Leishmania parasites responsible for visceral leishmaniasis. The compounds were screened for in vitro activity against L. infantum and L. donovani and for cytotoxicity in two monocytic cell lines (J774A.1 and THP-1) using the methyl thiazol tetrazolium assay. Reactive oxygen species formation, apoptosis, and mitochondrial impairment were measured by flow cytometry. The compounds exhibited fair to moderate anti-proliferative activity against promastigotes of the 2 Leishmania species, with IC50 values ranging from 13.0 ± 1.7 µM to 793.0 ± 37.2 µM. Tetraoxanes LC132 and LC138 demonstrated good leishmanicidal activity on L. infantum amastigotes (IC50 13.2 ± 5.2 and 23.9 ± 2.7 µM) with low cytotoxicity in mammalian cells (SIs 22.1 and 118.6), indicating selectivity towards the parasite. Furthermore, LC138 was able to induce late apoptosis and dose-dependent oxidative stress without affecting mithocondria. Compounds LC132 and LC138 can be further explored as potential antileishmanial chemotypes.Leishmaniasis remains one of the ten Neglected Tropical Diseases with significant morbidity and mortality in humans. Current treatment of visceral leishmaniasis is difficult due to a lack of effective, non-toxic, and non-extensive medications. This study aimed to evaluate the selectivity of 12 synthetic endoperoxides  and uncover their biochemical effects on  Leishmania that create impacting illness burden in 98 countries. The WHO estimates 12 million human infections worldwide, with over 1 million new cases annually, leading to different forms of the disease, namely cutaneous (CL), mucocutaneous (MCL), or visceral leishmaniasis (VL). The latter occurs in up to 90,000 cases and is fatal if untreated, with an estimated 65,000 deaths per year  released during parasite haemoglobin digestion, leading to cleavage of the endoperoxide ring and consequent formation of reactive species that react with relevant parasite targets, e.g., proteins or lipids, through alkylation or peroxidation, promoting parasite death , the highest among the compounds synthesized , being a common apoptosis-necrosis process that occurs in an attempt to rescue the life of ATP-deprived cells from the mitochondria  was added to a final concentration of 5 µg/mL in 1× phosphate-buffered saline , with an additional 4 h incubation at 36 °C, as previously described [50 values were calculated fitting the data as a non-linear regression with variable slope using a dose-response inhibitory model, in the GraphPad Prism V 8.0 program. Each compound was tested in three independent assays.Cell lines J774A.1 and THP-1 were used to estimate CCompounds  were addescribed . The for50, promastigotes of L. infantum and L. donovani were plated in 96-well flat bottom culture plates at a concentration of 5 × 106 parasites/mL and incubated with six serial concentrations of each compound  and anti-Leishmania spp. activity (IC50) (SI = CC50/IC50).SI was determined for each compound, expressed by the ratio between cytotoxicity in cell lines (CC5 cells/mL in supplemented DMEM medium. 200 μL/well of the cell suspension was placed in 16 well Lab Tek culture glass slides (Nunc), and incubated at 37 °C ± 1 °C, 5% CO2, for 3 h, to enable the cells to adhere to the slides. After a washing step with DMEM to discard non-attached cells, the cells were infected with 2.5 × 106 cells/mL promastigotes in a 10:1 parasite-host cell proportion [v/v). Slides were incubated for 48 h at the same temperature. After a washing step with 1× PBS (Sigma), slides were fixed and stained, and the infection index was calculated according to [50 of each compound, which was calculated by fitting the data as a non-linear regression with variable slope using a dose-response inhibitory model, in the GraphPad Prism V 8.0 program. SI was determined as described above.The intracellular assay using the macrophage J774A.1 cell line was performed with the compounds that presented best SI on the promastigote’s susceptibility assay. J774A.1 cells were adjusted to 2.5 × 10oportion  for 24 hrding to  by countL. infantum strain (MHOM/PT/1988/IMT151) was chosen for the cytometry analysis and promastigotes at a concentration of 1 × 107 parasites/mL were tested against the three compounds evaluated in the intracellular amastigotes assay . Flow cytometry analysis were performed in a CytoFlex flow cytometer (Beckman Coulter), at an excitation wavelength of 488 nm (blue laser). Data acquisition were collected and registered using channels FL1  with a BP filter (525 ± 40 nm) for Fluorescein and FL2 (IP) with a BP filter (570 ± 20 nm) for propidium iodide (PI) and 10,000 total events per sample were analysed using CytExpert v2.0.0.153 software. Samples were analysed in triplicates and two to three independent experiments were performed. Values are expressed as a percentage of cells for a given marker, relatively to the number of cells analysed ± standard error of mean (SEM).The L. infantum promastigotes were incubated with IC50 and 2 × IC50 of the tested compounds and amphotericin B, for 24 h and 48 h, at 24 °C ± 1 °C. H2O2 (2 mM) was used as positive control and nontreated parasites as negative control. In addition, amphotericin B was used as control drug. After a washing step with cold 1 × PBS , cells were resuspended in 1 × Binding Buffer. 100 μL of cell suspension was incubated with 5 μL of Annexin V and 5 μL of PI for 15 min at RT in the dark. After adding 400 μL of 1 × Binding Buffer, samples were analysed in Cytoflex flow cytometer. Annexin V-FITC and PI were measured in FL1 and FL2 channels, respectively. The following controls were used to set up compensation and quadrants: unstained cells, cells stained with FITC Annexin V (no PI), cells stained with PI (no FITC Annexin V).To study the apoptosis inducing capacity of the most selective compounds, PS translocation from the inner to the outer leaflet of the plasma membrane was accessed, as it is a characteristic event of early apoptosis. To distinguish apoptotic cell death from necrotic cell death, cells were counterstained with PI, a non-permeable stain with affinity for nucleic acids, as it selectively enters necrotic cells. Parasites were stained with Annexin V and PI according to the manufacturer’s instructions . Briefly, Parasites populations were characterized as follows: (1) viable: Annexin V-/Pi-; (2) early stages of apoptosis: Annexin V+/Pi−; (3) late stages of apoptosis: Annexin V+/Pi+ and (4) other type of cell death: Annexin V-/Pi+.L. infantum promastigotes were incubated with IC50 and 2 × IC50 of the tested compounds and amphotericin B for 24 h, at 24 °C ± 1 °C. Nontreated parasites were included as negative control and, as positive control, promastigotes were incubated with a mitochondrial membrane potential disrupter, carbonyl cyanide 3-chlorophenylhydrazone . Treated and nontreated parasites were incubated with JC-1  cationic dye, according to the manufacturer’s instructions. This dye exhibits potential-dependent accumulation in mitochondria, indicated by a fluorescence emission shift from red (~590 nm), JC-1 aggregates to green (~529 nm) JC-1 monomers indicating membrane depolarisation. Briefly, parasites were incubated with JC-1 in PBS  at 24 °C ± 1 °C, for 30 min. Then, parasites were washed once with warm PBS, centrifuged  and resuspended in 500 μL PBS. Cells were analysed by flow cytometry. Cells were gated to exclude debris and standard compensation was done with CCCP-treated parasites.Mitochondrial damage was evaluated through the analysis of the mitochondrial membrane potential with MitoProbe™ JC-1 Assay Kit for Flow Cytometry . Treated 2DCFDA  was used. Briefly, promastigotes were incubated with IC50 and 2 × IC50 of the tested compounds and amphotericin B for 5 h and 24 h, at 24 °C. Nontreated control and a positive control (H2O2 1 mM) were included. Following incubation step, parasites were washed in pre-warmed PBS, centriguged  and resuspended in PBS with CM-H2DCFDA in a final concentration of 5 μM for 30 min, at 24 °C, and then washed again with PBS. Parasites were resuspended in normal medium (M199 medium) and fluorescence was measured by flow cytometry (blue laser), and the signals registered in the FL1/FITC channel.For the detection of reactive oxygen species (ROS), a cell permeable probe CM-HLeishmania nuclear DNA, promastigotes (1 × 107 parasites/mL) were incubated for 24 h with IC50 concentrations of LC132, LC137 and LC138. 2 mM H2O2 was used as DNA damage inductor and non-treated cells were used as control. After centrifugation  DNA was extracted using a commercial extraction kit , according to the manufacturer’s instructions. Extracted DNA was observed in a 2% (w/v) agarose gel at 100 V for 90 min.To detect whether the compounds induced fragmentation on 50 and CC50 were expressed as the mean ± standard diviation (SD) of three independent experiments. Cell population retrieved from cytometry analysis were expressed as % ± SEM GraphPad Prism 8  was used to compare differences among groups in flow cytometry data using two-way ANOVA with Dunnet’s multiple comparisons test using different significance levels .Results of ICLeishmania sp. upon treatment should also be evaluated, in order to have more insigths on compounds targets and mode of action.Our findings showed that the selected endoperoxides LC132 and LC138 induce some biochemical and morphological alterations, causing oxidative stress. More biochemical studies are needed to better elucidate the mode of action and the cell death pathway displayed by these compounds. Ultrastrutural alterations of Leishmania species- L. infantum and L. donovani- but LC138 showed to be more selective in L. infantum when cytotocicity was considered. Upon optimization and pharmacomodulation of these compounds, efficacy evaluation in a VL in vivo model with toxicity evaluation and host immune system involvement, we expect that these class of endoperoxides may furnish promising therapeutic leads against Old-World Leishmania parasites worthy of pharmacological consideration.LC132 presented better anti-leishmanial intracelular activity on both Old-World


E. coli fermentation followed by cell disruption and filtration, and a single purification step via cation exchange chromatography. The process was developed at bench scale and then pilot scale, where the PCV2d subunit protein yield was approximately 0.93 g/L fermentation volume in a short production time. Moreover, we have successfully implemented this production process at two different sites, in Southeast Asia and Europe. This demonstrates transferability and the high potential for successful industrial production.Porcine circovirus type 2d (PCV2d) is becoming the predominant PCV genotype and considerably affects the global pig industry. Nevertheless, currently, no commercial PCV2d vaccine is available. Preventing and controlling the disease caused by PCV2d is therefore based on other genotype-based vaccines. However, their production platforms are laborious, limited in expression level, and relatively expensive for veterinary applications. To address these challenges, we have developed a simple and cost-efficient platform for a novel PCV2d vaccine production process, using fed-batch  E. coli production can be performed to high cell density cultivation, and the process can be readily conducted on a large scale with a short production process time. Moreover, current processes render yields as high as g/L scales of product [Porcine circovirus type 2 (PCV2) is one of the most pervasive viral pathogens repeatedly affecting the swine meat production industry worldwide, as infection causes loss in pig production, mostly due to growth retardation, reduction in average daily weight gain, and reproductive issues. Importantly, this leads to immune suppression, increasing the risk of other pathogenic infections. PCV2 is also associated with the occurrence of a postweaning multisystemic wasting syndrome (PMWS), porcine dermatitis and nephropathy syndrome (PDNS), and porcine respiratory disease complex (PRDC) ,2. Over ®) , inactiv PCV G2) , and bac G2) [®) ,15. Thes G2) [®) ,17,18. T product ,20,21.Recently, a novel PCV2d vaccine and a laboratory-scale production process were developed in order to produce small quantities of test material for animal experiments and vaccine characterization . HoweverE. coli W3110 containing pET23/ptac Δ2–40 PCV2d-His6 were received from the Robinson group at the University of Kent (UK) [TM, Sparks, NV, USA) containing 5 g/L glycerol and 100 µg/mL ampicillin at 30 °C and 200 rpm in a shaking incubator . Cells were mixed with 40% w/v glycerol in a 1:1 ratio and stored at −80 °C for use in this whole study.Cell stocks of ent (UK) . ResearcThe preculture was started with 1% inoculation of RCB to 50 mL TB in a shake flask and incubated at 30 °C and 200 rpm overnight. For cell expansion to a pilot-scale fermenter, the first preculture was inoculated into a second preculture with 10% cell density in 200 mL SM6Gc media supplemented with antibiotics and cultured at 30 °C and 200 rpm until OD600 reached 5–7.4)2SO4, 4.4 g/L NaH2PO4·H2O, 4.03 g/L KCl, 1.04 g/L MgSO4·7H2O, 4.55 g/L citric acid monohydrate, 0.25 g/L CaCl2·2H2O, and 95 g/L glycerol. The pH was adjusted to 7.0 using 25% NH4OH. The trace elements were prepared at 100× stock solutions containing 113.48 g/L citric acid monohydrate, 5.22 g/L CaCl2·2H2O, 2.06 g/L ZnSO4·7H2O, 2.028 g/L MnSO4·H2O, 0.81 g/L CuSO4·5H2O, 0.42 g/L CoSO4·7H2O, 10.06 g/L FeCl3·6H2O, 0.03 g/L H3BO3, and 0.02 g/L Na2MoO4·2H2O. The trace element solutions were filtered through a 0.22 µm sterile syringe filter and stored in dark bottles at 4 °C. Filters and chemicals were all purchased from Merck KGaA unless otherwise stated.SM6Gc comprises SM6 and its trace elements, which were prepared separately. SM6 media consisted of 5.2 g/L (NH3PO4 or 25% NH4OH. The pH dead band was set to 0.1. Dissolved oxygen (DO) concentration (%DO) was calibrated at 0 by unplugging the probe’s cable. Calibration of 100% DO was achieved by keeping the mixing speed at 1050 rpm and the airflow rate at 60 Nl/h. The %DO setpoint was 30 and was controlled by a cascade of agitation speeds (450–1050 rpm), airflow, and O2 flow. A total gas flow of 1 vvm (volume of gas sparged per fermenter working volume per minute) was used. The starting temperature was set at 30 °C and was reduced to 25 °C once the cells reached an OD600 of 60–70. The temperature was controlled through perfused stainless-steel baffles connected to the heating circuit with a circulation pump, electrical heater, and cooling water valve. In addition, 5% Antifoam 204  was used to prevent overflow of the media during cultivation through control of the level probe and peristaltic pump.A 3.7 L Ralf Advanced  stirred-tank glass fermenter, equipped with a 2-flat-blade Rushton agitator, a 6-blade agitator for radial mixing, 4 stainless steel baffles, and a ring sparger, was used in this work. A quantity of 1.5 L of SM6 media was transferred to the fermenter followed by autoclaving at 121 °C for 20 min . An amount of 15 mL 100× trace elements solution was filter-sterilized via a 0.22 µm syringe filter into the vessel before inoculation. The pH set point of 7.0 was controlled by a cascade using internal peristaltic pumps for the addition of 10% H4·7H2O when OD600 reached 38–42, 1.164 g/L NaH2PO4·H2O at OD600 of 54–58 and 1.63 g/L NaH2PO4·H2O when starting the fed-batch process at OD600 around 66–70. The feeding media, 800 g/L glycerol, was fed constantly at 0.15 mL/min (per 1.5 L working volume) using the internal pump. When OD600 exceeded 75, cells were induced with 0.039 g/L Isopropyl β-d-1-thiogalactopyranoside (IPTG). Cells were grown until an OD600 of 200 was reached, then the temperature was lowered to 15 °C prior to proceeding to cell harvest by centrifugation at 7500× g for 30 min . Wet cell paste was collected and stored at −20 °C for downstream processing. Fermentation samples were collected throughout the cultivation for OD600 measurements. A 1 mL cell suspension was taken, before and after induction, and spun down, and the pellet was kept for SDS-PAGE and Western blot analysis.Fermentation was performed with a starting OD600 of around 0.8. A series of supplements were added through a 0.22 µm sterile syringe filter as follows: 1.972 g/L MgSO® D-DCU, Sartorius Stedim Biotech GmbH, Guxhagen, Germany) with 4 stainless-steel baffles and 3 six-bladed Rushton turbines. A quantity of 20 L of SM6 media was prepared and transferred to the fermenter through an external peristaltic pump. Then, automatic sterilization in place (SIP) was conducted using a recipe pre-programmed in BioPAT® MFCS|win. After cooling down, 200 mL 100X trace element solution was added to the sterile media using a 0.2 µm sterile disc filter . Fed-batch processes at pilot scale were performed in a 42 L stainless-steel stirred-tank bioreactor . Fermentation was terminated after cells were grown for more than 48 h and OD600 was over 200. Wet cell paste was collected after centrifugation and stored for future use. Samples were collected and analyzed, as described in the sample preparation and analytical methods section.E. coli W3110 producing PCV2d capsid protein was resuspended in resuspension buffer  at various NaCl concentrations depending on which CIEX equilibration buffer was used. A disperser was used for mixing at 3000 rpm for 15 min . The resuspended cell sample was lysed by cell disruption using a high-pressure homogenizer  at 700, 1400, and 2100 bars for 10 passages until low viscosity of lysed cells was observed. Cell debris was separated by centrifugation at 20,000x g for 30 min at 10 °C. The supernatant was collected and filtered through a depth filter  and a 0.45 µm PES membrane disc filter . A turbidity meter  was used to measure sample clarification before applying the sample onto the chromatographic column.Next, 15% of wet cell paste of Cation exchange chromatography was conducted in a one-step purification . A small-scale experiment used 2× HiScreen SP Sepharose Fast Flow columns (Cytiva) connected in series. The linear velocity was kept at 300 cm/h. The binding conditions were optimized based on our preliminary studies  using 50A HiScale 16/40 column  was packed with SP Sepharose Fast Flow resin  to 20 cm bed height, requiring 40 mL of resin. Column efficiency was tested according to Cytiva’s protocol. The flow rate was kept constant at a linear velocity of 300 cm/h for the entire run. The column was equilibrated with 50 mM Tris-HCl, 300 mM NaCl, pH 7.0 in 3 column volumes (CV). The clarified supernatant from the product recovery step was applied onto the column for 5 CV using the sample pump. Unbound proteins and impurities were first washed with 4 CV of equilibration buffer and then with a step gradient at 40% of elution buffer  for another 4 CV. PCV2d was eluted at 100% of elution buffer for 4 CV. Flow through, washing steps, and elution were all collected in 40-, 40-, and 10-mL fractions, respectively. The purified fractions were collected for SDS-PAGE and Bradford analysis.Cell pellets from fermentation runs were resuspended in 50 mM Tris-HCl, 2.5 mM EDTA, pH 7.0 with the same volume that was taken out after centrifugation. Resuspended cells were lysed by sonication using a microtip (6.4 mm) with 70% amplitude at 5 s On-time and 10 s Off-time for a total of 1 min On-time . The lysed samples were centrifuged at 14,000 rpm for 10 min. The supernatant and inclusion bodies (IB) were separately collected and mixed with 4× Laemmli sample buffer with reducing agent 2-mercaptoethanol and boiled for 10 min. For SDS-PAGE and Western blot, 12% SDS-PAGE gels were used. After transfer onto a polyvinylidene difluoride membrane, the membrane was blocked overnight with 3% blotting-grade blocker in Tris-buffered saline with 1% Tween-20 (TBST) at 4 °C. After washing with TBST, the membrane was incubated for 1 h with anti-porcine circovirus antibody at 1:2,000,000, obtained from the Robinson lab at University of Kent, UK. The secondary antibody, Goat Anti-rabbit IgG H&L , was used at 1:10,000 dilution. The membrane was imaged on ChemiDoc™  with Western ECL Substrate. The Precision Plus protein dual color standard was used as a molecular weight marker. Protein concentrations were determined using Bradford assay. All reagents were sourced from Bio-Rad unless otherwise stated.The Global Challenges Research Fund (GCRF) consortium has recently published a relatively inexpensive PCV2d vaccine candidate and a PCV2-PCV3 chimera vaccine candidate shown to effectively induce PCV2d- and PCV2-PCV3-neutralizing antibodies in immunized animals. In brief, their work showed the production of antisera to PCV2d and the neutralization effect of antisera with regard to PCV2d infection in cell-based assays. The virus-neutralizing (VN) titers were 28 ± 7 and 19 ± 2 for Rabbit 1 and Rabbit 2, respectively. The VN titers for pre-immunization sera were insignificant. This convinced them that the construct shows high potential for being further developed as a subunit vaccine against PCV2d, and hence that proceeding to large-scale process development should be carried-out .Building on this work, the PCV2d subunit vaccine manufacturing process was further developed and verified at a pilot scale. Here, we present proof that the PCV2d vaccine candidate can be easily manufactured at a commercial scale. The process flow diagram is summarized in At first, fermentation of PCV2d was attempted at a working volume of 1.5 L. Seed trains were prepared in 50 mL TB reaching an overnight OD600 of 10–12 and transferred into each fermenter where cells were cultured in chemically defined media (SM6Gc). Defined media provides benefits over other media, such as reducing batch-to-batch variability observed in complex media, increased process control for simple downstream processing, and lower vaccine production costs ,24.−1. At the end of the fermentation, cell densities reached up to OD600 of 230, with the lowest cell density observed at OD600 of 170  to the fed-batch stage for induction of the l growth ,26. PCV2−1. Cells were grown to an OD600 of 220 corresponding to 6.59 kg wet cell paste with a total cultivation weight of 27.99 kg. This resulted in the calculation of 235 g wet cell paste per kg culture weight. Samples from various time points after IPTG induction were analyzed for PCV2d expression on SDS- PAGE and Western blot, as illustrated in To this end, and using the established bench scale parameters, the PCV2d vaccine was produced in large scale, demonstrating production capability in a 42 L stainless-steel fermenter with a 20 L initial working volume. The growth rate in large-scale fermentation was shown to be comparable to the 1.5 L fermentation runs shown in −1.To illustrate the process transferability and reproducibility of this simple upstream process, large-scale production was carried out in parallel at the consortium partner site in the UK, as shown in Next, PCV2d was purified from 60 g cell mass derived from large-scale fermentations. The cells were resuspended in 400 mL Tris-EDTA buffer and disrupted by homogenization in continuous mode. The homogenized pressures were varied between 700 and 2100 bar, and 700 bar was observed to be the optimal operating condition, as seen from the measurement of turbidity after centrifugation as well as the band intensity of protein release on the SDS-PAGE. The number of passages was set to 10, as the lowest viscosity of the mixture was observed with this number of passages. After centrifugation was carried out for liquid–solid separation, the supernatant fraction was then collected. A series of depth filtration and membrane filtration steps were then performed through 0.5–15 µm retention rating filters and 0.45 µm membrane filters. The turbidity of the filtered sample was measured at 215 NTU. Although turbidity seemed to be rather high, no untypical increase in column pressure was observed during sample application on the ion exchange column.The PCV2d protein has an isoelectric point (PI) of 9.4. Thus, a cation exchange column with bind/elute mode at pH 7.0 was selected for downstream purification. SP Sepharose Fast Flow has been widely used in preparative protein separations and is well known for high speed and low cost. Our preliminary studies see  with linHere, 200 mL of clarified sample was loaded with a flow rate of 10 mL/min. The chromatogram shown in We have presented a biomanufacturing process that is simple, transferable, and scalable to a large-scale protein subunit vaccine production for PCV2d. This process is based on a proof of concept of a novel PCV2d vaccine candidate that successfully induces PCV2d-neutralizing antibodies in immunized animals. All steps were carefully developed and evaluated to avoid excessive investment costs in low- and middle-income regions or high processing costs while maintaining high quantities for veterinary applications. We are optimistic that this developed process can benefit vaccine manufacturers, particularly in the region where the PCV2d vaccines are inaccessible. Additionally, we are convinced by our comparative studies between production sites that the presented PCV2d vaccine platform is highly likely to produce effective vaccines with high yields. However, other characterization parameters for veterinary vaccine products  may be required to meet the regulatory aspects. Further investigation in a pig’s field trial to assess the protection of our vaccine against a virulent PCV2d challenge may need to be considered.


Using the OHD4–12, we show that the CXCL4–CXCL12 chemokine heterodimer inhibits the CXCL12-driven migration of triple-negative MDA-MB-231 breast cancer cells. We also show that the CXCL4–CXCL12 chemokine heterodimer binds and activates the CXCR4 receptor.Chemokines form a family of proteins with critical roles in many biological processes in health and disease conditions, including cardiovascular, autoimmune diseases, infections, and cancer. Many chemokines engage in heterophilic interactions to form heterodimers, leading to synergistic activity enhancement or reduction dependent on the nature of heterodimer-forming chemokines. In mixtures, different chemokine species with diverse activities coexist in dynamic equilibrium, leading to the observation of their combined response in biological assays. To overcome this problem, we produced a non-dissociating CXCL4–CXCL12 chemokine heterodimer OHD Chemokine signaling is mediated by intermolecular interactions with G protein-coupled receptors (GPCRs), cell surface glycosaminoglycans (GAGs), and through chemokine homooligomerization4. Additionally, different chemokines can interact with each other to form heterodimers. These heterophilic interactions have been directly detected by several methods, including co-immunoprecipitation and ligand blot12, surface plasmon resonance11, mass spectrometry13, and NMR (Nuclear Magnetic Resonance) spectroscopy17. A pairwise bidirectional immunoblot chemokine screening shows that heterophilic interactions are abundant in the chemokine family identifying approximately 200 distinct interactions12.Chemokine signaling is essential in normal physiologic and pathological conditions24. Treatment with chemokine mixtures demonstrated either a synergistically enhanced or a reduced activity dependent on the microenvironment and the nature of heterodimer-forming chemokines tested25. For example, the heterodimerization of platelet-derived CXCL4 with CXCL8 chemokine inhibits the activation and proliferation of endothelial cells, and the CXCL8-induced migration of cells transfected with the CXCR2 chemokine receptor15, whereas CXCL4-CCL5 chemokine heterodimerization promotes the arrest of CCL5-stimulated monocytes on activated endothelium11. In addition, chemokines CXCL10 and CCL22, co-expressed in the inflamed skin, synergistically enhance the CCR4-mediated chemotaxis of T cells25. Likewise, CCR7 chemokine ligands CCL19 and CCL21 enhance monocytes recruitment by forming heterodimers with CCL7 and CCL222, while heterodimers formed by CXCL9 and CXCL12 chemokines co-expressed in the perivascular area of the tumor enhance CXCR4-mediated recruitment of tumor-infiltrating lymphocytes and malignant B cells6.The role of heterophilic interactions in chemokine signaling remains to be understood. However, it is established that cell responses to chemokine mixtures differ from individual chemokines26. For example, the disruption of the CCL5-CXCL4 heterodimer decreased the CCL5-mediated neutrophil influx, edema formation, and destruction of lung tissue in acute lung injury26 and attenuated monocyte recruitment, thereby reducing atherosclerosis in mice9. However, the use of heterophilic interactions as drug targets requires an understanding of the mechanism of action of chemokine heterodimers, which is currently lacking.The in vivo co-localization, particularly near GAGs, produces favorable conditions for chemokine heterodimerization. In fact, a few studies have already demonstrated the relevance of the heterophilic interactions in in vivo animal models and the possibility of targeting such interactions for therapeutic benefit28). The measured biological outcomes represent the combined response to chemokine species, rendering the direct functional assessment of the heterodimer biological activity challenging. Here, we overcome this limitation by generating a new, non-dissociating heterodimer of CXCL4 (platelet factor 4) and CXCL12  chemokines, OHD4–12 (Obligate HeteroDimer CXCL4–CXCL12). CXCL4 and CXCL12 are both stored and released upon stimulation of platelets29, thus having ample opportunities to interact in vivo and modulate cell responses in chemokine-rich microenvironments. We also provide proof-of-principle evidence that OHD4–12 is a valuable tool for investigating the biological activities and the mechanism of action of the CXCL4–CXCL12 chemokine heterodimer. In particular, we show that the OHD4–12 binds and activates the CXCL12 receptor CXCR4 and inhibits the CXCL12-driven migration of MDA-MB-231 breast cancer cells. The mechanistic insight obtained from using the OHD4–12 may extend to other chemokine heterodimers and inform experiments testing their mode of action.In chemokine mixtures, competing homophilic and heterophilic interactions lead to the equilibrium coexistence of different chemokine species including monomers, dimers, heterodimers, and, in some cases, higher-order oligomers  experiments using antibodies specific to CXCL4 and CXCL12. The CXCL4-S26C or CXCL12-L29C fractions were first immunoprecipitated with magnetic microbeads coated with anti-CXCL4 or anti-CXCL12 antibodies and then detected using anti-CXCL12 antibodies, demonstrating the presence of the OHD4–12 heterodimer  NMR spectrum of the 15N-OHD4–12 displayed well-dispersed cross-peaks, which confirmed the presence of a folded structure . Unlike CXCL4, the CXCL12 chemokine produces the 15N-HSQC NMR spectrum, in which a single resonance represents each amino acid. Some heterodimer cross-peaks originating from the CXCL12 counterpart can be tracked by comparing the 15N-HSQC NMR spectra of the 15N-OHD4–12 (black cross-peaks) and 15N-CXCL12 (red cross-peaks). Some of these peaks are labeled in Fig. 4–12 and CXCL12. Supplementary Fig. 4–12, CXCL12, and CXCL4 with known assignments for CXCL4 and CXCL12 labeled.We then used NMR spectroscopy to assess the folding state of the OHD16. Here, we similarly assessed the effect of the OHD4–12 on the migration of MDA-MB-231 cells. As expected, the wild-type CXCL12 (CXCL12wt) and the mutant CXCL12-L29C, used to produce the disulfide-trapped heterodimer, induced the migration of MDA-MB-231 cells  and dimer (CXCL12D) on cell migration35  molar ratios  or CXCR3b, respectively41. Previously, we determined that MDA-MB-231 breast cancer cells strongly express CXCR4 and CXCR3b receptors16. The expression of the CXCR7 receptor on MDA-MB-231 cells is marginally low43. In our experimental setting, the percent of CXCR7 receptor expression on MDA-MB-231 cells was 1.6 ± 0.1, whereas the CXCR4 and CXCR3b expression was 23.5 ± 2.7 and 38.0 ± 0.2%, respectively  incubated with different amounts of the OHD4–12. The addition of OHD4–12 to MDA-MB-231 cells induced a dose-dependent increase of cytoplasmic Ca2+  does not permit the investigation of cell response to chemokine heterodimers apart from chemokine monomers, homodimers, and, possibly, higher-order oligomers. Therefore, a powerful strategy to address this challenge is to use obligate, non-dissociating chemokine heterodimers in functional studies36. Here, we generated a new, obligate CXCL4–CXCL12 chemokine heterodimer, named OHD4–12. The CXC-type arrangement of chemokine monomers forming the CXCL4–CXCL12 heterodimer16 is particularly suitable for introducing cysteine substitutions at the intermonomer interface for disulfide bond formation. Therefore, we followed the disulfide-trapping strategy31 for forming the OHD4–12.The existence of the equilibrium of interconverting chemokines species in situ , including CXCL12 and CXCL4. The GAG-binding of a few chemokine heterodimers was characterized31 and it was also shown that binding to GAGs stabilized chemokine heterodimers13. Thus, it is plausible that the two heterodimer-forming chemokines dissociate from GAGs as a heterodimer, or remain close to each other to form a heterodimer, readily available for receptor binding. Further interest in CXCL4–CXCL12 heterodimer comes from the critical role of the CXCL12 chemokine and its receptor, CXCR4, in invasion, migration, and proliferation steps of tumor progression in more than 75% of all cancers, including breast, ovarian, lung, colon, prostate, kidney, melanoma, brain, esophageal, pancreatic, and various forms of leukemia51. Indeed, organs with the highest levels of CXCL12 expression  are the most common sites of metastasis for breast cancer cells52. CXCL4 is abundant in platelets of healthy individuals. The levels of CXCL4 and CXCL12 can be further elevated in cancer patients54 leading to micromolar concentrations at the tumor or metastasis site that could favor CXCL4–CXCL12 heterodimerization in vivo. Therefore, we tested the activity of the OHD4–12 on the migration of invasive MDA-MB-231 breast cancer cells. MDA-MB-231 are triple negative breast cancer cells extensively used in the investigation of therapeutic targets as well as mechanistic studies of cancer cell migration57. Although other cells, including different types of cancer cells, may trigger stronger than MDA-MB-231 cell migration responses58, MDA-MB-231 cells uniquely express CXCR4 and CXCR3b but not CXCR7 , inducing conformational changes that lead to signaling61. The formation of the CXC-type CXCL4–CXCL12 heterodimer does not involve the N-terminus, N loop, or the 40 s loop of the chemokine ligand, and they remain available for interaction with the receptor  and the globular core, N loop, and 40 s loop of the chemokine4–12 binds and activates the CXCR4 receptor is in accordance with previous observations that disulfide-trapped CXCL1-CXCL7 heterodimer binds and activates the CXCR2 receptor17, whereas CC-type obligate CCL5-CCL17 heterodimer involves both corresponding receptors CCR4 and CCR512. These observations suggest that chemokine heterodimers may act by involving receptors of both heterodimer-forming chemokines. Our data show that the addition of the specific CXCR4 inhibitor AMD3100 abrogates the OHD4–12-induced Ca2+ release, whereas the addition of the specific CXCR3 inhibitor AMG487 does not  required for the activation of CXCL4-CXCR3 Ca2+ signaling. Indeed, Korniejewska et al.64 used 10 μM CXCL4, and Mueller et al.65 reported that 500 nM CXCL4 was required to induce intracellular calcium flux in activated T cells. The 20-fold concentration discrepancy was proposed to be related to the difference in stimuli used for cell activation64. Further investigation will reveal whether the OHD4–12 can bind and activate the CXCR3 receptor.The observation that the OHDM  and CXCL12D (disulfide-linked) on the migration of colon cancer HCT116 and HT29 and monocytic leukemia THP-1 cells was previously observed35. CXCL12D competitively blocked CXCL12wt-induced cell migration and, in contrast to CXCL12M, was not able to stimulate the migration of these cells at the concentration of 10 nM35. Our data parallel this result as we observe a reduced migration of MDA-MB-231 cells treated with 100 nM CXCL12D as compared to CXCL12wt or CXCL12M and CXCL12-L29C variants. The difference in CXCL12M and CXCL12D activity stems from different binding modes to the CXCR4 receptor62. In the case of CXCL12M, the N-terminal domain of the CXCR4 receptor wraps around the chemokine and forms a beta-sheet with its first beta-strand β1, leading to an active signaling complex that promotes chemotaxis66. The lack of the CXCL12D chemotactic activity is caused by the inaccessibility of the β1 strand of the CXCL12 monomer for the interaction with the receptor because it participates in the inter-monomer contact with the β1 strand of an opposing CXCL12 monomer62.The difference in effects of CXCL12D, we observe no significant effect of the OHD4–12 on the migration of MDA-MB-231 breast cancer cells and the inhibition of the CXCL12wt-induced MDA-MB-231 cell migration by the OHD4–12. The design of the OHD4–12  vector (Novagen) were purchased from Genscript. Unlabeled or uniformly 15N-enriched proteins were expressed in BL21(DE3) pLysS E. coli bacteria (Novagen) grown in LB or M9 media, respectively, at 37 °C with shaking at 250 rpm in the presence of 60 μg/mL kanamycin. M9 medium was supplied with 15N-NH4Cl  (Cambridge Isotope Laboratories) as a sole source of nitrogen. Protein production was induced by the addition of IPTG  to the final concentration of 0.5 mM when the optical density of bacterial culture at 600 nm (OD600) reached 0.6. After the addition of IPTG, bacteria were grown for 4 h at 37 °C and harvested by centrifugation for 30 min at 3000 rpm. Bacterial pellet was resuspended in the lysis buffer (3 ml per gram of pellet), contained 50 mM Tris, 1% Triton, 100 mM PMSF (phenylmethylsulfonyl fluoride), and 0.1% of beta-mercaptoethanol (BME), prepared at pH 8.0. Bacteria were disrupted by sonication  on ice at 40% amplitude for 2 s on/0.5 s off with breaks between cycles to prevent overheating. After sonication, bacteria lysates were centrifuged for 1 h at 20,000 rpm and 4 °C. All proteins expressed as inclusion bodies. Inclusion bodies were homogenized by stirring overnight at 4 °C in the extraction buffer prepared at pH 8.0 (12 ml per gram of pellet with inclusion bodies), containing 50 mM Tris, 8 M Urea, and 0.1% BME. Cell debris were removed by centrifugation at 20,000 rpm and 4 °C for 1 h, and the supernatant containing unfolded soluble proteins was used in subsequent purification steps.Plasmids with DNA encoding CXCL4wt, CXCL12wt, and CXCL4-S26C proteins were eluted using a 0–100% gradient of elution buffer containing 100 mM Tris, 6 M urea, 2 M NaCl, pH 8.0. Following cation exchange chromatography, the refolding of CXCL4wt, CXCL12wt, and CXCL4-S26C was performed by 1:50 drop-wise dilution with constant stirring in 100 mM Tris buffer containing 10 mM cysteine and 1 mM cystine at pH 8.0. CXCL12-L29C was refolded on-column by gradually decreasing the concentration of urea from 8 to 1 M using 50 mM Tris buffer containing 10 mM reduced glutathione and 1 mM oxidized glutathione at pH 7.3. Following overnight incubation, the column was washed with 50 mM Tris buffer, pH 7.3, containing 50 mM NaCl. Folded CXCL12-L29C was eluted from the column with a 0–2 M gradient of NaCl. All refolded proteins were further purified by heparin affinity chromatography using HiPrep Heparin FF 16/10 column (Cytiva). The proteins were eluted in the 50 mM Tris buffer at pH 7.3 using a 0–2 M NaCl gradient. The CXCL4wt and CXCL4-S26C, eluting at high NaCl concentration, were sufficiently pure and were ready for subsequent experiments. The purity of CXCL12wt and CXCL12-L29C, eluting at ~ 0.3 M NaCl, was additionally improved by size-exclusion chromatography using HiPrep 26/60 Sepharcryl S-200 HR column (Cytiva) and 50 mM sodium phosphate buffer containing 150 mM sodium chloride at pH 7.0.The proteins were initially purified by cation exchange chromatography using the ÄKTA pure 25 M FPLC system and HiTrap SP/FF 16/10 column (Cytiva). CXCL44–12, mutants CXCL4-S26C and CXCL12-L29C obtained after the heparin affinity chromatography step, were mixed at 1:1 molar ratio and dialyzed against 50 mM sodium phosphate buffer, containing 150 mM NaCl and 10 μM CuCl2 at pH 7.0 for 18 h at 4 °C. Following the dialysis, the mixture was centrifuged to remove the precipitated proteins and the OHD4–12 was purified using HiTrap Heparin FF 16/10 column (Cytiva). The OHD4–12 eluted at 0.5–0.6 M of NaCl. The purity of OHD4–12 was polished by size-exclusion chromatography using a Superdex 75 Increase 10/300 GL column (Cytiva).To form the disulfide-trapped heterodimer OHDM) and dimer (CXCL12D) were a gift from Protein Foundry, LLC.Obligate CXCL12 monomer . Membranes were blocked in Tris-buffered saline containing 0.1% Tween-20 and 5% non-fat milk at 4 °C and incubated with the primary anti-CXCL4 and anti-CXCL12 antibodies  overnight. The membranes were then incubated with species-specific horseradish-peroxidase (HRP)-conjugated secondary antibodies . Following the incubation with ECL substrate (BioRad), the presence of specific proteins was determined based on chemiluminescence detected using a ChemiDoc Imaging System (BioRad). For co-IP analysis, chemokine mixtures were incubated with magnetic microbeads pre-coated with mouse anti-CXCL4 or anti-CXCL12 antibodies (R&D Systems) at 4 °C for 2 h. After incubation, microbeads were magnetically bound and washed to remove the unbound fractions. After elution, the Western blot analysis of the immune-precipitated samples was performed using goat anti-CXCL12 monoclonal antibodies (R&D Systems).Chemokine mixtures or the OHD2O in the presence of 20 mM of NaCl at pH 5.0. 15N-CXCL12 and 15N-OHD4–12 (both counterparts were 15N-labeled) were prepared at concentrations 129 and 68 μM, respectively, in 90% H2O/10% D2O in the presence of 20 mM of NaCl at pH 6.9. The two-dimensional 15N-HSQC (heteronuclear single quantum coherence) spectrum of CXCL4wt was recorded on the Bruker Avance 950 MHz spectrometer at David H. Murdock Institute . 15N-HSQC spectra of CXCL12wt and OHD4–12 were recorded on the Bruker Avance III 700 MHz NMR spectrometer equipped with a helium-cooled cryoprobe at METRIC, North Carolina State University. All spectra were recorded at 40 °C.15N-CXCL4 was prepared at the concentration of 150 μM in 90% H2O/10% Dl-glutamine, Amphotericin B and Gentamycin (Corning). Following an overnight coating with Collagen type I  at 37 °C with 5% CO2 and > 85% humidity and washes of the unbound Collagen I with sterile PBS, 96-well tissue culture plates (Greiner) were seeded (4 × 105 cells/well) with MDA-MB 231 cell suspension in culture media. Cells were grown to confluence and then incubated overnight with fresh media without FBS (0%). The confluent MDA-MB 231 cell monolayers were then scratched using a sterile pipet tip and the wells washed to remove non-adherent cells. Thereafter, cells were incubated with various chemokines and overlapping microphotographs encompassing the entire area of each scratch/wound were taken at the start of the treatment and following the incubation for 9 h using an IX71 Olympus microscope equipped with a DP70 camera and the associated software (Olympus). Overlapping microphotographs were stitched together, and the area of the wound was determined using ImageJ software (NIH). After normalization to the area measured at time 0, results were expressed as percentage of wound healing. Differences between treatments were tested by ANOVA and individual treatment compared through post-hoc Tukey tests with a priori significance threshold set at p < 0.05.MDA-MB 231 breast cancer cells (ATCC) were cultured in DMEM/F12 media (Corning) supplemented with 10% FBS (Atlanta Biologics), 5 cells in 100 μL per well on 96-well tissue culture plates (Greiner) and grown to confluence. After 48 h, cells were starved in serum-free media for 6 h and then incubated  with the Ca2+ intracellular indicator Fura-2 (2 μM). Cells were washed with phosphate buffered saline (PBS). Following injections of increasing concentrations of chemokines (0–250 nM), variations in cytoplasmic Ca2+ were measured every 1.5 s for up to 60 s by detecting the 510 nm fluorescence emission ratio following excitation at 340 nm and 380 nm, respectively, using the ID5 plate fluorescence reader (Molecular Device). Background fluorescence was measured for 30 s prior to the addition of chemokines. Fluorescent signals were subsequently normalized to the average background reading.MDA-MB 231 breast cancer cells (ATCC) were seeded at 4 × 10Supplementary Figures.


Amyloid β-protein (Aβ) plays an initiating role in Alzheimer’s disease (AD), but only a small number of groups have studied Aβ extracted from human brain. Most prior studies have utilized synthetic Aβ peptides, but the relevance of these test tube experiments to the conditions that prevail in AD is uncertain. Here, we describe three distinct methods for studying Aβ from cortical tissue. Each method allows the analysis of different ranges of species thus enabling the examination of different questions. The first method allows the study of readily diffusible Aβ with a relatively high specific activity. The second enables the analysis of readily solubilized forms of Aβ the majority of which are inactive. The third details the isolation of true Aβ dimers which have disease-related activity. We also describe a bioassay to study the effects of Aβ on the neuritic integrity of iPSC-derived human neurons. The combined use of this bioassay and the described extraction procedures provides a platform to investigate the activity of different forms and mixtures of Aβ species, and offers a tractable system to identify strategies to mitigate Aβ mediated neurotoxicity. Compelling evidences from genetic, neuropathological, biochemical and biomarker studies have shown that the amyloid β-protein (Aβ) plays a central role in Alzheimer’s disease (AD) . Recent Our prior work has shown that the primary structure of Aβ from human brain is highly heterogeneous with a diversity of N- and C-termini . In contHere, we describe three protocols for the preparation of Aβ-containing AD brain extracts and a paradigm to investigate AD relevant bioactivity. The first section of the protocol details two methods to extract aqueous soluble forms of Aβ. Method I involves soaking thin slices of brain in artificial cerebrospinal fluid (aCSF). Method II utilizes a more traditional homogenization procedure. Method III describes a protocol that enables isolation of size exclusion separated pure Aβ dimers and monomers. Finally, we detail the use of human iPSC-derived glutamatergic neurons and live-neuron imaging to measure the neurite disrupting activity of various Aβ-containing extracts.Here we provide details of vendors and catalog numbers for all chemicals and reagents we used for the experiments described. However, common reagents/chemicals from other sources should work equally well.•Milli-Q water.•NaCl .•KCl .•2PO4 .NaH•3 .NaHCO•Leupeptin .•Aprotinin .•Pepstatin A .•EGTA .•Pefabloc .•NaF .•0.5 M EDTA .•Protein A sepharose .•Anti-Aβ polyclonal antibody S97 .•Pre-immune serum .•Trizma-base .•Hydrochloric Acid .•BSA .•Sodium tetraborate .•Dimethyl pimelimidate (DMP) .•Ethanolamine .•Glycine .•3 .NaN•20% Sodium dodecyl sulfate .•β-mercaptoethanol .! CAUTION This reagent is highly caustic. Do not allow its contact with the skin or eyes. Use safety goggles and an 8-inch face shield when handling.•Sucrose .•Congo red .•88% formic acid .! CAUTION This reagent is highly caustic. Do not allow its contact with the skin or eyes. Use safety goggles and an 8-inch face shield when handling.•Ammonium bicarbonate .•Blue dextran .•Gel filtration standard .•mTeSR Matrix .•mTeSR medium-1L kit .•Penicillin/Streptomycin .•Y-27632 .•Accutase .•PBS .•DMSO .•Neurobasal™ Medium .•DMEM/F12 .•GlutaMAX™ Supplement .•Puromycin .•Doxycycline Hyclate .•Dextrose .•N2 supplement B .•MEM non-essential amino acids .•KnockOut™ DMEM .•Knockout Replacement Serum .•Blocker A .•4xMSD Read Buffer P .•Streptavidin SULFO-TAG Labeled .•Tissue chopper .•Bench top centrifuge .•Ultracentrifuge .•Overhead stirrer .•15 mL tissue grinder .•Lyophilizer .•FPLC system .•Light microscope with a polarizing filter (Leica).•Water bath and dry bath.•Lab vacuum system.•IncuCyte live-cell video microscopy system (Essen Bioscience).•Double-Edge Stainless blade .•Scalpel blade .•Superdex 75 10/300 column .•HiTrap desalting column .•Zeba spin 40k MWCO desalting column .•Slide-A-Lyzer G2 dialysis cassettes, 2000 MWCO .•Cover slips .•96-well micro plate .•0.2 μm Syringe filters .•0.2 μm Membrane filters .•1 mL Syringes .•10 mL Syringes .•50 mL falcon tubes .•13.2 and 38.5 mL Ultracentrifuge tubes .•1.5 mL protein low bind tubes .•2 mL protein low bind tubes .•1.5 mL protein low bind tubes .•40 μm Sterile Cell Strainers .•112 μm nylon mesh filter (cat. no. U-CMN-112-A).•38 μm nylon mesh filter (cat. no. U-CMN-38-A).•30 mL Syringe .2PO4, and 21.8 g NaHCO3 in 800 mL distilled water. Bring to 1 L with Milli-Q water and filter sterilize. Store at 4°C.Dissolve 72.5 g NaCl, 2.1 g KCl, 1.5 g NaHAdd 10 mL of 10 × aCSF-B to 80 mL distilled water. Add 1 mL of 0.5 mg/ml leupeptin, 1 mL of 0.5 mg/ml aprotinin, 1 mL of 0.2 mg/ml pepstatin, 1 mL of 12 mg/ml pefabloc, 1 mL of 0.1 M EGTA, 1 mL of 0.5 M EDTA, and 1 mL of 0.5 M sodium fluoride. Bring to 100 mL with Milli-Q water and filter sterilize. Store at 4°C.Dissolve 3.03 g Trizma-base in 300 mL Milli-Q water and adjust pH to 7.6. Add 4.38 g NaCl and 10 g BSA. When dissolved bring to 500 mL with Milli-Q water and filter sterilize.Dissolve 38.14 g sodium borate in 300 mL Milli-Q water and heat to dissolve. Adjust pH to 9.0 with HCl. Bring to 500 mL with Milli-Q water and filter sterilize.Add 6.035 mL Ethanolamine  to 500 mL with Milli-Q water. Adjust pH to 8.0 with approximately 18 mL of 5N HCl and filter sterilize.Dissolve 3.755 g of glycine in 400 mL Milli-Q water. Adjust pH to 3.0 with HCl. Bring to 500 mL with Milli-Q water and filter sterilize.Dissolve 12.11 g Tris-base in 400 mL Milli-Q water. Adjust pH to 8.0 with HCl. Bring to 500 mL with Milli-Q water and filter sterilize.Dissolve 3.03 g Tris-base in 300 mL Milli-Q water and adjust pH to 7.6. Add 50 mL of 20% SDS and 3.5 mL β-mercaptoethanol. When dissolved bring to 500 mL with Milli-Q water and filter sterilize.Dissolve 1 g Congo red in 90 mL Milli-Q water, bring to 100 mL and filter sterilize.Dissolve 4.38 g NaCl in 300 mL Milli-Q water, add 2.5 mL 20% SDS. When dissolved bring to 500 mL with Milli-Q water and filter sterilize.Dissolve 4.84 g Tris-base in 200 mL Milli-Q water and adjust pH to 7.6, add 40 mL of 20% SDS. Mix thoroughly and split into 4, 50 mL lots placing each in a separate bottle. Add 82.15 g (1.2 M), 95.84 g (1.4 M), 109.54 g (1.6 M), and 123.23 g (1.8 M) sucrose into the four bottles, respectively. When dissolved, bring each solution to 200 mL with Milli-Q water.Dissolve 50 mg blue dextran in 25 mL of Milli-Q water and filter sterilize.Dissolve 3.953 g ammonium bicarbonate in 900 mL of Milli-Q water. Adjust pH to 8.5 with ammonium hydroxide. Bring to 1 L with Milli-Q water and filter sterilize.Dissolve one vial of 10 μg BDNF in 1 mL of PBS containing with 0.1% BSA. Mix thoroughly and aliquot into 100 μL vials. Store at −20°C.Dissolve one vial of 10 μg CNTF in 1 mL of PBS containing with 0.1% BSA. Mix thoroughly and aliquot into 100 μL vials. Store at −20°C.Dissolve one vial of 10 μg GDNF in 1 mL of PBS containing with 0.1% BSA. Mix thoroughly and aliquot into 100 μL vials. Store at −20°C.Dissolve one vial of 1 mg Y-27632 compound in 312.2 μL of DMSO. Filter sterilize and aliquot into 50 μL vials. Store at −20°C.Dissolve 1 g doxycycline hyclate (DOX) in 50 mL Milli-Q water. Filter sterilize and aliquot into 100 μL vials. Store at −20°C.Dissolve 20 g of D-(+)-Glucose in 100 mL of Milli-Q water. Filter sterilize and store at 4°C.Add 100 mL of mTeSR™1 5× Supplement and 5 mL of Penicillin/Streptomycin  to 400 mL of mTeSR™1 Basal Medium. Filter sterilize and store at 4°C.Add 75 mL of KnockOut Serum Replacement, 5 mL of MEM non-essential amino acids, 5 mL of GlutaMAX™ Supplement and 0.5 mL of β-mercaptoethanol to 414.5 mL of KnockOut™ DMEM. Filter sterilize, store at 4°C, and shield from light. This medium is designated as KSR base medium. Prior to use, take 10 mL of the KSR base medium and add 1 μL of DOX stock solution.Add 5 mL of GlutaMAX™ Supplement, 7.5 mL of 20% Dextrose stock solution and 5 mL of N2-supplement B to 482.5 mL of DMEM/F-12 Medium. Filter sterilize and store at 4°C. This medium is designated as N2B base medium. Prior to use, add 5 mL of the KSR base medium to 5 mL of the N2B base medium, and then add 10 μL of Puromycin and 1 μL of DOX stock solution.Prior to use, add 100 μL of B27 supplement, 10 μL of Puromycin and 1 μL of DOX stock solution to 10 mL of N2B base medium.Add 5 mL of GlutaMAX™ Supplement, 7.5 mL of 20% Dextrose stock solution and 2.5 mL MEM non-essential amino acids to 485 mL of Neurobasal™ Medium. Filter sterilize and store at 4°C.For 50 mL iN cell culture medium, add 1 mL of B27 supplement, 50 μL of BDNF, 50 μL of CNTF, 50 μL of GDNF, 50 μL of Puromycin, 5 μL of Doxycycline to 49 mL of Supplemental Neurobasal medium. Y-27632 is only required when plating iN cells, but not for iN maintenance.For 100 mL buffer exchange medium, add 2 mL of B27 supplement and 1 mL of GlutaMAX™ Supplement to 97 mL of Neurobasal™ Medium. Filter sterilize and store at 4°C. Note: since buffer-exchanged extracts will be applied to iN cultures all manipulations are done under sterile conditions.Extreme care should be taken when handling human tissue. Experimenters should wear appropriate personal protective equipment and consumable materials should be disinfected in bleach prior to disposable in hazardous waste. Where possible experimenters should work in an isolated area, and the area should be thoroughly clean at the end of each session.1)Before starting the procedure, set up tissue chopper  and pre-cool the cutting board on ice.2)Remove brain tissue from −80°C storage and thaw slightly on dry ice. Cut the brain tissue into half gram chunks using a razor blade. Carefully remove blood vessels and white matter, keep gray matter.3)Install pre-cooled cutting board on tissue chopper and chop brain tissue into ø 0.5 mm chunks. Divide chopped tissue in two: one half is used for Method 1 and the other for Method 2.Note: We here describe an example procedure appropriate to 20 g of dissected brain tissue. But we recommend adjusting experimental scales optionally for particular purposes. Brain tissue were obtained from four patients who both died with AD at Braak stage VI and temporal cortex used in the current study.1)Prepare two, 5 g lots of chopped brain tissue and transfer each into two separate 50 mL ice-cold falcon tubes. Add 25 mL (20% w/v) of ice-cold extraction buffer  to each tube and incubate with gentle side-to-side shaking at 4°C for 30 min.Note: Avoid harsh shaking. Thirty minutes incubation enables soluble proteins, including Aβ, to diffuse out of brain tissue.2)g and 4°C for 10 min, collect the upper 90% supernatant and transfer to clean ice-cold ultracentrifuge tubes.Centrifuge the suspension at 2,000 × 3)g and 4°C for 110 min using a SW41 Ti rotor. Carefully collect the upper 90% supernatant. The resulted sample is designated S extract.Centrifuge at 200,000 × Note: Low speed centrifugation described in step 2 prior to ultracentrifugation is used to minimize physical disruption of plaques.1)Add 50 mL (20% w/v) of ice-cold aCSF-B (pH 7.4) with protease inhibitors to 10 g of chopped brain tissue and homogenize with 25 strokes of a Teflon-glass Dounce homogenizer.2)g for 110 min and 4°C in a SW41 Ti rotor. Carefully remove the upper 80% supernatant. The resulted sample is designated H extract.Centrifuge at 200,000 × 1)During ultracentrifugation, prepare pre-cooled dialysis buffer  with a volume of at least 100-fold excess of designated brain extracts.2)Add 50 mL of pre-cooled dialysis buffer to a new dialysis cassette and leave at 4°C for at least 10 min.Note: Ensure the cassette is cold prior to adding brain extract and there is no buffer leaking.3)Remove dialysis buffer in the dialysis cassette and then add brain extract using a 10 mL pipette. Place cassette in the 5 L bucket with pre-cooled dialysis buffer and stir slowly for 24 h.4)Replace dialysis buffer. Repeat the above step 3 twice.® Red Glutamic Acid/Glutamate Oxidase Assay Kit  following manufacturer’s instructions.Note: We recommend dialysis against >100-fold excess of aCSF-B at 4°C, with three buffer changes over a 72 h period. The efficiency can be estimated by measuring the amount of glutamate in the final dialyzate vs. the starting extract . Typical5)Remove sample from the cassette using a 5 mL pipette and aliquot into 1.5 mL protein-low bind tubes. Freeze on dry ice and store at −80°C.Note: As Aβ are aggregation prone and sensitive to temperature changes, brain extracts can be thawed once and used. In general, remove samples from −80°C and allow to thaw at room temperature for 20 min.Antibodies are used to deplete brain extracts of Aβ to provide a negative control for bioactivity studies. Polyclonal antibodies are covalently cross-linked to Protein A sepharose (PAS) beads to minimize the potentially confounding effects of free antibody.1)Dissolve 0.25 g of PAS in 10 mL of 50 mM Tris-HCl, pH 7.6 containing 150 mM NaCl and 2% BSA. Incubate the PAS suspension with gentle side-to-side shaking at room temperature for 30 min.2)During incubation, use a Zeba spin desalting column to buffer exchange 3∼4 mg of protein A purified anti-Aβ polyclonal antibody  or control antibody  into 50 mM Tris-HCl, pH 7.6 containing 150 mM NaCl.Note: Pre-immune serum is a negative control to anti-Aβ polyclonal antibody and does not recognized Aβ. If polyclonal antibody such as S97 is not available, a commercial antibody 4G8 is optional .3)Centrifuge PAS suspension at 3,000 rpm for 5 min at room temperature and carefully discard supernatant.4)Keep the PAS pellet (∼1 mL from 0.25 g dried PAS) and add 3 mg anti-Aβ polyclonal antibody. Leave stand for 1 h at room temperature and then incubate with gentle side-to-side shaking at room temperature for 1 h.5)Centrifuge at 3,000 rpm for 5 min at room temperature and discard the supernatant. Wash PAS pellet with 10 mL of antibody coupling buffer . Centrifuge and discard supernatant. Repeat one time.6)Resuspend PAS with 10 mL of antibody coupling buffer and add 50 mg of DMP, then incubate with gentle side-to-side shaking at room temperature for 30 min.Note: This coupling reaction requires pH > 8.3, adjust with NaOH if necessary.7)Centrifuge at 3,000 rpm for 5 min at room temperature and discard supernatant. Resuspend PAS pellet with 10 mL of 0.2 M ethanolamine.8)Centrifuge at 3,000 rpm for 5 min at room temperature and discard supernatant. Resuspend PAS pellet with 10 mL of 0.2 M ethanolamine and incubate with gentle side-to-side shaking at room temperature for 2 h.9)Centrifuge at 3,000 rpm for 5 min at room temperature and discard supernatant. Resuspend PAS pellet with 1.75 mL PBS and transfer the suspension into two 2 mL Eppendorf tubes.10)Centrifuge at 3,000 rpm for 5 min at room temperature and discard supernatant. Resuspend PAS pellet with 1.75 mL of 100 mM glycine for a minimal time interval.11)Centrifuge at 10,000 rpm for 30 s at room temperature and discard supernatant. Resuspend PAS pellet with 1.75 mL of 100 mM Tris-HCl, pH 8.0.12)3, yielding a 50% S97-PAS or PIS-PAS slurry.Centrifuge at 3,000 rpm for 5 min at room temperature and discard supernatant. Resuspend each tube of 0.5 mL PAS pellet with 0.5 mL PBS containing 0.02% NaNNote: Covalently conjugated antibodies can be stored at 4°C for up to 3 months and maintain high immunoreactivity.1)Thaw appropriate aliquots of brain extracts on ice. Add 30 uL S97-PAS [immunodepletion (ID)] or PIS-PAS [mock immunodepletion (mock-ID)] to each 0.5 mL brain extracts, gently mix by hand and then rotate at 4°C overnight.2)Centrifuge at 6,000 rpm for 5 min at 4°C, carefully collect supernatant using gel loading tip and transfer into new 1.5 mL pro-low bind tubes. Keep the pellet as IP’d material.3)Repeat the above step 2 twice. The final supernatant is designated as Aβ-ID’d or mock-ID’d brain extract.4)Analyze the IP’d material from each round by Western Blot if necessary.Note: We recommend using a combination of anti-Aβ monoclonal antibodies specific for Aβ40  and Aβ42  to probe blots.5)Measure Aβx-42 monomer concentration in Aβ-ID’d or mock-ID’d brain extracts using Aβx-42 immunoassay. Quantitate the efficiency of immunodepletion by comparing Aβ-ID’d and mock-ID’d brain extract. Perform assays using the Meso Scale Discovery (MSD) platform and reagents from Meso Scale .Note: The Aβx-42 immunoassay preferentially detects Aβ monomers ending at Ala42. It employs m266 (3 μg/mL) for capture and biotinylated 21F12 (0.4 μg/mL) for detection .6)Measure aggregated Aβx-42 in Aβ-ID’d or mock-ID’d brain extracts by first dissociating aggregate in chaotrop. Incubate 20 μL of extract with 50 μL of 7 M GuHCl overnight at 4°C. Thereafter dilute samples 1:10 with assay diluent so that the final GuHCl concentration is 0.5 M. Quantitate the efficiency of immunodepletion by comparing Aβ-ID’d and mock-ID’d brain extract.Note: Previously we have shown that Aβ aggregates in AD brain extracts can be dissociated with chaotrops such as GuHCl, and that incubation of aggregate-containing extracts with GuHCl allows their disassembly and quantitation using monomer-preferring immunoassays. To match the buffer composition of standards with samples, prepare monomeric stock of Aβ1–42 in assay diluent containing 0.5 M GuHCl.Amyloid plaque isolation is time-consuming and the distribution and abundance of amyloid plaques in different brains and in different areas of the same brain is highly variable. Therefore, we recommend conducting a less time-consuming preliminary screen which involves randomly selecting a small piece of tissue from different areas, and comparing the abundance of Congo red stained plaques using light microscopy. Thus with this approach we identify regions of brain most likely to give good yields of plaque. Here we show an example of using temporal cortex from an end-stage AD patient.1)Remove brain tissue from −80°C storage place on dry ice and allow to thaw slightly. Dissect frozen cortical tissue with a scalpel blade and selecting ∼100 mg aliquots of tissue from the different areas of the available tissue.2)Mince tissue aliquots with a scalpel and transfer to 1 mL homogenizing tube.3)Add 5 volumes of 2% SDS buffer and homogenize 20 strokes with a Teflon-glass Dounce homogenizer.4)Transfer the homogenate to labeled 2 mL low-bind tubes. Heat at 100°C for 5 min and then cool down to room temperature.5)Centrifuge at 10,000 rpm for 10 min at room temperature. Carefully remove supernatant using gel-loading tip attached to vacuum suction unit.6)Vortex pellet in remaining buffer and remove 5 μL using wide-orifice tip and place onto a slide marked with Super Pap Pen.7)Add 100 μL of 1% Congo red staining solution to each slide. Ensure the solution spread over sample.8)Incubate slide at room temperature for 30 min and mount with a coverslip.9)Visualize stained amyloid plaques by using a light microscope fitted with a polarizing filter. Count the number of birefringent cores. Based on this preliminary screen select the tissue that contain the largest number of Congo red positive plaques for plaque isolation.Note: Amyloid structures can be visualized using polarized light. Amyloid plaques have a characteristic Maltese cross appearance and undergo red/green birefringence when view under polarized light. This birefringence is most apparent when gently rotating the polarizing filter.In general, a large amount of cortical tissue  is required for the isolation of amyloid plaques. Here we describe an example procedure appropriate to 100 g of brain tissue.1)Slightly thaw brain tissue on dry ice and dissect tissue away visible blood vessels and white matter. Mince 50 g of gray matter with a scalpel and then transfer to a 500 mL glass beaker.2)Add 250 mL of plaque extraction buffer and incubate at room temperature for 2 h without shaking.Note: During the incubation period, set water bath to 100°C and the bench-top centrifuge to 26°C.3)Homogenize the suspension with 20 strokes with a Teflon-glass Dounce homogenizer. Pool homogenates (approximately 300 mL from 50 g of dissected gray matter) from the same subject into a 500 mL glass beaker.4)Place the homogenates in 100°C water bath for 15 min and then cool to room temperature.5)Pass the homogenates through 112 um nylon mesh, transfer the flow through into six 50 mL tubes.6)g for 30 min at 26°C using a Bench top centrifuge.Centrifuge at 300 × 7)Remove the supernatant leaving at least 15 mL liquid above the soft pellet.Note: Critical step. Avoid disrupting the soft pellet.8)Combine the material from 6 tubes into 3 tubes. Rinse the emptied tubes three times with 5 mL 0.1% SDS solution and combine all materials together. Bring the content of each tube to 50 mL with 0.1% SDS solution.Note: At this stage, samples are quite sticky, thus we recommend rinsing emptied tubes thoroughly and retain all the materials.9)g for 30 min at 26°C using a Bench top centrifuge.Centrifuge at 300 × (10) Remove the supernatant leaving at least 10 mL liquid above the pellet.Note: The pellet should be more compact than that in step 7, but still needs to avoid disruption.11)Combine the material from 3 tubes to 2 tubes. Rinse emptied tubes in the same way as described in step 8 and bring each tube to 50 mL with 0.1% SDS solution.12)g for 30 min at 26°C using a benchtop centrifuge.Centrifuge at 300 × 13)Remove the supernatant leaving at least 5 mL liquid above the pellet.14)Combine the material from 2 tubes to 1 tube. Rinse emptied tube in the say way as described in step 8 and bring to 50 mL with 0.1% SDS solution.15)g for 30 min at 26°C using a Bench top centrifuge.Centrifuge at 300 × 16)Remove the supernatant leaving 4 mL liquid above the pellet.17)Resuspend the material by homogenizing with 20 strokes of a Teflon-glass Dounce homogenizer.18)Pass the homogenate through 38 um Nylon mesh, collect flow through into a 50 mL tube. Rinse homogenizer tube with 1 mL 0.1% SDS solution, pass through the same filter and collect flow through. Repeat 2 more times.19)The above procedure typically yields ∼7–7.5 mL crude plaque materials from 50 g of gray matter. Store sample at 4°C overnight.1)Prepare a linear sucrose gradient in a 38.5 mL ultracentrifuge tube. First add 5 ml of 1.8 M sucrose solution to the bottom, then carefully add 8 mL of 1.6 M sucrose solution on top of the 1.8 M sucrose solution. Thereafter, add 8 mL of 1.4 M sucrose solution and 7 mL of 1.2 M sucrose solution on top of the former. Carefully apply 7.5 mL crude plaque suspension to the top of the gradient.2)g for 60 min at 26°C.Centrifuge in an SW28 rotor at 72,000 × Note: Ensure the centrifuge is set at 26°C as SDS will crystallize at lower temperature.3)Beginning at the top collect each layer and transfer to individual 50 mL tubes.Note: Typically most amyloid plaque are found in the 1.6 M sucrose layer. Nonetheless, we recommend collecting materials from all layers for further analysis.4)g for 30 min at 26°C, then transfer each pellet to a 1.5 mL Protein-low bind Eppendorf tube.Dilute each layer sample 1:5 with 0.1% SDS in 150 mM NaCl. Centrifuge at 300 × 5)g and room temperature for 10 min. Discard supernatant and keep 100 uL liquid above the pellet. Add 1 mL filtered MQ water into each tube and vortex thoroughly.Centrifuge at 12,000 × 6)Repeat step 6 one more time.7)g and room temperature for 10 min. Discard supernatant and resuspend pelleted plaques with 100 uL of MQ water. Take 10 uL of this material, stain with 0.2% Congo red and visualize using polarized microscopy (see section 2.2. Equipment).Centrifuge at 12,000 × 8)Freeze the remaining material and lyophilize.9)g and room temperature for 10 min.To concentrate lyophilizate at the bottom of the tube centrifuge at 12,000 × Note: The weight of isolated plaques can be estimated by subtracting the weight of pre-weighed empty tubes from the weight of tubes containing plaques.1)Add 1.2 mL of 88% formic acid to each vial of lyophilized plaques, vortex and incubate at room temperature overnight, e.g., 12–14 h, with gentle agitation.Note: Do not incubate plaques in formic acid longer than 24 h.2)g for 15 min at room temperature and carefully transfer the upper 90% of supernatant into a 1.5 mL Protein-low bind Eppendorf tube.Centrifuge at 12,000 × 3)Equilibrate Superdex 75 10/300 GL size exclusion column (SEC) in 50 mM Ammonium Bicarbonate, pH 8.5 at 0.4 mL/min.4)Inject 1 mL of blue dextran solution and collect 0.5 mL fractions. The peak fraction containing blue dextran is designated as “void volume” and labeled as Fx0. Equilibrate system with 2 column volumes (CV) of Mobile Phase solution.5)Inject 1 mL of Gel filtration standard (Bio-Rad) and calibrate standards elution by using UV detector. Equilibrate system with 2 CVs of Mobile Phase solution.6)Inject 1 mL solubilized plaque material and collect 0.5 mL fractions from Fx-1 to 17. Take 5 uL of each fraction and measure Aβ by Western Blot as described previously . Store r1)5 per well in mTeSR media.Plate human iPS cells on Matrigel coated 12-well plate with 3.8 × 102)On the day of viral infection, prepare 1 mL of fresh mTeSR media containing pTet-O-NGN2-puro lentivirus, Tet-O-FUW-eGFP lentivirus and Fudelta GW-rtTA lentivirus. Remove cell culture medium and add lentivirus-containing medium.Note: Lentiviruses were prepared using the plasmids from Addgene  as showed previously . The eff3)After 24 h, remove lentivirus-containing medium. Dissociate cells with Accutase and resuspend with mTeSR supplemented with Y-27632. Transfer cells to 10 cm dishes and feed with fresh mTeSR media every day. When cells reach >80% confluent, split for expansion or freeze down.Note: To avoid batch effect of lentivirus infection, we recommend freezing down a large number of cryovials of transduced iPS cells. Transduced iPS cells can be passaged no more than 10 times to maintain high efficiency of neuron differentiation.4)6 transduced iPS cells using mTeSR with Y-27632 on Matrigel coated 10 cm dish. Feed with 7 mL fresh mTeSR media every day until cells reach 50% confluence.Plate 4 × 105)On day 1, remove cell culture medium and add 7 mL KSR media containing with 2 μg/mL Doxycycline (iN induction medium A).Note: At this stage, Ngn2 and GFP expression are induced.6)On day 2, replace cell culture medium with KSR:N2B (1:1) media containing with 5 μg/mL Puromycin and 2 μg/mL Doxycycline (iN induction medium B).Note: At this stage, cells without Ngn2/Puromycin resistance would die off.7)On day 3, replace cell culture medium with N2B media containing with 1% B27, 5 μg/mL Puromycin and 2 μg/mL Doxycycline (iN induction medium C).8)On day 4, split cells for neuron differentiation or freeze down. For freezing down, wash cells with warm PBS and detach with Accutase. Resuspend with freezing medium and store in liquid nitrogen.Note: The neuronal differentiation efficiency may differ from batch to batch. To minimize batch effect, we recommend freezing down a large down a large number of cryovials of iN4 (induced neuron at day 4) and using the same batch of cells in one study.9)4 iN4 cells/cm2 using iN culture media with Y-27362 onto Matrigel coated cell culture vessels. Feed cells with iN culture media.For neuron differentiation, plate 1.5 × 10Note: Select appropriate cell culture vessels according to practical requirements. When changing iN culture medium, we recommend removing half of the medium and add the same volume of fresh medium. Ara-C is optional if divided cells are seen in the iN culture.10)Change iN culture media every 3∼4 days. In our preliminary experiments, iN cells mature at iN21, i.e., 17 days after plating, and can be used throughout a time window (approximately a week).1)3 iN4 cells onto Matrigel coated 96-well plate. Feed cells with iN culture medium every 3–4 days.For Aβ toxicity measurement using live-cell imaging system, plate 5 × 10Note: To minimize the effect of evaporation cell culture medium from neurons, we recommend plating iN cells into the inner 60 wells of 96-well plate and adding PBS into the outer wells.2)Approximately 7 h prior to treatment, place cell culture plates into IncuCyte live-cell imaging instrument. Set up scanning procedure following manufacturer’s instructions. For long-term recording, schedule repeating scans every 2 h for up to 96 h.Note: We recommend recording iN cells 7 h prior to treatment. This allows acquisition of three baseline scans which provide a solid reference for time course studies. During this period Aβ-containing human brain extracts are buffer-exchanged into neurobasal medium supplemented with B27/GlutaMAX.3)Buffer exchange is accomplished using a 5 mL HiTrap desalting column and a peristaltic pump. Wash the column with 25 ml of 20% Ethanol and then 25 mL of filtered MQ water. Then equilibrate with 25 mL of neurobasal medium supplemented with B27/GlutaMAX.4)Apply 0.5 mL of human brain extract using a 1 mL syringe at a flow rate of ∼1 mL/min. Elute proteins with iN culture medium and collect eight 0.5 mL fractions.5)Remove 50 μL of each fraction and measure Aβ concentration by ELISA. Pool fractions 4 and 5  and use for further toxicity assays.Note: Prior studies indicate that fractions 4 and 5 consistently contain the peak Aβ fractions and exclude low molecular weight brain-derived compounds. Confirmatory ELISA analysis is done on the day after iN cell treatment.6)During the time after the 3rd baseline scan, remove cell culture plates from IncuCyte instrument. Remove approximately half of cell culture medium leaving ∼100 μL in each well. Then add 50 μL of fresh iN culture medium and 50 μL of buffer-exchanged human brain extract. This results in a 1:8 dilution of the original brain extract.Note: If the scan interval is 2 h, we should be cautious of the time window that allows us to complete the sample application. In fact, the scan interval is optional depending on practical requirements and manipulation feasibility.7)Place treated cells into IncuCyte instrument and continue recording for 3∼4 days.8)When experiment is completed, select representative images from different wells to a new NeuroTrack image collection. Launch New Processing Definition and adjust parameters following manufacturer’s instructions to define neurite processes and cell bodies based on phase contrast images. Once the training is completed, save the processing definition as a confirmed analysis template.2; Neurite Filtering–Best; Neurite sensitivity–0.4; Neurite Width–2 μm.Note: Typical settings were: Segmentation Mode–Brightness; Segmentation Adjustment–1.2; Cell body cluster filter–minimum 500 μm9)Launch new analysis job to the recorded database of the whole plate using the above confirmed analysis template.Note: The training of processing definition can be conducted using the baseline scan data. Thus, we can launch real-time analysis job and monitor the scheduled scanning to the whole cell culture plate. Alternatively, an analysis job can be launched when the experiment is completed or even during the scheduled scanning.10)Export Metric Graph/data and quantify neurite length and number of branch points. The neurotoxicity of Aβ is determined to the average value measured during the 6 h period prior to sample addition.post-hoc tests or Student’s t-tests using GraphPad Prism 8.Figures showing MSD Aβ immunoassay and live-cell IncuCyte imaging data are representative of at least three independent experiments. Differences between groups were tested with one-way analysis of variance (ANOVA) with Bonferroni We have used the above protocols to isolate and characterize Aβ from multiple AD brains–but here we show only a few representative cases. The amount of Aβ extracted by Method I (gently soaking minced tissue in aqueous buffer–S extract) and Method II  was measured using highly sensitive Aβ immunoassays. Typically, S extracts contain substantially less Aβ than H extracts . Since AHuman brain extracts may vary in their content of bioactive small molecules such as neurotransmitters and/or pharmacologic agents, thus it is important to take precautions to remove any such material prior to assessing bioactivity. For this reason we dialyzed brain extracts immediately after preparation. For routine analysis we measured glutamate levels in human brain extracts before and after dialysis. We found that both H and S extracts contained ∼300 μM of glutamate, and that >90% of the starting concentration of glutamate was removed by three rounds of dialysis . Prior sTo determine whether the neuritotoxicity induced by AD brain extracts was mediated by Aβ, we removed Aβ using a pan anti-Aβ polyclonal antibody, . S97 readily removes >70% of Aβx-42 from S extracts . Consistp < 0.001) or monomers (p < 0.001)  . Prior a< 0.001) .Although not shown in the current study we have previously used brains from non-Alzheimer subjects as a control  and founIn contrast to the thousands of studies that have used synthetic Aβ, rodent neurons, or irrelevant cell lines and measures of cell death , we pionMethod III enables isolation of highly pure Aβ dimers and monomers from amyloid plaques. While we have shown dimers to have disease relevant activity , certainIn future studies, CRISPR-based approaches could be employed to interrogate pathways of both Aβ toxicity and synaptogenic effects. Thus, the combination of the three extraction methods described and the further adaptation of our bioassay should enable a deeper understanding of the positive and negative activities of Aβ and may facilitate discovery of agents highly specific for the most noxious forms of Aβ.Limitations of our approach include the requirement for large amounts of postmortem brain tissue, and relatively long duration experiments. Typically, preparation and characterization of brain extracts and/or plaque-derived Aβ takes on the order of 2 weeks. The bioassay is medium throughput and requires data collection periods of 2–4 days. To exclude artifacts arising due to postmortem delays and/or freeze-thawing of tissue, for certain studies it will be important to investigate the use of fresh biopsy tissue. Nonetheless, other investigators have already employed our experimental paradigms  and we eThe original contributions presented in this study are included in the article, further inquiries can be directed to the corresponding author.This study was reviewed and approved by Partners IRB. Human specimens were obtained from the Massachusetts ADRC Neuropathology Core, Massachusetts General Hospital and used in accordance with the Partners Institutional Review Board . Written informed consent was obtained from the individual(s) to participate in this study.DMW conceived the research. DMW and WH designed the protocols and wrote the manuscript. WH performed the biochemical and IncuCyte experiments, analyzed the data, and prepared the figures. WL cultured iPSC-derived neurons and assisted with biochemical experiments. WL and AD isolated and characterized the amyloid plaques. TY-P provided the viral-infected iPS cells. All authors critically reviewed the manuscript.


After the successful characterization of the prepared nanoparticles by various techniques such as FT-IR, FE-SEM, EDX/MAP, XRD, TEM, TGA, VSM, and ICP-OES, the catalytic efficiency of them were evaluated in the cyanation reaction of aryl halides in the presence of nitromethane as a non-toxic and cost-effective cyanation source. The cyanation products were obtained in desirable yields. Notably, the magnetic nanocatalyst can be easily recovered and reused at least five times without a significant decrease in its performance.In the current study, a highly efficient heterogeneous copper catalyst has been developed by supporting copper acetate on a magnetically separable triazine–vitamin B 5. Its catalytic efficiency was evaluated in the cyanation of aryl halides with nitromethane as a non-toxic and cost-effective cyanating source.An efficient heterogeneous copper catalyst was developed by supporting copper acetate on magnetically separable triazine–vitamin B The diazotization of anilines followed by the Sandmeyer reaction3 and also the Rosenmund–von Braun reaction4 have been known as classic synthetic methods toward aryl nitriles. Since then, a series of transition metal-catalyzed protocols have been reported for the cyanation reactions by using diverse cyano sources such as KCN,5 NaCN,6 Zn(CN)2,7 CuCN,8 and TMSCN.9 Nevertheless, most of them are confronted with significant drawbacks including the use of toxic metal cyanides in stoichiometric amounts and the requirement of harsh reaction conditions. To overcome this problems, several less-toxic metal-free cyanide sources including malononitrile,10 butyronitrile,11 acetonitrile,12 benzyl cyanide,13 AIBN,14etc. have been disclosed. Furthermore, some other organic compounds, which can generate cyano groups in situ have been employed as indirect cyanation sources. Explored examples of such cyanation reagents include NH4I/DMF,15 NH4HCO3/DMSO,16 NH4HCO3/DMF,17t-BuNC,18etc. Recently, nitromethane has also been introduced as a potentially valuable indirect organic cyano source for the cyanation of organic scaffolds, because of its chemical stability, and low-cost commercial availability. Nevertheless, there are very few reports on the use of nitromethane as a cyanation reagent in the field of metal-catalyzed aromatic CCN bond formation. So, it is still demanding to make attempts in this direction.19Aromatic nitriles are an important class of versatile scaffolds present in natural compounds and many synthetic organic products, such as dyes, pharmaceuticals, agrochemicals, and herbicides.20 Co,21 Ni,22 Cu,10 Rh23 and Zn24 have been utilized to catalyze cyanation reactions of aryls and aryl halides using different cyanation sources. Among them, the use of Cu as a readily available and inexpensive transition metal can be desirable.17,19 However, most of these reported metal-catalyzed cyanation reactions have adopted a homogeneous approach, which suffer from serious limitations including high expenses, problematic separation (in some cases non-separable), and non-reusability of metal catalysts. To meet these restrictions as well as minimize possible pollution caused by toxic metal discharge in some homogeneous protocols, a wide range of heterogeneous metal-incorporating catalytic systems have been developed for cyanation transformations.25 Magnetic nanoparticles have been well known as one of the most promising heterogeneous nanostructures, which have unique physicochemical features in addition to being easily recoverable and reusable.26Over the recent past decades, various transition metals, such as Pd,27–30,19 we chose vitamin B5 as a suitable ligand for this reaction.Until now, various reports have been presented about the use of ligands in the cyanidation reaction, and by examining the used ligands and the importance of ligands containing nitrogen and oxygen in binding to various metals and forming stable complexes,5 system for the synthesis of aryl nitriles from aryl halide substrates in presence of nitromethane.To the best of our knowledge, no heterogeneous copper catalyst has been reported for the cyanation of aryl halides using nitromethane as an indirect organic nitrile source. In this work, we designed and synthesized a novel heterogeneous copper-incorporating nanocatalyst by loading copper acetate on a magnetically recyclable triazine–vitamin B22.1.−1 in the range of 25–800 °C under the following air.All required materials were purchased from Aldrich and Merck companies and were used without any further purification. The reaction was monitored by thin-layer chromatography. TLC was performed on glass plates incorporated with silica-gel 60 F-254. Infrared spectrums (IR) were determined by grade KBr on a Nicolet FT-IR 100 spectrometer. X-ray diffraction (XRD) data were obtained at room temperature by a Philips X-pert 1710. The size and morphology of the nanoparticles were determined by scanning electron microscopy (SEM) using a TESCAN MIRA III FE-SEM. Also, the elemental composition of the nanoparticles was studied by energy-dispersive X-ray (EDAX). Transmission electron microscopy (TEM) was performed using a Philips CM 120 at 120 kV. Thermal gravimetric analysis (TGA) was performed by a thermal analyzer with a heating rate of 10 °C min2.2.5 in anhydrous THF (5 mL) and ultrasonicated at 60 °C. Potassium carbonate (1 mmol or 0.138 g) was added to the reaction mixture, and the solution was kept under the same conditions for 3 h and then stirred for 12 h at 70 °C. Finally, the reaction mixture was centrifuged, and the product was separated, washed with anhydrous THF, and dried in a vacuum oven at 70 °C.Cyanuric chloride (1 mmol or 0.18 g) was dissolved in 5 mL of anhydrous tetrahydrofuran (THF). The resulting solution was then slowly added to a solution of vitamin B2.3.3·6H2O (5 mmol or 1.35 g) and FeCl2·4H2O (2.5 mmol or 0.5 g) in 50 mL deionized water was ultrasonicated for 10 min to be completely dispersed. While the solution was being stirred at room temperature, 20 mL of ammonia solution (37%) was added dropwise until reaching pH = 11. The solution was then stirred under a nitrogen atmosphere at 80 °C for 5 h. Thereafter, the solution was cooled down to 40 °C, 40 mL of ethanol was added, and the reaction mixture was stirred for 30 min. Tetraethyl orthosilicate (10 mL) was gradually added to the reaction mixture and it was then placed under a nitrogen atmosphere at 40 °C for 22 h. The Fe3O4@SiO2 product was separated by applying an external magnetic field, washed several times with water and ethanol, and dried in a vacuum oven at 80 °C.A mixture of FeCl2.4.3O4@SiO2, and the solution was ultrasonicated for 15 min to be completely dispersed. Then, (3-chloropropyl)trimethoxysilane (10 mL) was added to the dispersion, and the mixture was stirred at 60 °C for 18 h under a nitrogen atmosphere. The nanoparticles were separated by applying an external magnetic field, washed several times with anhydrous toluene, and dried in a vacuum oven at 80 °C. First, potassium iodide (2 mmol or 0.332 g), then potassium carbonate (2 mmol or 0.276 g), and, finally, TA/B5 (0.2 g) were added to Fe3O4@SiO2–Cl (0.2 g) dispersed in 10 mL acetonitrile, and the mixture was refluxed for 12 h. The nanoparticles were then separated by applying an external magnetic field, washed several times with water and ethanol, and dried in a vacuum oven at 80 °C.Anhydrous toluene (50 mL) was added to 0.2 g Fe2.5.3O4@SiO2–TCT/B5 (0.2 g) was dispersed in 50 mL ethanol through 15 min ultrasonication. Next, 1 mmol of copper(ii) acetate monohydrate dissolved in ethanol was added dropwise to the dispersion. The resulting mixture was then refluxed for 16 h. The nanoparticles are separated by applying an external electric field, washed several times with ethanol, and dried in a vacuum oven at 60 °C.Fe2.6.First, 4-iodoanisole (1 mmol or 0.234 g), nitromethane (3 mmol or 0.183 g), potassium carbonate (1 mmol or 0.138 g), and the catalyst (30 mg) were weighed and poured into a glass tube. Then, 3 mL dimethyl sulfoxide (solvent) was added and the mixture was stirred at 100 °C for 12 h. Thin-layer chromatography was used to monitor the progress of the reaction. Upon the completion of the reaction, the reaction mixture was cooled down to room temperature, and the catalyst was removed by a magnet. The reaction mixture was then extracted by water and ethyl acetate. After any remaining water droplets were removed from the organic phase by using anhydrous sodium sulfate, the solvent was evaporated under a vacuum. Finally, the purification process was carried out using silica gel plates and ethyl acetate : hexane (1 : 9) solvent.33.1.3O4@SiO2–TCT/B5–Cu(ii) nanocatalyst is illustrated in The preparation process of the proposed Fe3.2.3.2.1.3O4@SiO2, TCT/B5, and Fe3O4@SiO2–TCT/B5–Cu(ii) are shown in 3O4@SiO2, the stretching and bending vibrations of the O–H bond are observed at around 3429 cm−1 and 1632 cm−1, respectively. Also, peaks appearing at 1095 cm−1 and 466 cm−1 are associated with the stretching and bending vibrational modes of Si–O–Si, respectively. In addition, the characteristic peak that appeared at 574 cm−1 can be related to the Fe–O stretching. In the FT-IR spectrum of TCT/B5, the bands located at 1717 cm−1, 1634, cm−1 and 1602 cm−1 can be assigned to ester, acid and amid carbonyl groups, respectively. The appearance of a characteristic peak at 1717 cm−1 can be ascribed to ester, indicating the formation of the covalent bond between TCT and vitamin B5. There are two absorption bands in the area of 1400–1460 cm−1 and 1100–1290 cm−1, which are associated with the stretching vibrations of the C–N bond and C–O bond, respectively. Also, peaks observed at around 3422 cm−1 and 2925 cm−1 are allocated to O–H and C–H aliphatic stretching vibrations, respectively. The characteristic peaks in the FT-IR spectrum of Fe3O4@SiO2–TCT/B5–Cu(ii) are mainly observed at 3406 cm−1, 1577 cm−1, 1440 cm−1, 1092 cm−1, and 465 cm−1, which can be respectively related to the stretching vibrations of O–H, C PBM data was replaced with SVG by xgml2pxml:<glyph-data id="z.dbd" format="PBM" resolution="300" x-size="8" y-size="10" xml:space="preserve">00000000000000000000000000000000111111110000000011111111000000000000000000000000</glyph-data><svg xmlns="http://www.w3.org/2000/svg" version="1.0" width="13.200000pt" height="16.000000pt" viewBox="0 0 13.200000 16.000000" preserveAspectRatio="xMidYMid meet"><metadata>Created by potrace 1.16, written by Peter Selinger 2001-2019</metadata><g transform="translate scale" fill="currentColor" stroke="none"><path d="M0 440 l0 -40 320 0 320 0 0 40 0 40 -320 0 -320 0 0 -40z M0 280 l0 -40 320 0 320 0 0 40 0 40 -320 0 -320 0 0 -40z"/></g></svg>N triazine ring, C–N, Si–O–Si and Fe–O bonds, indicating the successful attachment of TCT/B5 to the support. In addition, due to the coordination of some ligand groups with Cu(ii) species, the intensity of some TCT/B5 peaks in the final catalyst has decreased.FT-IR spectra of Fe3.2.2.5 and Fe3O4@SiO2–TCT/B5–Cu(ii) were depicted in 5, there are peaks to those of ligand structure in the final catalyst XRD pattern. As illustrated in the XRD pattern of Fe3O4@SiO2–TCT/B5–Cu(ii), the diffraction peaks at 2θ values of 30.74°, 35.89°, 43.44°, 53.94°, 57.64° and 63.24° correspond to (220), (311), (400), (422), (422), (511) and (440) planes of Fe3O4, which can be readily indexed to the standard spinel cubic magnetite (JCPDS no. 19-0629). Also, the broad peak at 2θ = 20 to 30 can be assigned to the amorphous SiO2 phase. The diffraction peaks at 2θ values of 29.54° and 32.19° can be attributed to ligand structure in the final catalyst. As shown in the XRD pattern of TCT/B5, there are peaks similar to those of ligand structure in the final catalyst XRD pattern, confirming the success of ligand binding to the magnetic support.XRD results of TCT/B3.2.3.5 and Fe3O4@SiO2–TCT/B5–Cu(ii) were depicted in 5 nanoparticles are porous and irregular in shape. The TCT/B5 nanoparticles have mean diameter of 36 nm. The morphology of the prepared Fe3O4@SiO2–TCT/B5–Cu(ii) nanoparticles is a combination of the spherical Fe3O4@SiO2 nanoparticles with the porous and irregular structure of the TCT/B5 ligand with mean diameter of 38 nm, which is well showed in The morphology and size of nanocatalysts were investigated by FE-SEM analysis. The SEM images of TCT/B3.2.4.5 and Cu(ii) have a uniform distribution on the surface of Fe3O4@SiO2.EDAX analysis confirmed that the final catalyst contains the expected elements of C, O, N, Fe, Si, and Cu with wt% of 7.89, 49.77, 4.27, 15.27, 16.38 and 6.42, respectively. Additionally, from EDAX element mapping images  can be a3.2.5.3O4@SiO2–TCT/B5–Cu(ii) have been presented in 3O4@SiO2–TCT/B5–Cu(ii), which can be attributed to the ligand and Fe3O4 or Cu(ii) NPs. The magnetic core can be seen as a dark spot inside the bright, spherical Cu(ii) triazine-based complex, according to the observation from the TEM images.The TEM images of Fe3.2.6.5 and Fe3O4@SiO2–TCT/B5–Cu(ii) nanoparticles were estimated by TGA-DTG analysis. The TGA of TCT/B5 , the prepared nanoparticles showed higher stability than TCT/B5. The first main weight loss of Fe3O4@SiO2–TCT/B5–Cu(ii) (about 6%) was observed in the range of 220–340 °C, indicating the degradation of organic contents. The second one (mass loss of 5%) was observed in the range of 460–550 °C, which can be attributed to the decomposition of the remaining organic moieties and oxidation of carbon.The thermal degradation and thermal stability of TCT/Bf TCT/B5  showed t3.3.3O4@SiO2–TCT/B5–Cu(ii) catalyst, we studied its catalytic performance in the cyanation reaction of aryl halides using CH3NO2 as the cyanide resource . DMSO was chosen as the best solvent. Comparison between different bases used (entries 10–13), exhibited that K2CO3 has the best efficiency to form the cyanated product. Based on the catalyst loading study, various amounts of catalyst were tested in this reaction. The amount of 30 mg was found to be the best value of catalyst for high conversion under the reaction conditions. The model reaction in presence of lower values of the catalyst resulted in a decrease in the yield of the cyanation product (entries 14 and 15). Decreasing the reaction temperature to 90 °C brought down the yield to 81% (entry 18). By increasing the temperature to 110 °C (entry 17), no change in the yield of the product was observed. The cyanation reaction was carried out in the presence of copper on substrates Fe3O4 and Fe3O4@SiO2. These substrates are free of vitamin B5 to investigate the importance of the ligand in this reaction. According to the results, the importance of vitamin B5 in the prepared composite is concluded. The presence of vitamin B5 in the structure of catalyst is essential and enhances efficiency (entries 20 and 21).After characterization of the Feresource . In ordereaction . First, O-substituted aryl halides  gave lower yields compared to P-substitutes due to the hindrance effect.In order to evaluate the role of substituent groups, various aryl halides bearing both electron-rich and electron-poor groups were tested in this reaction . Under t3O4@SiO2–TCT/B5–Cu(ii) and effectiveness of the present approach were compared with the previous researches. As shown in 3O4@SiO2–TCT/B5–Cu(ii) catalyst.The catalyst efficiency of Fe3.4.19 Thermochemical studies show that CH3NO2 can be decomposed into CO and NO at high temperatures. The released NO in the reaction medium interacts with Cu(ii) in the catalyst and reduces Cu(ii) to Cu(i). The 16-electron Cu(i) complex undergoes an oxidative addition reaction with aryl halide to form complex A. The 18-electron A complex performs ligand exchange in presence of HCN (generated from CH3NO2 using base) and produces B complex. Finally, complex B after a reductive elimination affords the cyanation product and the catalyst returns to the catalytic cycle  which did not show a significant decrease compared to that of the original catalyst.We examined the recyclability and reusability of our catalyst in the cyanation reaction of 4-ome-iodobenzene in presence of CH43O4@SiO2–TCT/B5–Cu(ii) nanocatalyst and nitromethane as a non-toxic, inexpensive and readily available cyanating source has been developed. The aryl nitriles were obtained in moderate to good yields. The prepared heterogeneous nanocatalyst can be readily separated from the reaction mixture by magnetic filtration and recovered for at least five cycles without a significant loss of its activity.In conclusion, a novel and efficient catalytic approach for the synthesis of aryl nitriles in presence of FeThis research does not involve human participants and/or animals.The data that supports the findings of this study are available in the ESIConceptualization: Farzaneh Karimi, Masoumeh Jadidi Nejad; methodology: Farzaneh Karimi, Masoumeh Jadidi Nejad; formal analysis and investigation: Farzaneh Karimi and Masoumeh Jadidi Nejad; writing-original draft preparation: Masoumeh Jadidi Nejad, Arefe Salamatmanesh; writing-review and editing: Arefe Salamatmanesh, Masoumeh Jadidi Nejad; funding acquisition: Akbar Heydari; resources: Akbar Heydari; supervision: Akbar Heydari.There are no conflicts of interest to declare.RA-013-D2RA06104J-s001