Seeding of aSyn can be recapitulated in cellular systems of aSyn aggregation; however, the mechanism of aSyn seeding and its regulation are not well understood. We developed an mEos-based aSyn seeding assay and performed saturation mutagenesis to identify with single-residue resolution positive and negative regulators of aSyn aggregation. We not only found the core regions that govern aSyn aggregation but also identified mutants outside of the core that enhance aggregation. We identified local structure within the N terminus of aSyn that hinders the fibrillization propensity of its aggregation-prone core. Based on the screen, we designed a minimal aSyn fragment that shows a ~4-fold enhancement in seeding activity and enabled discrimination of synucleinopathies. Our study expands the basic knowledge of aSyn aggregation and advances the design of cellular systems of aSyn aggregation to diagnose synucleinopathies based on protein conformation. Mutagenesis of α-synuclein coupled with crosslinking and NMR provides insights into how local structure modulates aggregation. We followed up these experiments with truncation analyses, cross-linking with mass spectrometry (MS), and nuclear magnetic resonance (NMR) spectroscopy. These experiments provided insight into how local structure controls aSyn propensity for aggregation. We propose a model in which the N terminus of aSyn modulates the fibrillization propensity of its aggregation-prone core.We sought to create a saturation screen that would rely on intracellular expression of one mutant aSyn per cell. Thus, we developed an aSyn(A53T)-mEos stable cell line similar to that which we have previously reported  We used sequential degenerate codon primers to construct a saturation library of single-point mutants of aSyn at every position. The pooled library encoded 2780 aSyn variants C-terminally fused to the mEos tag (aSyn-mEos). Deep sequencing of the input library revealed consistent distribution of mutations across the length of the gene and the absence of multiple mutations within the same construct . We thenTo identify which mutants decreased or increased aSyn seeding propensity, we calculated the relative enrichment (RE) for each variant within sorted populations . This reTo test these mutations for seeding potential, we picked 16 de-enriched and 13 enriched mutants, cloned them upstream of cyan or yellow fluorescent protein (aSyn-CFP or aSyn-YFP), and created high-expressing polyclonal biosensor lines. The literature suggests that these tags do not have any substantial effect on aSyn aggregation . Each muTo systematically assess aSyn domains most sensitive to any substitution, we calculated the median RE across all mutations for each residue (missing data and data for stop codons were excluded) and applied a variance-based Otsu thresholding method . This reThe mutagenesis predicted an important role for the aSyn core in seeded aggregation. Thus, we tested the influence of N- and C-terminal sequences on aggregation by truncating aSyn-CFP/YFP fusions and stably overexpressing them in HEK293T cells . DeletioThe prion hypothesis predicts differential seeding efficiencies of strains based on the amino acid composition of the monomer. To test this idea for aSyn, we purified sarkosyl-insoluble fibrils from MSA, PD, and LBD brain tissue and transfected a panel of various biosensors . MSA fibDeletion analysis suggested that the N terminus of aSyn might modulate aggregation. Cross-linking with MS (XL-MS) has proven useful not only to model protein assemblies but also to infer changes in conformations of intrinsically disordered proteins such as tau  aggregated more efficiently . We hypo15N-labeled recombinant aSyn(WT) and aSyn(E13K) by NMR spectroscopy. We collected heteronuclear single quantum coherence (HSQC) of aSyn(WT) and aSyn(E13K) (Models based on XL-MS predicted that aSyn(E13K) has a different conformation within the N terminus. To test that hypothesis, we analyzed yn(E13K)  and usedWe have used saturation mutagenesis, deletion analysis, XL-MS, and NMR spectroscopy to identify the sequence determinants of aSyn aggregation. We began by identifying aSyn residues enriched or de-enriched in biosensor cells seeded with recombinant aSyn fibrils. A selected number of mutants were retested and confirmed, validating the primary screen and allowing identification of residues important for aggregation. These mapped onto regions predicted to be important based on cryo-EM of multiple aSyn fibrils. Deletion analysis revealed an inhibitory role for both the N and C termini and allowed us to augment the sensitivity of aSyn biosensors by at least fourfold. In addition, we identified mutants that added discriminatory power to aSyn biosensors. XL-MS suggested that the N terminus has local structure, and the E13K mutation suggested that disruption of local electrostatic interactions might underlie increased seeding. We tested this idea using NMR spectroscopy. This mutation perturbed local structure in the region beyond that expected in the immediate vicinity of the mutated residue. We conclude that transient local structure in the aSyn N terminus mediates interactions that inhibit aggregation. These studies may enable understanding of cellular controls of aSyn aggregation and provide new tools for the analysis of synucleinopathies.et al.  . The amit-distributed stochastic neighbor embedding (t-SNE)  that may inhibit aggregation. This was destabilized in case of aSyn(A53T) and almost eliminated in aSyn(E13K). XL-MS also suggested that these variants change the exposure of the hydrophobic core, which would presumably be masked by intramolecular folding. Together, these findings indicated that regulation of aSyn aggregation may hinge on exposure of its hydrophobic core , frozen, and stored at −80°C until use. The resuspended pellets were quickly thawed, and the cells were lysed with a French press at 10,000 to 15,000 psi in a 5-min cycle until the liquid became transparent. The cell solution was centrifuged at 15,000g for 30 min at 4°C. Then, the supernatant was passed through a 22-μm cutoff filter. For further protein purification, we packed a column with 40 ml of slurry of chitin beads . The resin was washed with 200 ml of the lysis buffer. Next, we loaded the filtered supernatant into the prepared chitin column at 0.5 to 1 ml/min, ensuring that the resin was never dry. Then, we added 50 ml of cleavage buffer , quickly collecting 30 ml of flow-through (discarded) and incubating chitin beads with the rest of the cleavage buffer for 48 hours in darkness at 4°C. Then, we collected the flow-through containing eluted aSyn. Next, we added another 40 ml of cleavage buffer and incubated chitin beads for another 48 hours. Flow-through was combined and dialyzed twice overnight at 4°C into 5 liters of 10 mM ammonium bicarbonate using dialysis cassettes with 10-kDa protein molecular weight cutoff with a delicate mixing. After dialysis, the sample was passed through a 22-μm cutoff filter, flash-frozen, and stored at −80°C. Next, we lyophilized the protein sample and stored it at −80°C until further use. The sample was then dissolved in 100% trifluoroacetic acid (TFA) and diluted with water to 50% TFA, and resolved using a reversed-phase column . The protein was eluted with a gradient of increasing concentration of acetonitrile with constant 0.05% TFA. We combined collected fractions containing aSyn monomer and dialyzed them twice overnight at 4°C into 5 liters of 10 mM ammonium bicarbonate. The dialyzed sample was filtered with a 22-μm cutoff filter and flash-frozen. The sample was then lyophilized and stored at −80°C.We designed aSyn gene blocks with sequences of interest, which were synthesized by Integrated DNA Technologies (IDT). We used present NdeI and SpeI restriction sites to subclone these genes into ampicillin-resistant pTXB1 plasmid (NEB N6707S) by ligation. Lyophilized WT aSyn monomer was resuspended in a buffer containing 20 mM tris, 100 mM NaCl at pH 8.0 to a concentration of 2 mg/ml (138 μM). The sample was incubated at 37°C in a Thermomixer (Eppendorf) with shaking at 1000 rpm for 7 days. The sample was stored at 4°C.A total of 4.5 μl of sample [0.1 mg/ml of recombinant aSyn(WT) fibrils or 0.2 mg/ml of brain-derived sarkosyl-insoluble material] was applied to a glow-discharged grid  and incubated for 1 min at room temperature (RT). After that, the grid was blotted and quickly washed with 4.5 μl of water. Next, the grid was stained with 4.5 μl of 2% uranyl acetate for 45 s, blotted, and dried at RT. Images were acquired using a Tecnai G2 spirit transmission electron microscope , serial number D1067, equipped with a LaB6 source at 120 kV and a Gatan ultrascan charge-coupled device camera.We ordered 35– to 61–nucleotide (nt)–long primer pairs with degenerated codons NNS in the middle of each primer , flanked by 10 to 48 nt matching the WT nucleotide aSyn sequence (IDT). We individually cloned 139 saturated mutants for each residue and pooled plasmids with all the mutants to one tube in a mass ratio of 1:1. Cloning of each mutated residue consisted of seven steps.Two halves of aSyn nucleotide sequence were amplified using two primer pairs . For each reaction, we added 0.5 μl of iProof polymerase , 1 μl of 10 mM deoxynucleotide triphosphate (dNTP) , 1 μl of 100% dimethyl sulfoxide (DMSO), 1 μl of template , 1 μl of 10 μM primer mixture (forward + reverse) at a molar ratio of 1:1, 10 μl of 5× iProof buffer, and 36 μl of nuclease-free water.Polymerase chain reaction (PCR) program:We mixed two halves of aSyn gene from the previous step to build one fragment (insert). For each reaction, we added 0.5 μl of iProof polymerase , 1 μl of 10 mM dNTP , 1 μl of 100% DMSO, 1 μl of template (0.5 μl + 0.5 μl of each fragment from the extension PCR), 1 μl of 10 μM primer mixture at a molar ratio of 1:1 (forward + reverse vector primers), 10 μl of 5× iProof buffer, and 36 μl of nuclease-free water.PCR program:Reaction mixtures after the overlap PCR, containing inserts for the following steps, were purified with a DNA purification kit (Zymo Research Genomic DNA Clean & Concentrator-10 D4011), eluting DNA with 43 μl of nuclease-free water.We digested purified inserts with an Esp3I restriction enzyme. For each reaction, we added 42.5 μl of purified insert from step (iii), 2.5 μl of 20 mM DTT, 5 μl of 10× Fast Digest buffer, and 1 μl of Esp3I restriction enzyme. The reactions were incubated at 37°C for 1 hour, then at 65°C for 10 min, and at 4°C until later use.We ligated digested inserts with a digested target vector containing a cytomegalovirus (CMV) promoter, an mEos C-terminal tag, and ampicillin resistance. For each reaction, we added 2 μl of 10× T4 DNA ligase buffer, 50 ng of vector DNA, 9 μl of digested insert DNA from step (iv), and nuclease-free water to fill it up to 20 μl. Next, we added 1.1 μl of T4 DNA ligase, the mixture was incubated at 16°C for 18 hours and then at 65°C for 10 min, and the temperature was lowered to 4°C until further use.E. coli  with a Bio-Rad E. coli Pulser  and 0.1-cm gap cuvettes . Then, we immediately added 980 μl of super optimal medium with catabolic repressor (SOC) medium and incubated the mixture at 37°C with 220 rpm shaking for 1 hour. Then, the electroporated bacteria were plated on LB agar plates with ampicillin at 100 μg/ml using glass beads and incubated overnight at 37°C.We electroporated 1 μl of each ligation reaction into 19.5 μl of 10-Beta electrocompetent g for 10 min. Plasmids were purified from the pelleted bacteria with a Qiagen kit , eluted with 50 μl of nuclease-free water, and stored at −20°C.We harvested all colonies from each plate by scrapping them with 10 ml of LB medium with 100 μg/ml of ampicillin. Bacteria were spun down at 4°C, 3901g for 5 min. Polybrene (Sigma-Aldrich Polybrene Infection TR-1002-G) and Hepes were added to the supernatant to a final concentration of 4 μg/ml and 50 nM, respectively. The mixture was passed through a 22-μm filter, aliquoted, and stored at −80°C.Lentivirus was produced in HEK293T cells [available from the American Type Culture Collection (ATCC)] through a triple transfection with a packaging plasmid Gag-Pol and an envelope plasmid pVSV-g together with a lentiviral transfer vector carrying an aSyn gene variant (or a pooled aSyn library). HEK293T cells were plated at 400,000 cells per well in six-well plates and incubated at 37°C. The next day, 800 ng of Gag-pol, 200 ng of pVSV-g, and 1000 ng of lentiviral transfer vector were mixed with 7.5 μl of TransIT-293 (Mirus) and 92.5 μl of reduced serum medium  and incubated at RT for 15 min. The cell medium was changed to 1 ml of DMEM  with 3% fetal bovine serum . The transfection mixture was added dropwise to the wells and incubated at 37°C. After 6 hours, the medium was changed to fresh 1.5 ml of DMEM with 3% FBS. After 48 hours, the medium was collected, stored at 4°C, and replaced with fresh 1.5 ml of DMEM with 3% FBS. The next day, the medium was collected, combined with the previous one, and centrifuged at 10006 cells per plate in 9 ml of compete cell medium , and incubated at 37°C. The next day, 25 μl of aSyn library lentivirus diluted with 975 μl of Opti-MEM was added dropwise to each of the five dishes with HEK293T cells (resulting in a multiplicity of infection below 0.3). Cells were incubated at 37°C for 48 hours. Then, the cell medium was replaced (10 ml). aSyn fibrils were sonicated (or buffer for a negative control) at an amplitude of 65 for 5 min (1 min ON/1 min OFF) at 4°C in a water bath using a Q700 Sonicator (QSonica). Next, in one tube, we diluted 100 μl of sonicated aSyn fibrils or buffer with 650 μl of Opti-MEM. In the second tube, we mixed 50 μl of Lipofectamine  with 700 μl of Opti-MEM and incubated the mixture at RT for 5 min. After that, both mixtures were combined and incubated at RT for 30 min. The total of 1.5 ml of prepared treatment (with fibrils or Lipofectamine only) was added to each of three plates with HEK293T cells. The fibrils were diluted to a final concentration of 200 nM. Cells were incubated with a treatment at 37°C for 48 hours. Next, the cells were harvested by removal of cell medium, treatment with 2 ml of 0.05% trypsin , and incubation at 37°C for 2 min. 8 ml of complete medium was added to stop trypsinization. The cells were centrifuged at 1000 rpm for 5 min and fixed with 5 ml of 2% paraformaldehyde (PFA) in Dulbecco’s phosphate-buffered saline  at RT for 10 min. The cells were centrifuged at 1000 rpm for 5 min, resuspended in 1 ml of DPBS, and diluted by adding another 4 ml of DPBS and placed into 6-cm dishes. The cells were treated with UV light (~405 nm) for 1 hour to convert green mEos to red. The cells were centrifuged at 1000 rpm for 5 min, resuspended in 1 ml of DPBS, passed through 50-μm filters, and diluted in 5 ml of DPBS. Moreover, we prepared fixed HEK293T cells with aSyn A53T-dsRed, aSyn A53T-mEos, and WT HEK293T cells for FACS compensation controls. After cell preparation, the cells were sorted at 4°C to DPBS into FRET-positive and FRET-negative gate using FACSAria II SORP with the four-way purity sorting method. The gating strategy was followed as previously described (g for 10 min. Each pellet was resuspended in 500 μl of DPBS and stored at 4°C.HEK293T cells (available from ATCC) were plated into five 10-cm dishes, 1.5 × 106 cells per sample) in 500 μl of DPBS. Samples were incubated overnight at 55°C with 550 rpm shaking. After that, samples were heated up at 95°C for 10 min and then cooled at 4°C. Next, we added 2 μl of ribonuclease A  to each sample, vortexed, and incubated at 37°C with 550 rpm shaking in a Thermomixer (Eppendorf) for 1 hour. Cell solutions were added to pelleted phase lock gel (PLG) tubes . We added 500 μl of buffered phenol (pH 7.9)  to each sample, gently mixed, and centrifuged at 16,000g at RT for 5 min. Next, we added 500 μl of phenol-chloroform-isoamylalcohol 25:24:1 (v/v) , gently mixed, and centrifuged at 16,000g at RT for 5 min. Upper aqueous phases were transferred into centrifuged PLG tubes, and then 500 μl of chloroform  was added to each tube, gently mixed, and centrifuged at 16,000g at RT for 5 min. Next, upper aqueous phases were transferred into 2-ml tubes with 1.3 μl of glycoblue  and gently mixed. We added 1 ml of 100% ethanol to each sample, gently mixed, and incubated at −80°C for 1 hour. After that, samples were centrifuged at 15,000g at 4°C for 15 min. Pelleted DNA was washed with 1 ml of 75% ethanol (stored at −20°C) and centrifuged at 15,000g at 4°C for 15 min. The supernatants were aspirated, and DNA pellets were dried for several minutes. We added 50 μl of nuclease-free water and let DNA solubilize at 4°C overnight and left it at 4°C until further use.We added 500 μl of lysis buffer  to each sample containing cell pellets ; in the case of FRET-negative cells, we used 1.6 μg of genomic DNA in the first PCR. All used primers were synthesized by IDT.For each reaction, we mixed 0.5 μl of iProof polymerase , 1 μl of 10 mM dNTP , 1 μl of 100% DMSO, template (genomic DNA), 1 μl of 10 μM mixed primers (forward and reverse) at a molar ratio of 1:1, 10 μl of 5× iProof buffer, and nuclease-free water with a total volume of 50 μl. The polymerase was added by the end of the first step of PCR .PCR program:*temperature decreased by 1°C per cycle.Used primers:Forward:1–4GCTAGCGCCGCCACCATGTGACTGGAGTTCAGACGTGtgctcttccgatctNNNNN(N)Reverse:1–4GCAGCGGAGCCAGCAGAACCACTCTTTCCCTACACGACgctcttccgatctNNNNN(N)The PCR product was run on a 3% agarose gel  at 80 V for 6 hours. The 532– to 538–base pair (bp) PCR product was cut out from the gel and purified with a purification kit . The eluted product was purified with a DNA purification kit (Zymo Research Genomic DNA Clean and Concentrator-10 D4011), eluting DNA with 10.5 μl of nuclease-free water.For each reaction, we mixed 0.5 μl of iProof polymerase , 1 μl of 10 mM dNTP , 1 μl of 100% DMSO, 6 ng of template (purified product from step 2), 1 μl of 10 μM mixed primers (forward and reverse) at a molar ratio of 1:1, 10 μl of 5× iProof buffer, and nuclease-free water with a total volume of 50 μl. The polymerase was added by the end of first step of PCR .PCR program:We used combination of total of eight following primers for each sample:Forward:CAAGCAGAAGACGGCATACGAGATcgagtaatGTGACTGGAGTTCAGACGTGCAAGCAGAAGACGGCATACGAGATtctccggaGTGACTGGAGTTCAGACGTGCAAGCAGAAGACGGCATACGAGATaatgagcgGTGACTGGAGTTCAGACGTGCAAGCAGAAGACGGCATACGAGATggaatctcGTGACTGGAGTTCAGACGTGReverse:AATGATACGGCGACCACCGAGATCTACACtatagcctACACTCTTTCCCTACACGACAATGATACGGCGACCACCGAGATCTACACatagaggcACACTCTTTCCCTACACGACAATGATACGGCGACCACCGAGATCTACACcctatcctACACTCTTTCCCTACACGACAATGATACGGCGACCACCGAGATCTACACggctctgaACACTCTTTCCCTACACGACThe PCR product was run on a 3% agarose gel (Invitrogen UltraPure Agarose-1000 16550100) at 80 V for 7 hours. The 603- to 609-bp PCR product was cut out from the gel and purified with a purification kit (Promega Wizard SV Gel and PCR Clean-Up System A9281). The eluted product was purified with a DNA purification kit (Zymo Research Genomic DNA Clean & Concentrator-10 D4011), eluting DNA with 25 μl of nuclease-free water.Deep sequencing was performed at the UTSW Next Generation Sequencing Core, using Illumina MiSeq v2 (500-cycle) Reagent Kit MS-102-2003. The 4 nM DNA sample was denatured with 0.2 N NaOH at 1:1 (v/v), diluted to 8 pM, and spiked with a denatured PhiX control library.iyf is the frequency of protein i in sample y, and the reference sequence (WT) is the WT aSyn protein sequence. This RE value indicates the fold change of abundance for each protein variant relative to WT.Paired-end reads were joined using NGmerge .To identify which residues had strong inhibitory signal, we took the median signal across all mutations  for all residues and applied the Otsu threshold to these data. All residues with a median value equal to or less than the threshold (−1.2152) were classified as having strong inhibitory signal for aggregation.To evaluate the relationship of different mutations across all residues, we applied t-SNE dimensionality reduction to the mutagenesis data. Each residue’s 20 mutations  were used as input, t-SNE was performed with a perplexity value of 30, and Euclidean distance was used as the distance metric.We designed aSyn gene blocks either WT or containing mutations of interest, which were synthesized by IDT. We subcloned these genes by ligation into ampicillin-resistant vectors with a CMV promoter in variants containing CFP or YFP C-terminal tags. All aSyn biosensor variants were engineered through a transduction of HEK293T cells (available from ATCC) with a lentivirus carrying an aSyn gene variant under a CMV promoter.HEK293T cells were plated at 200,000 cells per well in 2 ml of complete DMEM in six-well plates. Two hundred fifty microliters of the lentivirus with a CFP variant and 250 μl with a YFP variant were added dropwise to the well. The cells were incubated at 37°C and maintained until the full expression of CFP and YFP was obtained. Then, the cells with aSyn variants were sorted using FACS into double positive for CFP and YFP single cells or polyclonal cell lines . Cells were grown, tested, and stored in liquid nitrogen.Biosensor cells were plated at 25,000 cells per well in 130 μl of complete DMEM in a 96-well plate and incubated at 37°C. After 24 hours, recombinant full-length WT aSyn fibrils were diluted in a buffer [20 mM tris (pH 8.0) and 100 mM chloride sodium] and sonicated at an amplitude of 65 for 5 min (1 min ON and 1 min OFF) using a Q700 Sonicator (QSonica). 1 μl of sonicated fibrils (or buffer for a negative control) were diluted with 9 μl of Opti-MEM. 0.6 μl of Lipofectamine 2000 was mixed with 9.4 μl of Opti-MEM and incubated for 5 min at RT. The diluted fibrils were mixed with Lipofectamine and incubated for 30 min at RT. Twenty microliters was added dropwise to each well. Each treatment was performed in triplicate. Cells were incubated at 37°C for 48 hours. The medium was removed and 50 μl of 0.05% trypsin was added to each well and incubated for 2 min at 37°C. 100 μl of 2% PFA in DPBS was added to each well, mixed, and incubated for 10 min at RT. Plates were centrifuged at 1000 rpm for 5 min. The supernatant was removed, and cells were resuspended in 130 μl of DPBS for analysis by flow cytometry. FRET-positive cells were quantified by flow cytometry using the gating strategy described as previously , 0.15 M NaCl, 10% sucrose, and 0.2% sarkosyl] and centrifuged for 30 min at 166,000g, 20°C. The supernatant was discarded, and the pellet was washed twice with PBS, followed by centrifugation. Pellets were resuspended in PBS. In case of low solubility, the pellets were sonicated at 4°C in a water bath up to 20 min at an amplitude of 65 (1-min ON and 1-min OFF) using a Q700 Sonicator (QSonica). Protein concentrations were measured with micro-BCA protein assay kit . Samples were flash-frozen and stored at −80°C.aSyn fibrils were extracted from tissue following the protocol published by Schweighauser Fifteen micrograms of brain-derived sarkosyl-insoluble material was mixed with sample buffer  supplemented with β-mercaptoethanol and boiled for 6 min. Samples and a protein MW ladder  were loaded onto a NuPage bis-tris polyacrylamide (4 to 12% gradient) gel . Electrophoresis was performed in MES-SDS buffer  for 1 hour 10 min at 150 V. Protein was transferred from the gel into a polyvinylidene difluoride (PVDF) membrane  at 20 V for 50 min using a Bio-Rad Trans-blot semi-dry transfer apparatus. The membrane was blocked in 5% nonfat milk  in TBS  with 0.1% Tween-20 (TBST) for 1 hour at RT with agitation. Then, the membrane was incubated with an anti-aSyn rabbit antibody targeting C-terminal residues 118 to 123  diluted 1:1000 in TBST with 5% milk overnight at 4°C with agitation. Then, the membrane was washed with TBST and incubated with a secondary antibody against rabbit immunoglobulin G (IgG)  diluted 1:5000 in TBST with 5% milk for 1 hour at RT with agitation. The membrane was washed with TBST and then incubated for 2 min with a chemiluminescence detection reagent .Lyophilized aSyn was resuspended in buffer containing 20 mM Na-Hepes and 100 mM NaCl (pH 8.0) and brought to 0.4 mg/ml. Thirty microliters of DMTMM (120 mg/ml) was added to 270 μl of protein solution and incubated at 37°C with 750 rpm shaking in a Thermomixer (Eppendorf) for 10 min. The reaction was quenched by adding 60 μl of 1 M ammonium bicarbonate and incubated at 37°C with 750 rpm shaking in a Thermomixer for 30 min. Next, the samples were flash-frozen and stored at −80°C.We added Laemmli buffer to samples and boiled at 100°C for 5 min. The samples were resolved on a 1.5-mm bis-tris polyacrylamide (4 to 12% gradient) gel  at 150 V for 1 hour and 10 min and stained with Coomassie G-250  overnight at RT with mild rotatory shaking. The next day, gels were washed with water for 2 hours at RT with mild rotatory shaking. Bands with monomeric aSyn (~14 kDa) were cut out and divided into small pieces and transferred into 1.5-ml tubes. Gel pieces were washed twice with 200 μl of water for 10 min at RT with 600 rpm shaking in a Thermomixer (Eppendorf). Gel pieces were sonicated six times in acetonitrile/50 mM ammonium bicarbonate in a ratio of 2:3 (v/v) at 65 A, 5 min each cycle  at 4°C in a water bath, until gel pieces became transparent. Gel pieces were then washed with 200 μl of 100% acetonitrile for 5 min at RT with 600 rpm shaking in a Thermomixer. Acetonitrile was removed, and residual liquid was vacuumed for 5 min in a freeze-dryer device (Labconco). Then, we added 150 μl of 10 mM DTT in 25 mM ammonium bicarbonate, and samples were incubated for 1 hour at 56°C with 600 rpm shaking in a Thermomixer. The tubes were cooled down, and solution was switched to 150 μl of 55 mM iodoacetamide in 25 mM ammonium bicarbonate. Samples were incubated for 45 min at RT in darkness. Next, gel pieces were washed with 200 μl of 25 mM ammonium bicarbonate, then 200 μl of 50% acetonitrile, and last 200 μl of 100% acetonitrile. After removing the last solution, samples were vacuumed for 5 min. We added 200 μl of 50 mM ammonium bicarbonate to each tube with gel pieces and 1.5 μl of MS-grade trypsin  in water. The samples were incubated overnight (15 to 17 hours) at 37°C with 600 rpm shaking in a Thermomixer. The next day, the reaction was stopped by adding 0.6 μl of formic acid to each tube (until pH dropped to ~3). The samples were flash-frozen and stored at −80°C. Later, the samples were lyophilized and stored at −80°C.m/z) 350 to 1600, charge state 3+ and above] from a preview scan and subjecting them to collision-induced dissociation . Fragment ions were detected at low resolution in the linear ion trap. Dynamic exclusion was enabled .The lyophilized samples were reconstituted in water/acetonitrile/formic acid  to a final concentration of ~ 0.5 μg/μl. In total, 2 μl each was injected for five technical replicates for LC-MS/MS analyses , frozen, and stored at −80°C. The 15N-labeled aSyn was purified following the regular protocol described above for the unlabeled protein.We designed aSyn gene blocks either WT or containing mutations of interest, which were synthesized by IDT. We used present NdeI and SpeI restriction sites to subclone these genes into ampicillin-resistant pTXB1 plasmid  by ligation. 1H-15N HSQC spectra were recorded at 15°C on an Agilent DD2 spectrometer at 800 MHz using a cold probe. The samples contained 30 μM 15N-labeled monomeric aSyn (WT or E13K mutant) in either 20 mM Hepes (pH 7.4), 100 mM NaCl, 10% D2O or 20 mM sodium phosphate (pH 6.0), 10% D2O. Total acquisition time was 2 hours for each sample. NMR data were processed with NMRPipe (et al. (The


Therefore, the ultra-trace electrochemical determination of Hg2+ in the environment is of critical importance. In this work, a new electrochemical Hg2+ sensing platform based on porous activated carbon (BC/Cu2O) modified with cuprous oxide was developed using a simple impregnation pyrolysis method. Differential pulse anodic stripping voltammetry (DPASV) was used to investigate the sensing capability of the BC/Cu2O electrode towards Hg2+. Due to the excellent conductivity and large specific surface area of BC, and the excellent catalytic activity of Cu2O nanoparticles, the prepared BC/Cu2O electrode exhibited excellent electrochemical activity. The high sensitivity of the proposed system resulted in a low detection limit of 0.3 ng·L−1 and a wide linear response in the ranges from 1.0 ng·L−1 to 1.0 mg·L−1. In addition, this sensor was found to have good accuracy, acceptable precision, and reproducibility. All of these results show that the BC/Cu2O composite is a promising material for Hg2+ electrochemical detection.Mercuric ion (Hg Since Hg2+ has a high affinity for thiol groups to form Hg-S chemical bonds, it can cause severe damage to the brain, kidneys, the central nervous system, the immune system, and the endocrine system 3−/4− solution with 0.1 M KCl at scan rates ranging from 10 to 80 mV·s−1 are shown in p is the peak current value. A represents the effective area of the working electrode. C represents the [Fe(CN)6]3−/4− concentration. v represents the scan rate. D (7.6 × 10−6 cm2·s−1) and n (n = 1) represent the diffusion coefficient and the number of electrons involved in the redox reaction, respectively. Using Equation (1), it was found that the active surface area of the BC/Cu2O/GCE was 0.0803 cm2, which is higher than that of GCE (0.0707 cm2), indicating the superior electrocatalytic activity of BC/Cu2O composite in the redox reaction of Hg2+.The CV plots for BC/Cu2O/GCE were measured using EIS studies. 2O/GCE. As can be seen, there are clear semicircle curves in the EIS spectra of bare GCE, BC/GCE, and BC/Cu2O/GCE. The inset in 2O increased the electron-transfer resistance compared with BC/GCE, which is due to the low electrical conductivity of Cu2O.The inherent interfacial properties of bare GCE, BC/GCE, and BC/Cu−1 Hg2+ on bare GCE, BC/GCE, and BC/Cu2O/GCE was investigated using DPASV in 0.1 M of ABS solution. As shown in 2+ at 0.2 V. Compared with bare GCE, the DPASV signal of BC/GCE slightly increased, which may be attributed to outstanding electrical conductivity and large specific surface area of BC. Furthermore, the GCE modified with BC/Cu2O showed the highest peak current. The unique sensing performance mainly originates from the fact that Cu2O possesses a synergistic adsorption effect with BC to enhance the response current of Hg2+.The electrochemical behavior of 100 μg·L2+, the optimization of pH, deposition potential, deposition time, and material volume is very important.To obtain the best response for the detection of Hg−1 of ABS buffer  on the 100 μg·L2+ signals was investigated in the potential range from −0.6 V to −1.0 V  = 0.0179 + 0.05 C (μg·L−1) (R2 = 0.9975), and the LOD is calculated as 0.3 ng·L−1 (S/N = 3). From 2O composite. Therefore, the proposed BC/Cu2O composite is expected to be a promising candidate for the detection of Hg2+.Under the optimal experimental conditions, the electrocatalytic activity of the BC/Cu2O/GCE were evaluated using the DPASV technique. The repeatability of BC/Cu2O/GCE was examined by detecting 100 μg·L−1 Hg2+ for 15 tests continuously , indicating that the BC/Cu2O/GCE sensor has good anti-interference ability for the determination of Hg2+.To assess the selective sensing ability of BC/Cu2O/GCE was evaluated for the determination of Hg2+ in a lettuce sample. Lettuce was purchased from a local vegetable market, which was chopped and then homogenized in a juicer for 5 min. Afterward, the acquired lettuce juice was further centrifuged and diluted with 0.1 M ABS (pH = 5.0). Under optimal experimental conditions, Hg2+ standard solutions were spiked into the lettuce sample and detected using DPASV. As shown in 2O/GCE can be used as an accurate and efficient sensing platform for Hg2+ detection in real samples.The practicality of BC/Cu4, 98%), ethanol , copper nitrate (Cu(NO3)2·5H2O, ≥99%), and mercury nitrate (Hg(NO3)2, ≥99%) were purchased from Sigma-Aldrich. An acetate buffer solution  was prepared by mixing 0.1 M of sodium acetate and acetic acid. All chemicals are of analytical grade and used without further purification. All the reagent solutions and buffer solutions were prepared using ultra-pure water .Potassium hydroxide , zirconium tetrachloride  spectrophotometer procured from Bruker, Hamburg, Germany. The sample was mixed with KBr and crushed into pellets before the test. The scan range was fixed at 400–4000 cm−1. All electrochemical experiments were carried out on a CHI 660E electrochemical workstation , including electrochemical impedance spectroscopy (EIS) and differential pulse stripping voltammetry (DPASV), with a three-electrode system. A saturated calomel electrode (SCE) and a platinum wire electrode were used as reference electrodes and counter electrodes, respectively. Bare or material-decorated glass carbon electrode  served as the working electrode. X-ray diffraction  measurements of products were characterized with an X-ray diffractometer using Cu Kα radiation (λ = 1.5418 Å) at 40 kV and 40 mA in the 2θ region of 10–50°. The morphology and dimensions of the samples were obtained via scanning electron microscopy . The electronic states of the surfaces of the samples were analyzed using X-ray photoelectron spectroscopy . Thermogravimetric analysis (TGA) was performed using a Q50 Universal V20.13 Build 39 . The sample was tested using a ramp profile (10 °C/min) from 30 °C to 800 °C under N−1, 100 mL). After that, the obtained suspension was sealed in a 150 mL hydrothermal synthesis reactor, followed by a hydrothermal process . After drying at 80 °C overnight, the activated Litsea cubeba was carbonized at 800 °C for 2 h under an Ar atmosphere, with a heating rate of 5 °C·min−1. When cooled down to room temperature, the activated BC was ultrasonically washed with 1 mol·L−1 of HCl and deionized water in sequence until pH = 7. After drying and grinding, BC was obtained.BC derived from Litsea cubeba was produced by slowly pyrolyzing the feedstock under Ar protection. First, the waste Litsea cubeba samples were washed with deionized (DI) water repeatedly to remove physical contaminants on the surface. Subsequently, 2.8 g of Litsea cubeba was dispersed in a KOH solution (1.0 mol·L3)2.5H2O. After being sonicated for 10 min, the mixture was soaked at room temperature for 24 h. Then, the BC rich in Cu2+ ions was dried in a 60 °C drying oven and transferred into a tube furnace. The mixture was heated at 500 °C for 2 h (temperature rise 5 °C/min) in an Ar flow. Finally, the obtained material was washed with 1 mol·L−1 HCl and deionized water until neutral.Briefly, 1 g of BC was immersed in 35 mL of a KOH solution (1 M) containing 1 g of Cu containing Hg2+ with varying concentrations under optimized conditions. DPASV was applied at a deposition potential of −0.8 V for 210 s and was performed in the potential range of 0.10 to 0.26 V under the following conditions: an increment potential of 4 mV, a frequency of 15 Hz, an amplitude of 50 mV, and a pulse width of 50 ms. Prior to the next cycle, BC/Cu2O/GCE was dipped in a fresh supporting electrolyte at a potential of +0.6 V for 180 s in order to remove the residual metal ions on the surface of the working electrode. All measurements were performed at room temperature. The preparation procedure for the BC/Cu2O/GCE composite and the DPASV method for Hg2+ detection is shown in In order to carry out electrochemical sensing of Hg6]3−/4− and 0.1 M of KCl at the formal potential of 0.25 V, with a frequency changing from 0.01 Hz to 100 kHz and a signal amplitude of 5 mV.Electrochemical impedance spectroscopy experiments were performed in a solution including 5 mM of [Fe(CN)2O composites was successfully developed for Hg2+ detection. Benefitting from the synergistic effects of enrichment capacity, strong catalytic activity, high conductivity, and large specific surface area, the BC/Cu2O composite material exhibited excellent electrochemical performance. Using the optimal analytical parameters, the BC/Cu2O/GCE sensor possesses a wider linear range, high reproducibility, and good practical application performance. In addition, the proposed sensor was validated for Hg2+ detection in a real sample with very satisfactory outcomes. Thus, the BC/Cu2O/GCE has great application potential as a sensitive and reliable electrochemical platform for the determination of Hg2+ in food samples.In this work, an electrochemical sensor of BC/Cu


Myeloperoxidase (MPO) is a neutrophil-derived enzyme that has been recently associated with tumour development. However, the mechanisms by which this enzyme exerts its functions remain unclear. In this study, we investigated whether myeloperoxidase can alter the function of A549 human lung cancer cells. We observed that MPO promoted the proliferation of cancer cells and inhibited their apoptosis. Additionally, it increased the phosphorylation of AKT and ERK. MPO was rapidly bound to and internalized by A549 cells, retaining its enzymatic activity. Furthermore, MPO partially translocated into the nucleus and was detected in the chromatin-enriched fraction. Effects of MPO on cancer cell function could be reduced when MPO uptake was blocked with heparin or upon inhibition of the enzymatic activity with the MPO inhibitor 4-aminobenzoic acid hydrazide (4-ABAH). Lastly, we have shown that tumour-bearing mice treated with 4-ABAH had reduced tumour burden when compared to control mice. Our results highlight the role of MPO as a neutrophil-derived enzyme that can alter the function of lung cancer cells. Non-small-cell lung cancer (NSCLC) is a chronic inflammatory disease  that accNeutrophils are important innate immune cells that respond to inflammation  and repr2O2) as a substrate to oxidize halide and pseudohalide ions to produce hypohalous acids, such as hypochlorous acid (HOCl)   and incubated for 15 min in the dark at room temperature. A total of 100 μL of 1× Binding Buffer was added to each sample and analysed within 1 h using a FACS Canto II flow cytometer. The percentages of non-apoptotic cells (AnV−/PI−), early apoptotic cells (AnV+/PI−), late apoptotic cells (AnV+/PI+) and necrotic cells (AnV−/PI+) were recorded.A total of 5 × 106 cells were seeded in a 10 cm dish and incubated for up to 24 h. Cells were then serum starved for 2–4 h or overnight, followed by MPO treatment for the indicated durations. Heparin was incubated for 1 h prior to MPO treatment. Cells were washed with PBS and snap frozen in liquid nitrogen (stored at −80 °C until use) or used fresh for protein lysates.A total of 1–2 × 102 in H2O) containing a 1:100 protease/phosphatase inhibitor cocktail . After centrifugation at 18,620× g for 10 min at 4 °C), the total protein concentration was determined using a Pierce BCA assay . Furthermore, 4× NuPage sample buffer (90 µL 4× LDS sample buffer + 10 µL reducing reagent) was mixed with the samples and boiled for 10 min at 95 °C. Total protein lysates: Cells were scraped off of the plates and lysed using IP buffer , Cell Signaling] in PBS. Cells on ice were homogenized at medium speed for 15 sec, followed by the addition of 25 µL NP-40, and mixed by vortexing. After centrifugation at 2370× g for 5 min at 4 °C, the cytoplasmic fraction in the supernatant was transferred in a separate epi and kept on ice. To generate the nuclear cell fraction, the remaining cell pellet was mixed with 50 µL ice-cold buffer C . The gently dislodged pellet was actively rocked for 30 min in ice and centrifuged at 18,620× g for 10 min at 4 °C. The nuclear fraction in the supernatant was collected in a separate Eppendorf tube. The same volume of both fractions was applied and analysed by Western blot analysis. Nuclear and cytoplasmic fractions: Detached cells were centrifuged at 16,060× g for 5 min. The supernatant with the cytoplasmic fraction was collected in a pre-chilled tube. To dissolve the plasma, mitochondria and ER/golgi membranes, the MEB reagent was incubated with the remaining pellet for 10 min on ice. The nucleus was recovered by centrifugation at 500× g for 10 min and the membrane fraction in the supernatant was collected in a pre-chilled tube. The pellet was further incubated with NEB reagent for 30 min on ice to generate the soluble nuclear extract. After centrifugation at 5000× g for 5 min, the nuclear extract in the supernatant was collected. The remaining pellet was incubated with NEB reagent and micrococcal nuclease  to release chromatin-bound nuclear proteins. After centrifugation at 16,000× g for 5 min, the supernatant containing the chromatin-binding extract was collected in a separate tube. Samples were snap-frozen and stored at −80 °C until use.Subcellular fractions: were generated using the Subcellular Protein Fraction Kit for Cultured Cells  following the instruction manual. In brief, cells were incubated with the CEB reagent for 10 min on ice and centrifuged at 500× Western blot analysis: 20 µg protein from homogenates was electrophoresed for 45 min at 200 V using the NuPage, Bis-Tris gel . To investigate MPO uptake with heparin, equal volumes of homogenates were loaded on the gel. Using the iBlot transfer device, proteins were transferred to a membrane and blocked for 1 h with TBST (1× TBS + 0.1% Tween) containing 5% milk. Membranes were incubated at 4 °C overnight with the primary antibody. The following monoclonal antibodies were used: Anti-MPO , anti-GAPDH , anti-Lamin A/C , anti-tAKT , anti-pAKT (rabbit anti-human phospho-AKT (Ser473), clone: D9E, 1:1000), anti-AKT (rabbit anti-human AKT (pan), clone: 11E7, 1:1000), anti-ERK1/2 , anti-pERK1/2 (phosphor-p44/42 MAPK (Thr202/tyr204); rabbit anti-human pERK1/2, clone: D13.14.4E, 1:1000), anti-Histone3 , anti-HSP90  and anti-EGFR . All antibodies were purchased from Cell Signaling, USA. Afterwards, the membranes were washed for 30 min and incubated for 2 h with the secondary antibody HRP-goat anti-rabbit IgG . Washed membranes were developed using ECL solution .4 or 2 × 105 A549 cells were seeded in 4-well chamber slides or on coverslips in 6-well plates and incubated for up to 24 h at 37 °C. Cells were serum starved for 2–4 h followed by MPO incubation for the indicated timepoints. To investigate MPO uptake by A549 cells, the cells were incubated with heparin (150 µg/mL) for 1 h before adding MPO (10 µg/mL) for 2 h. Cells treated with ddH2O or DMSO (0.2%) were used as the controls. Cells were rinsed with PBS (3 × 2 min) before fixing and permeabilizing with ice-cold methanol for 20 min at −20 °C. Cells were washed with PBS (3 × 2 min on shaker) and incubated with blocking solution (5% goat serum + 5% BSA in PBS with 0.1% TritonX) for 60 min at room temperature (on shaker). The primary antibody  was prepared in an antibody diluent (1:10 blocking solution in PBS-TritonX) and incubated overnight at 4 °C. After washing the cells with PBS (3 × 2 min on shaker), they were incubated with the secondary antibody  for 1 h at room temperature in the dark. After washing the cells with PBS (5 × 2 min on shaker), they were mounted using VECTASHIELD with DAPI. The cells were visualized using an Olympus iX70 fluorescence microscope and the Olympus cellSens Dimension 2.3 imaging software with Z-stacking to allow imaging of different cell layers. A total of 1 × 105 A549 cells were seeded in a 6-well plate and incubated for up to 24 h. After serum starvation for 4 h, the cells were treated with indicated concentrations of MPO for 2 h. Otherwise, cells were treated with heparin (150 µg/mL) for 1 h prior to MPO treatment. Cells were detached using accutase and collected in FACS tubes for intracellular staining with a transcription factor buffer set . To exclude dead cells, FVD eFluor™ 780  was incubated for 30 min at 4 °C. After 2× washing with SB, cells were fixed and permeabilised with 1X TF Fix/Perm Buffer solution for 40 min at 4 °C in the dark. Cells were washed twice with 1X Perm/Wash followed by incubation with 1.5 µL humanTruStain FcX blocking solution  in 1× Perm/Wash and incubated at 4 °C for 10 min. Subsequently, cells were stained with anti-MPO FITC antibody  for 20 min at 4 °C in the dark. After 2× washing with 1× Perm/wash and SB, the cells were analysed with a FACS Canto II flow cytometer. A total of 3.5 × 102O2 production. A549 cells (1.5 × 104) were seeded into a black/clear-bottom 96-well plate and incubated for up to 24 h, followed by 4 h serum starvation. Cells were then incubated with 100 µL of the Fluorescent Peroxide Sensor (FITC) for 15 min and washed with PBS. To stain the nuclei, 1 drop of NucBlue™ Live ReadyProbes™ Reagent  was added to cells in PBS and they were visualized using an Olympus iX70 fluorescence microscope and the Olympus cellSens Dimension 2.3 imaging software. The Intracellular Hydrogen Peroxidase assay kit  was used for imaging of H5 cells were seeded directly or on coverslips in a 6-well plate and incubated for up to 24 h. Afterwards, cells were serum starved for 2 h followed by MPO (5 µg/mL) treatment for 2 h or 3 h for microscopy. For the inhibitor experiments, the cells were first treated with 10 µM 4-ABAH for 30 min before adding MPO (5 µg/mL) for 2 h. Cells treated with ddH2O or DMSO (0.1%) served as the vehicle control. Cells were washed with PBS and incubated with 10 µM R19-S  in serum-free media for 1 h. For signal quantification, cells were washed 2× with PBS, scraped off of the plate and transferred into a 96-well plate, and fluorescence was measured using a CLARIOstar microplate reader . For flow cytometry analysis, cells were collected in FACS tubes, fixed using IC fixation buffer (Thermo Fisher) for 10 min at 4 °C, and the fluorescent signal was quantified using a FACS Canto II flow cytometer (PE channel). For imaging, cells on coverslips were washed 2× with PBS, and fixated and permeabilized with 3.7% paraformaldehyde (PFA) for 10 min at room temperature, followed by 6 min incubation with ice-cold methanol at −20 °C. Cells were rinsed with PBS and incubated in PBS up until the next day. Cells were mounted using VECTASHIELD with DAPI and visualized using an Olympus iX70 fluorescence microscope and the Olympus cellSens Dimension 2.3 imaging software with Z-stacking. A total of 2–2.5 × 105 KP cells suspended in 450 µL PBS (Gibco) were subcutaneously (s.c.) injected into the flank of 6–14-week-old MPO B6 mice. The injections were performed with mice under inhaled isoflurane anaesthesia. A total of 40 mg/kg 4-ABAH was injected intraperitoneally (i.p.) every day until the tumours were harvested. The control group (vehicle) was injected with 200 µL PBS containing 2% DMSO. Tumour growth was monitored and measured (length and width) every day using a digital calliper during the duration of the experiment. After 15 days, tumours were harvested, weighed, and measured with a digital calliper ex vivo. Tumour volume was calculated using the formula: v = length × width × height × π/6 [All animal experiments were performed at the animal facilities of the Medical University of Graz. The experimental protocols were approved by the Austrian Federal Ministry of Science and Research (GZ 2022-0.748.851). Wild-type C57BL/6J (B6) mice were purchased from Charles River, Germany. A total of 5 × 10ht × π/6 .t-tests with Welch´s correction; otherwise, a paired t-test was performed. For comparing three or more groups, one-way ANOVA was used with the Dunnett´s or Tukey´s post hoc test for correction of multiple comparisons. Results are presented as the mean ± SD where the p-value < 0.05 was considered as statistically significant and is shown as 1, 2, 3 or 4 asterisks when lower than 0.05, 0.01, 0.001 or 0.0001. Statistical analyses were performed using GraphPad Prism 6.1 . Significant differences between the two experimental groups were assessed using unpaired Student’s −/−) have up to a 40% reduction in tumour size compared to MPO wild-type mice (C57BL/6 MPO+/+) [Previous studies in our laboratory have shown that MPO-deficient mice  . To furt MPO+/+) . With thA role for MPO in the apoptosis of endothelial and epithelial cells has been reported . For insOur data suggest that MPO can alter the proliferation and apoptosis of A549 cells to benefit their survival. We therefore investigated the effect of MPO in the activation of intracellular signalling pathways that control cell growth, including the AKT and ERK pathways ,34. A549It has been reported that MPO binds to and is internalised into epithelial cells . We therTo further investigate the localisation of MPO after cellular uptake, immunofluorescence staining was performed. Besides the already reported cytoplasmic uptake, a minor MPO signal was also detected in the nuclear region after 2 and 24 h of MPO exposure, while untreated control cells showed no MPO signal A. To valThe mechanism by which MPO binds and internalises into cells is still unclear. It has been reported that highly cationic MPO binds to the negatively charged parts of glycosaminoglycans (GAGs) on the surface of cells and internalises into cells ,24. A stTo determine if MPO can alter the function of A549 cells without binding or entering the cells, we exposed cells to heparin prior to MPO treatment, with effects on apoptosis determined after 15 h of MPO treatment. Blocking MPO uptake with heparin abolished the protective role of MPO in apoptosis. We found no difference in the percentage of early apoptotic cells in untreated control vs. heparin + MPO-treated cells D. These 2O2 in an oxidative reaction with halides to produce reactive oxygen species (ROS), such as HOCl [2O2 in A549 cells. We found that H2O2 is produced by A549 cells under basal conditions, although the signal was not found in every cell.  with an epithelial origin to examine the role of MPO in cancer development. We found that in vitro MPO-treated A549 cells had increased proliferation and reduced apoptosis when compared to untreated control cells. Then, we investigated the activation of AKT and ERK signalling pathways as the major regulators of cell proliferation and survival. Indeed, after 15 min of MPO exposure, we observed an increased activation of these signalling pathways. These data could explain the mechanism by which MPO alters the function of A549 cells. It has been shown that MPO can interact with CD11b/CD18 (Mac-1 integrin) on the surface of neutrophils to suppress apoptosis by increasing the phosphorylation of AKT and ERK . The impWe found that MPO rapidly binds to the surface of lung cancer cells and that the amount of bound enzyme depends on the concentration/availability in the media. The intracellular uptake was observed in the majority (>60%) of cells. Interestingly, we found that MPO partially translocated into the nuclei of A549 cells. It was previously reported that MPO can be found in the nuclei of neutrophils during NETosis . MPO binds to the cell surface via glycosaminoglycans (GAGs), dependent upon its charge ,40. Ther2O2, needed for MPO activity. Using the R19S sensor, which has a high specificity for HOCl [We further investigated whether MPO can retain its enzymatic activity after intracellular uptake. First, we observed that A549 cells produce Hfor HOCl , we obsefor HOCl  and assefor HOCl . FurtherIn conclusion, our results demonstrate that MPO can directly alter cancer cell function in vitro, in a mechanism involving both binding to the cell surface and enzymatic activity. Blocking MPO from binding to cells or treatment with an MPO inhibitor reduced MPO effects on apoptosis and proliferation of lung cancer cells. Inhibiting enzymatic activity in a lung tumour engraftment model resulted in reduced tumour growth when compared to mice with active MPO. However, further studies on the mechanism of action are needed. Our data strongly suggest that MPO is an enzyme that promotes cancer cell growth and could be used as a target for lung cancer treatment.


Post-translational modifications of histones are important regulators of the DNA damage response (DDR). By using affinity purification mass spectrometry (AP-MS) we discovered that genetic suppressor element 1 (GSE1) forms a complex with the HDAC1/CoREST deacetylase/demethylase co-repressor complex. In-depth phosphorylome analysis revealed that loss of GSE1 results in impaired DDR, ATR signalling and γH2AX formation upon DNA damage induction. Altered profiles of ATR target serine-glutamine motifs (SQ) on DDR-related hallmark proteins point to a defect in DNA damage sensing. In addition, GSE1 knock-out cells show hampered DNA damage-induced phosphorylation on SQ motifs of regulators of histone post-translational modifications, suggesting altered histone modification. While loss of GSE1 does not affect the histone deacetylation activity of CoREST, GSE1 appears to be essential for binding of the deubiquitinase USP22 to CoREST and for the deubiquitination of H2B K120 in response to DNA damage. The combination of deacetylase, demethylase, and deubiquitinase activity makes the USP22-GSE1-CoREST subcomplex a multi-enzymatic eraser that seems to play an important role during DDR. Since GSE1 has been previously associated with cancer progression and survival our findings are potentially of high medical relevance.  DNA damage represents a challenge to the genomic integrity on a daily basis. To address this constant threat, the eukaryotic cell developed a system integrating multiple pathways that act synergistically to promote cell survival in response to DNA damage. Once DNA damage is sensed, a cell cycle arrest accompanied by DNA repair signalling is initiated. Impaired repair mechanisms have a severe impact on fitness of the organism, leading to the onset of diseases such as cancer (reviewed in ).Post-translational modifications (PTMs) of histones such as phosphorylation, methylation, monoubiquitination and acetylation were shown to be important regulators of the DNA damage response (DDR) (reviewed in ). For inThere is increasing evidence that all these PTMs must be orchestrated sequentially and many of them are interdependent for proper signal propagation . MoreoveIn this study, we propose a potential mechanism for regulating the DDR by identifying a multi-enzymatic eraser sub-complex of HDAC1/CoREST (co-repressor of Repressor Element 1 Silencing Transcription Factor). This sub-complex combines the activities of the deacetylase HDAC1 and the demethylase activity LSD1 with the deubiquitinase activity of USP22. Using affinity-purification mass spectrometry (AP-MS) we identified GSE1 (genetic suppressor element 1) as a prominent interaction partner of HDAC1, and more specifically, as an interactor of the CoREST complex. While loss of GSE1 did not affect the deacetylase activity of the CoREST complex, it resulted in reduced γH2AX formation and an impaired signalling response upon DNA damage. Further, by in-depth quantitative mass spectrometry, we determine common and individual effects caused by loss of GSE1 or HDAC1 activity. We show that GSE1 acts at an initial step of the DDR. Ultimately, we demonstrate that GSE1 is required for the recruitment of the deubiquitinase USP22 to the CoREST complex, potentially serving as a scaffold, and that loss of GSE1 inhibits deubiquitination of histone H2B at lysine 120, which was previously shown to facilitate γH2AX formation ,35.Through biochemical and phenotypic characterisation of GSE1, we unravel its role in response to DNA damage as a part of multi-enzymatic eraser complex together with HDAC1/2, USP22 and LSD1, which maintains proper chromatin environment and downstream DNA repair signalling. Hence, we emphasise the importance of understanding the cooperation of not only writers but also eraser enzymes within the kinetic window of DNA damage and the chromatin challenges that have to be resolved during DNA repair.2. Treatments of the cells were performed with mitomycin C, etoposide, hydroxyurea or trichostatin A that were purchased from Sigma Aldrich. Mitomycin C (10107409001) was used at final concentration of 500 nM, etoposide (E1383-25MG) at 0.1 μM, hydroxyurea (400046-5GM) at 200 μM and trichostatin A at 0.165 μM if not stated otherwise. The cells were treated either for 6 or 24 h with the chemicals in full culturing media. To harvest, cells were washed once with PBS on ice, scraped and collected by centrifugation . HAP1 cells expressing FLAG-tagged wildtype HDAC1 (HDAC1 WT) or HDAC2 (HDAC2 WT) or catalytically inactive HDAC1  or HDAC2  instead of the respective endogenous enzymes have been previously described , 293T HEK and MEF in Dulbecco's modified Eagle's medium . The culture media was supplemented with 10% foetal bovine serum  and 1% penicillin/streptomycin . The cells were kept in an incubator at 37°C and 5% COescribed .6 HAP1 cells. Human Gse1 plasmid was designed and purchased from VectorBuilder (pRP-hGse1).For transfection of HAP1 cells polyethyleneimine PEI/NaCl solution  and DNA/NaCl solution  were mixed, incubated at room temperature for 30 min, and added dropwise onto a plate of 0.5 × 10Gse1 in HAP1 cells with a V5-tag, Precise Integration into target chromosome (Crispr PITCh) was used as described before (https://eu.idtdna.com/site/order/designtool/index/CRISPR_SEQUENCE) of the IDTDNA company. The RNA duplex complex was assembled by adding 320 pmol guide RNA and 320 pmol tracrRNA (IDTDNA) in IDTE buffer and annealed in the following reaction: 94°C for 5 min, twelve times 93°C for 20 s with -1°C/cycle, 80°C for 4 min, three times 79°C for 20 s with -1°C/cycle, 75°C for 4 min, three times 74°C for 20 s with -1°C/cycle and finally at 70°C for 4 min. Next, the RNP complex was prepared using RNA duplex with 5 μg Cas9 protein in 2 μl cleavage buffer  and incubated for 10 min at room temperature. After incubation, 2 × 106 of HAP1 cells were electroporated using The Neon Transfection System™ (MPK5000) set to 3 pulses, 10 ms width and 1575 V. The obtained single clones were cultured and screened for V5-tag integration by PCR .Guide RNA sequence: GCT ACC TGA AGG GAT ATC CC. Homology arm sequence: 5′-CTG GAC CAC TTA CGA AAG TGC CTT GCC TTG CCT GCA ATG CAC TGG CCT AGG GGC TAC CTG AAG GGA TAT CCC AGG GGT AAG CCT ATC CCT AAC CCT CTC CTC GGT CTC GAT TCT ACG TGA CGG TTT CCC TTG CAC TAG GCC GAA CCT ATA GTA TAG AAA TAT TAT CTA TTT TAT TAC CTT GAA TAT TTA ATA TTT-3′.The GSE1 KO (HZGHC007762c010) and 53BP1 KO (HZGHC000413c010) HAP1 cell lines were obtained from Horizon Genomics (now Horizon Discovery). To tag endogenous human d before . The seqTM 3000  to produce lentiviral particles containing pBOB-EF1-FastFUCCI-PURO according to the manufacturer's protocol. 48 h post transfection, media containing virus was collected, filtered, and added onto HAP1 cells. After 48 h, the positive cells were selected by addition of 1 μg/ml puromycin for two days. FUCCI positive cells were cultured and washed once with phosphate-buffered saline  and fixed with 4% methanol-free formaldehyde  for 20 min at room temperature. Afterwards, the samples were washed twice with PBS and resuspended in the final volume of 200 μl PBS and taken for measurement. The samples were measured with Cytoflex Flow Cytometer (Beckman Coulter Life Sciences) and analysed with CytExpert software (Beckman Coulter Life Sciences) or FLowJo (TreeStar Inc.).To monitor changes in cell cycle, fluorescent ubiquitination cell cycle indicator system (FUCCI) was used as described before . The 293To generate whole-cell lysates, cells were washed in 1 ml PBS on ice and centrifuged at 300 × g/5 min. The pellets were lysed in Hunt buffer  supplemented with Complete Protease inhibitor cocktail (Roche 11697498001), Complete Phosphatase inhibitor cocktail , 10 mM sodium fluoride, 10 mM β-glycerophosphate, 0.1 mM sodium molybdate and 0.1 mM PMSF. The samples were frozen and thawed three times in liquid nitrogen and centrifuged . The supernatants were transferred, and protein lysates concentration measured by Bradford assay. Detection of proteins was performed by SDS-PAGE and western blots using nitrocellulose membranes and chemiluminescence. The blots were developed using ECL Western Blot detection reagents  and visualised via chemiluminescence using Fusion FX7-Spectra machine.Antibodies: GSE1 (KIAA0182) , HDAC1 SAT 208 , HDAC2 clone 3F3 , CoREST , β-actin , cleaved caspase-3 , γH2AX , vinculin , LSD1 , USP22 .3H]-acetate-labelled chicken reticulocyte core histones (1.5–2 mg/ml) were added. Following 3 h incubation on thermomixer , 35 μl of Histone stop solution  and 800 μl of ethyl acetate were added and vortexed for 15 s. The samples were centrifuged  and 600 μl of organic upper phase was mixed with 3 ml of Scintillation Solution . Released acetate was measured in counts per minute (CPM) with a Liquid Scintillation Analyzer (Packard).The deacetylase assay was conducted on beads carrying precipitated proteins in 20 μl volume as previously described ,38. 4 μl2, 50 mM NaHSO4, 1% Triton X-100, adjusted to pH 6.5) supplemented with Complete Protease inhibitor cocktail (Roche 11697498001), Complete Phosphatase inhibitor cocktail , 10 mM sodium fluoride, 10 mM β-glycerophosphate, 0.1 mM sodium molybdate and 0.1 mM PMSF and centrifuged . The nuclear pellet was washed three times with 1 ml of histone lysis buffer, once with histone wash buffer  and centrifuged . After each washing step the cells were briefly vortexed. Next, the pellets were resuspended in 100 μl H2O and concentrated H2SO4 was added to final concentration of 0.4 N. After 1 hr incubation on ice, the cells were centrifuged . The supernatants were transferred into a new tube with 10 times volume of acetone, vortexed for 1 min and precipitated o/n at −20°C. The precipitates were collected by centrifugation , air-dried and resuspended in H2O. 2 μg of histones were used for SDS-PAGE and western blot.To isolate histones, all the buffers were pre-cooled and kept on ice throughout the whole procedure. Cells were washed once with PBS on ice and centrifuged (300 × g/5 min). The pellets were lysed in histone lysis buffer  for 1 min and then fixed with 4% methanol-free formaldehyde  for 20 min at room temperature. The samples were washed two times with PBS and permeabilized with 0.5% Triton-X in PBS for 10 min at room temperature. After three times washing with PBS, the samples were blocked with 5% bovine serum in PBST for 1 h at room temperature. After blocking, the primary antibody was added in blocking solution and incubated overnight at 4°C. The samples were washed three times with 3% bovine serum in PBS for 5 min and stained with secondary antibody in blocking solution for 1 h at room temperature in the dark. Next, samples were washed three times with 3% bovine serum in PBS for 5 min, incubated with DAPI in PBS for 10 min at room temperature and washed again two times with PBS. Afterwards, the slides were mounted using DAKO mounting medium and dried. Antibodies: DAPI , γH2AX , H3Ser10ph , Alexa Fluor 546 , Alexa Fluor 488 .et al.  was measured. For quantification of G2 and M phase positive cells stained with H3Ser10ph antibody. As described in De La Barre et al.  the cellWhole-cell lysates were prepared as described above . 40 μl Protein A or Protein G Dynabeads  per sample were washed three times with Hunt buffer, resuspended in Hunt buffer containing 10% bovine serum albumin and incubated for 1 h rotating at 4°C for blocking. At the same time, the whole-cell extract (0.5–1 mg protein) was incubated with an antibody mix suitable for immunopurification of the protein of interest for 1 h rotating at 4°C. After blocking, the beads were added to the cell lysate-antibody mix and incubated overnight at 4°C rotating. Next day, the beads were washed twice with TBS for 5 min and once for 15 min at 4°C. Subsequently, the purified proteins were eluted from the beads in 1× SDS loading dye by heat-incubation at 95°C for 5 min. The eluate was used for SDS-PAGE and western blotting.Antibodies: 4 μg COREST , 4 μg USP22 , 2 μg GSE1 , 30 μl of monoclonal HDAC1 (clone 10E2) and HDAC2 (clone 3F3) generated in collaboration with Egon Ogris .Whole cell extracts were prepared as follows: cell pellets were resuspended in Hunt buffer , sonicated twice for 3 s , and centrifuged at 21 000 × g for 15 min at 4°C. Protein concentration of supernatants was determined via Bradford. 1 mg of protein was used for each sample.80 μl of V5-beads  per sample were used for isolation of Gse1-V5, 120 μl of crosslinked M2-FLAG beads  were used for purification of HDAC1-FLAG. Briefly, for antibody crosslinking the bead slurry was equilibrated with sodium borate buffer (200 mM sodium borate pH 9.0), resuspended in 20 mM dimethyl pimelimidate in sodium borate buffer, incubated rotating for 1 h at room temperature, washed and incubated with 200 mM ethanolamine pH 8.0 for 2 h, washed with 100 mM glycine and subsequently stored in 1× TBS. Beads were washed three times with Hunt buffer, resuspended in whole cell extracts and incubated overnight rotating at 4°C. Next day, the beads were washed twice with TBS for 5 min and once for 15 min rotating at 4°C. Beads were transferred to a 0.2 ml PCR tube, re-buffered to and washed once with ammonium bicarbonate (ABC)  buffer, pelleted by gentle centrifugation, and resuspended in 50 μl 2 M urea , 50 mM ABC.Predigestion of bound proteins was performed by addition of 200 ng/μl of a 1:1 LysC  trypsin  mix, followed by an incubation at RT for 90 min in the dark. The eluate containing predigested proteins and peptides was separated from the beads, and beads were subsequently rinsed with 50 μl ABC. The eluates were united.Predigested proteins and peptides were reduced with 10 mM dithiothreitol (DTT)  for 30 min at RT, and subsequently carbamidomethylated using 20 mM iodoacetamide (IAA)  followed by incubation for 30 min at RT in the dark. The alkylation reaction was quenched by adding 5 mM DTT and incubation for 10 min. Samples were digested overnight at 37°C using 150 ng/μl of the 1:1 LysC and trypsin mix, acidified using 10% trifluoroacetic acid (TFA)  to a final concentration of 0.5% and desalted on C18  StageTips as described in .m/z, automatic gain control (AGC) target = 3e6, and maximum injection time (IT) = 60 ms. MS2 analysis was performed with the following parameters: resolution 30 000, AGC target = 1e5, maximum IT = 250 ms, isolation window 1.6 m/z. The mass spectrometer was configured to pick the eight most intense precursor ions for data-dependent MS2 scans, applying HCD for fragmentation with a normalised collision energy (NCE) of 28. Dynamic exclusion time was set to 30 s.MS measurements were performed on a Q Exactive HF Orbitrap  mass spectrometer (Thermo Fisher), operated in data-dependent acquisition (DDA) mode with dynamic exclusion enabled. For peptide separation on the HPLC, the concentration of organic solvent  was increased from 1.6% to 32% in 0.1% formic acid at a flow rate of 230 nl/min, using a 2 h gradient time. Peptides were ionised with a spray voltage of 1,9 kV. MS1 survey scans were recorded with the following parameters: resolution 120 000, scan range 375–1500 MS raw files were searched using MaxQuant (version 2.1.4.0) against the Homo sapiens (human) (HAP1 samples) or the Mus musculus (mouse) (MEF samples) Uniprot database (release 2022.02) and a database of common laboratory contaminants. Enzyme specificity was set to ‘Trypsin/P’. Carbamidomethylation of cysteine was searched as a fixed modification. Protein N-term acetylation and Oxidation (M) were set as variable modifications. The data were filtered at 1% FDR (PSM and protein), reverse hits were removed. MaxLFQ and match-between-runs were activated. MaxQuant output was further processed in R using Cassiopeia (DOI: 10.5281/zenodo.5758975). After removing contaminants and proteins with less than two razor & unique peptides, missing LFQ protein intensities were imputed if at least two valid values were present in one experimental group. Imputation was performed in log2 space by drawing random values from a normal distribution . LIMMA was used to determine differentially enriched proteins at 1% FDR (Benjamini–Hochberg).Inspection of the HDAC1-FLAG IP data obtained from HAP1 cell extracts Figure  revealedThree biological replicates of each experimental condition were prepared. To induce DDR cells were treated with 0.1 μM etoposide  for 6 h. Cells were washed twice in 1× PBS, supplemented with 10 mM sodium butyrate  and 3 mM nicotinamide (NAM) , and pelleted prior to lysis.7 cells per replicate were resuspended in lysis buffer , 50 mM Tris/HCl pH8.0, 150 mM NaCl, 1 mM PMSF, 5 mM sodium butyrate , 10 ng/ml NAM , 10 ng/ml trichostatin A (TSA) , 1× cOmplete protease inhibitor cocktail (+EDTA) , 250 U/replicate benzonase ), lysed using a Bioruptor sonication device (Diagenode) , centrifuged at 15 000 × g for 10 min at 4°C and precipitated with 4× volumes of cold (−20°C) 100% acetone . Proteins were washed with cold (-20°C) 80% acetone, air dried for 5 min, dissolved in 8 M urea, 50 mM ABC , reduced with 10 mM DTT  for 45 min at RT, and subsequently carbamidomethylated with 20 mM IAA  for 30 min at RT in the dark. The alkylation reaction was quenched by adding 5 mM DTT for 10 min. The urea concentration was reduced to 4 M using 50 mM ABC. Samples were pre-digested for 90 min at 37°C with Lys-C  at an enzyme-to-substrate ratio of 1:100 and subsequently digested overnight at 37°C with sequencing grade trypsin  at an enzyme-to-substrate ratio of 1:50. Digests were stopped by acidification with TFA   and desalted on a 50 mg tC18 Sep-Pak cartridge . Peptide concentrations were determined and adjusted according to UV chromatogram peaks obtained with an UltiMate 3000 Dual LC nano-HPLC System , equipped with a monolithic C18 column (Phenomenex). Desalted samples were dried in a SpeedVac concentrator (Eppendorf) and subsequently lyophilized overnight.MS sample preparation was performed as previously described . Briefly2O, pH 7–8, dilute 1:10) on an UltiMate 3000 Dual LC nano-HPLC System  equipped with a XBridge Peptide BEH C18  column (Waters) (flow rate of 1.0 ml/min). Fractions were collected and subsequently pooled in a non-contiguous manner into 8 pools, dried for 30 min using a SpeedVac concentrator (Eppendorf) and subsequently lyophilized overnight.500 μg of trypsin-digested and desalted peptides were used for each sample for isobaric labeling. Lyophilized peptides were dissolved in 20 μl 100 mM triethylammonium bicarbonate (TEAB) . 60 μg of peptides in TEAB from each sample and replicate were additionally mixed  as reference sample (REF). 500 μg of each TMT 10plex reagent  were dissolved in 30 μl of acetonitrile (ACN) VWR, 83639.320) and added to the peptide/TEAB mixes. The REF sample was mixed with 1.5 mg (90 μl) of TMT 10plex reagent 131N and subsequently split in 3 aliquots. Labels used: 126C: wildtype (wt) replicate (rep) 1; 127C: wt rep 2; 128C: wt rep 3; 129C: HDAC1 CI rep 1; 130C: HDAC1 CI rep 2; 127N: HDAC1 CI rep 3; 128N: GSE1-KO rep 1; 129N: GSE1-KO rep 2; 130N: GSE1-KO rep 3; 131N: REF. Untreated and etoposide treated samples were labelled using 2 separate TMT 10plex reagent kits which were subsequently processed individually. Samples were labelled for 60 min at RT. 0.1 μl of each sample were pooled, mixed with 10 μl 0.1% TFA and analysed by MS to check labelling efficiency. Reactions were quenched using hydroxylamine  , pooled, desalted using Sep-Pak tC-18 (200 mg) cartridges , dried in a SpeedVac vacuum centrifuge (Eppendorf), and subsequently lyophilized overnight. Neutral pH fractionation was performed as previously described 39.320 an, using aAcetylated peptides were enriched, washed, eluted and desalted as described in , using t2 resin = 1:6) of TiO2  was washed twice with 50% methanol  and twice with glycolic acid solution , 70% ACN , 3% TFA ). Lyophilized peptides were dissolved in glycolic acid solution, mixed with the TiO2 resin, and incubated rotating at RT for 30 min. Samples were loaded onto Mobicol columns  and shortly centrifuged in a table centrifuge to remove unphosphorylated peptides. Phosphorylated peptides will bind to the TiO2 resin. The resin was washed twice with glycolic acid solution, twice with 200 μl 70%, ACN 3%, TFA and twice with 1% ACN, 0,1% TFA. Phosphorylated peptides were eluted twice using 150 μl 300 mM ammonium hydroxide , eluates were united and immediately acidified with conc. TFA to a pH of 2,5. Samples were desalted using a standard C18  StageTip protocol  target 4e5, max injection time of 50 ms, FAIMS CV −40 V, −55 V and −70 V, dynamic exclusion 45 s), and MS2 scans . RTS was enabled with full trypsin specificity, max. 2 missed cleavages, max. search time of 40 ms, max. 2 variable modifications, max. 10 peptides per protein, and considering the following modifications: Carbamidomethyl on cysteine and TMT-6-plex on Lys and peptide N-terminus as static, oxidation on Met and deamidation on Asn and Gln as variable modifications. The scoring thresholds were applied  and the Homo sapiens Uniprot database (release 2020.01) was used for the search. Quantitative SPS-MS3 scans were performed in the Orbitrap with the following settings: HCD normalised collision energy 50%, resolution 50 000; AGC target 1.5e5; isolation window 1.2 m/z, MS2 isolation window 3 m/z, 10 notches, max injection time of 150 ms. Total cycle time for each CV was set to 1.2 s.DDA-SPS-MS3 method with online real-time database search (RTS)  was performed on an UltiMate 3000 Dual LC nano-HPLC System , containing both a trapping column for peptide concentration  and an analytical column , coupled to a Orbitrap Eclipse mass spectrometer (Thermo Fisher) via a FAIMS Pro Duo ion source (Thermo Fisher). The instruments were operated in data-dependent acquisition (DDA) mode with dynamic exclusion enabled. For peptide separation on the HPLC, the concentration of organic solvent  was increased from 1.6% to 32% in 0.1% formic acid at a flow rate of 230 nl/min, using a 2 h gradient time for proteome analysis. Peptides were ionised with a spray voltage of 2.4 kV. The instrument method included Orbitrap MS1 scans  target = Custom, and maximum injection time (IT) mode = Auto, FAIMS voltages on, FAIMS CV −40 V, −55 V and −70 V, dynamic exclusion was set to 45 s. MS2 analysis was performed with the following parameters: resolution 45 000, normalised AGC Target 200%, AGC target = Custom, maximum IT mode = Custom, isolation window 0.7 m/z, and HCD normalised collision energy of 34%.DDA-MS2 method  was performed on an Orbitrap Exploris 480 mass spectrometer (Thermo Fisher) and similar HPLC and nanospray settings as described above, with the exception of a FAIMS Pro ion source and that 3 h gradient time was applied. MS1 survey scans were recorded with the following parameters: resolution 60 000, scan range 375–1500 MS raw files were split according to CVs  using FreeStyle 1.7 software (Thermo Scientific). All raw MS data were commonly analysed using MaxQuant  softwareData were analysed in R (4.1.0) using custom scripts . The analysis procedure covered: correction for isotopic impurities of labels, within-plex median normalisation, between-plex IRS-normalisation  and statPerseus  Python k-means algorithm was used. TMT reporter ion intensities were log2 transformed and z-scored. Mapping of human to mouse proteins was done using the ortholog database from OrthoInspector 3.0 (database version 2019)  , proteinon 2019) . For baret al. , thereby providing a comprehensive analysis of the interactions of the enzyme  C-terminally FLAG-tagged HDAC1 from the nearly haploid human cell line HAP1 ,51 and ae Figure .The presence of well-known subunits of co-repressor complexes such as NuRD, Sin3-related, MiDAC or CoREST  in the Het al. .To examine cross-species conservation in the composition of HDAC1 complexes we additionally analysed HDAC1-FLAG purifications from mouse embryonic fibroblasts (MEFs) Figure . Again, 2] = 12.37, adjusted P-value < 0.001, Figure BRemarkably, a previously reported interaction between HDAC1 and Genetic suppressor element 1 (GSE1) ,54–57, t3-acetylated histones as substrate . In addition, we demonstrate that GSE1-HDAC1/HDAC2 complexes are catalytically active, and their deacetylase potential is dependent on HDAC1 and HDAC2 activity. Moreover, GSE1 ablation has no impact on deacetylase activity of HDAC1 and HDAC2.et al. was first to identify GSE1 as a potential mediator of cellular sensitivity to the topoisomerase II inhibitor etoposide  and p53 binding protein 1 (53BP1) foci, which are indicative for single and double strand breaks respectively. Indirect immunofluorescence microscopy revealed a moderate increase in the number of 53BP1 foci in untreated conditions in GSE1 KO cells in comparison to wildtype, which is even more prominent in response to DNA damage treatment . NotablyIncreased sensitization of cancer cells with decreased homologous recombination repair capacity was observed upon HDAC inhibition in combination with antitumor drugs ,62–64. SDDR is vastly orchestrated by a sequential cascade of multiple signalling events that are tightly regulated. In addition, a functional role between phosphorylation and acetylation crosstalk was established in cardiomyocytes where inhibition of HDACs can lead to increased phosphorylation of proteins such as p38, ERK1, and AKT (reviewed in ).P-value of ≤ 0.05 were considered as significantly regulated. However, a higher level of variance was observed in untreated acetylome samples. We therefore used more stringent filtering for the acetylome dataset .To further investigate how HDAC1 and GSE1 can regulate DDR and whether their mode of action might be overlapping, we next performed a proteomic analysis using quantitative mass spectrometry based on the recently developed TMTpro 16-plex isobaric labelling technology . We deteFirst, we investigated etoposide-induced acetylome changes in HAP1 WT, GSE1 KO and HDAC1 CI cells. We detected a rather small population of increased lysine acetylation of sites in HAP1 WT upon etoposide treatment, which is in accordance with the previous studies, reporting only minor changes in acetylation levels upon 24 h of etoposide treatment  or in reP-value ≤ 0.01), suggesting that the mutants have to a certain extent individual effects on the acetylome  in wildtype cells but remain unaffected in both mutants  in response to DNA damage  up- and 1859 (7.6%) down-regulated phosphorylation sites in HAP1 WT cells  and to a lesser extent also in the HDAC1 CI mutant  Figure . NotablyPadj-value of ≤0.05, in wildtype cells). Analysis of the networks revealed proteins mostly associated with DDR and DNA repair, but also with spliceosome, ubiquitination machinery, and histone modifications  levels are affected in GSE1 KO cells, we followed this modification in a time-resolved manner using western blotting in combination with an anti-H2Bub antibody. In wildtype cells we observed a prominent and continuous decline in H2Bub levels from the 6 h to the 24 h time point of etoposide treatment. Interestingly, in GSE1 KO cells H2Bub levels remained constant throughout the same timeframe Figure , demonstInterestingly, GSE1 seems to exclusively bind to the CoREST complex as subunits of other known corepressor complexes such as SIN3, NURD or MiDAC were not enriched by the GSE1 pull down . In addiWe did, however, detect traces of UCH proteinases USP22 and USP15 in the affinity purification samples of GSE1-V5, but not in the background controls  complexes ,54–57. Het al., who demonstrated the heterogeneous composition and functional diversity of cellular CoREST complexes , providing an estimate of protein abundance, as well as data provided at ProteomicsDB , ImmProtomplexes . Consistntiation .What is the biological function of the GSE1-USP22-CoREST complex? Here we show that loss of GSE1 results in hampered γH2AX formation, affects the abundance of HDR as well as NHJE-related proteins , shows altered impaired phosphorylation of key sites in DNA damage signalling and an overall compromised DDR. These findings point to a connection between GSE1-USP22-CoREST and the regulation of DNA damage initiation. One of key requirements in this process seems to be the GSE1-dependent binding of USP22 to CoREST. We hypothesise that during DNA damage USP22 is essential for removal of H2B K120ub, which seems to be a prerequisite signal for γH2AX formation Figure . It was It also should be mentioned, however, that other studies reported an increase of H2B monoubiquitination levels in response to DNA damage . We coulOur identification of the GSE1-HDAC1-USP22 multi-eraser complex and its function in DNA damage might be of relevance for cancer research. Various studies recently connected GSE1 function to tumour growth in distinct types of cancer and highlighted its potential oncogenic and tumour suppressive functions . OverexpIn our study, we show that GSE1 is essential to integrate different enzymatic activities. It is, however, still an open question how these activities are regulated in response to DNA damage. We can rule out changes in deacetylase activity of HDAC1. We do not know how and whether the activity of USP22 is regulated, or whether the complex becomes localised to specific sites of damaged chromatin. However, we also dismiss DNA damage induced changes in complex composition. Future work will certainly focus on these aspects. The fact that inhibitors of separate enzyme activities combined in the GSE1-HDAC1-USP22 complex are currently applied and tested in the clinics for cancer patients is potentially putting our findings into the context of clinical relevance ,108.gkad911_Supplemental_FilesClick here for additional data file.