-1585; 1998. [17] Chomczynski, P.; Sacchi, N. Single-step method of RNA isolation by acid guanidinium thiocyanate-phenol-chloroform extraction. Anal Biochem 162:156-159; 1987. [18] Gross, E.; Sevier, C. S.; Heldman, N.; Vitu, E.; Bentzur, M.; Kaiser, C. A.; Thorpe, C.; Fass, D. Generating disulfides enzymatically: reaction products and electron acceptors of the endoplasmic reticulum thiol oxidase Ero1p. Proc. Natl. Acad. Sci. USA 103:299-304; 2006. [19] Haynes, C. M.; Titus, E. A.; Cooper, A. A. Degradation of misfolded proteins prevents ER-derived oxidative stress and cell death. Mol. Cell 15:767-776; 2004. [20] Song, B.; Scheuner, D.; Ron, D.; Pennathur, S.; Kaufman, R. J. Chop deletion reduces oxidative stress, improves beta cell function, and promotes cell survival in multiple mouse models of diabetes. J. Clin. Invest. 118:3378-3389; 2008. [21] Birk, J.; Ramming, T.; Odermatt, A.; Appenzeller-Herzog, C. Green fluorescent protein-based monitoring of endoplasmic reticulum redox poise. Front. Genet. 4:108; 2013. [22] van Lith, M.; Tiwari, S.; Pediani, J.; Milligan, G.; Bulleid, N. J. Real-time monitoring of redox changes in the mammalian endoplasmic reticulum. J. Cell Sci. 124:2349-2356; 2011. [23] Ramming, T.; Hansen, H. G.; Nagata, K.; Ellgaard, L.; Appenzeller-Herzog, C. GPx8 peroxidase prevents leakage of H2O2 from the endoplasmic reticulum. Free Radic. Biol. Med. 70:106-116; 2014. [24] Wang, D. H.; Tsutsui, K.; Sano, K.; Masuoka, N.; Kira, S. cDNA cloning and expression of mutant catalase from the hypocatalasemic mouse: comparison with the acatalasemic mutant. Biochim. Biophys. Acta 1522:217-220; 2001. [25] Fita, I.; Rossmann, M. G. The active center of catalase. J. Mol. Biol. 185:21-37; 1985.

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Figure 1

N-glycosylation potential

A 0.75

N244 N-glycosylation threshold

0.5 0.25

N481

N148

0 0

B

N439

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244 Asn

Leu

200 300 400 Amino acid position in human catalase protein

Ser

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Ala

Arg

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Gln

5´-AAC CTT TCT GTT GAA GAT GCG GCG AGA CTT TCC CAG-3´ mutagenesis of N244 glycosylation site 244 Asn

Leu

Pro

Val

Gly

Glu

Ala

Gly

Arg

Leu

Ala

Gln

5´-AAC CTT CCC GTT GGA GAG GCG GGG AGA CTT GCC CAG-3´

C

439 Asn

V al

Thr

Gln

5´-AAC GTT ACT CAG-3´ mutagenesis of N439 glycosylation site 439 Asn

Phe

Gly

Gln

5´-AAC TTT GGT CAG-3´

500

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Table 1

Table 1 Primers used for the deletion of the N-glycosylation motives at N244 and N439 of the human catalase and subcloning for eukaryotic expression Primers

Sequence

1. Deletion of N244 by site-directed mutagenesis

2.

frw

5´-GCGGGGAGACTTGCCCAGGAAGATCCT-3´

rev

5´-CTCTCCAACGGGAAGGTTTTTGATGCCCTG-3´

Deletion of N439 by site-directed mutagenesis frw

5´-GGTCAGGTGCGGGCATTCTAT-3´

rev

5´-AAAGTTATCATCATTGGCAGTGTTGAA-3´

3. Subcloning of the mutated catalase cDNAs into the pLenti6.3/V5-MCS plasmid frw

5´-ATGGCTGACGACAGCCGGGAT-3´

rev

5´-TCACAGATTTGCCTTCTCCCTT-3´

4. Subcloning of the mutated catalase cDNAs into the pCMV/myc/ER plasmid frw

5´-ATACTGCAGATGGCTGACAGCCGGGAT-3´

rev

5´-ATGCGGCCGCCAGATTTGCCTTCTCCCTTGC-3´

5. Transfer of the mutated catalase, ER-targeted cDNAs into the pLenti6.3/V5-MCS plasmid frw

5´-ATGGGATGGAGCTGTATCATCC-3´

rev

5´-CTACAGCTCGTCCTTCTCGCTT-3´

Primers for subcloning of the mutated catalase cDNAs into the pCMV/myc/ER plasmid: restriction enzyme recognition sites in italic, complementary sequences are underlined

1

Table 2

Table 2 Primers used for the quantification of ER-stress genes by qRT-PCR Primers

Sequence

Atf4 (activating transcription factor 4) frw

5´- TCCTGAACAGCGAAGTGTTG -3´

rev

5´- CGCACTGACCACTCTGTTTC -3´

Atf6 (activating transcription factor 4) frw

5´- GTCCCAGATATTAATCACGGA -3´

rev

5´- TATCATACGTTGCTGTCTCCTT -3´

Chop (C/EBP homologous protein) frw

5´- CCAGCAGAGGTCACAAGCAC -3´

rev

5´- CGCACTGACCACTCTGTTTC -3´

Tuba1a (α-Tubulin) frw

5´- ACGTGAGACGTACATCCCAAACTCA -3´

rev

5´- CTCCCAGCAGGCATTGCCCA -3´

Actb (β-Actin) frw

5´- GAACACGGCATTGTAACCAACTGG -3´

rev

5´- GGCCACACGCAGCTCATTGTA -3´

Ppia (peptidylprolyl isomerase A or cyclophilin A frw

5´- CATCTGCACTGCCAAGACTGA -3´

rev

5´- TGCAATCCAGCTAGGCATG -3´

1