TY - JOUR
AU - H. Sajedi,, Reza
AB - Abstract Amyloid-β (Aβ) peptide and tau protein are two hallmark proteins in Alzheimer's disease (AD); however, the parameters, which mediate the abnormal aggregation of Aβ and tau, have not been fully discovered. Here, we have provided an optimum method to purify tau protein isoform 1N4R by using nickel-nitrilotriacetic acid agarose chromatography under denaturing condition. The biochemical and biophysical properties of the purified protein were further characterized using in vitro tau filament assembly, tubulin polymerization assay, circular dichroism (CD) spectroscopy and atomic force microscopy. Afterwards, we investigated the effect of tau protein on aggregation of Aβ (25–35) peptide using microscopic imaging and cell viability assay. Incubation of tau at physiologic and supra-physiologic concentrations with Aβ25–35 for 40 days under reducing and non-reducing conditions revealed formation of two types of aggregates with distinct morphologies and dimensions. In non-reducing condition, the co-incubated sample showed granular aggregates, while in reducing condition, they formed annular protofibrils. Results from cell viability assay revealed the increased cell viability for the co-incubated sample. Therefore, the disassembling action shown by tau protein on Aβ25–35 suggests the possibility that tau may have a protective role in preventing Aβ peptide from acquiring the cytotoxic, aggregated form against oxidative stress damages. Open in new tabDownload slide Open in new tabDownload slide Aβ25–35, Alzheimer’s disease, fibrillation, intrinsically disordered protein, tau purification Alzheimer’s disease (AD) is neuropathologically characterized by the presence of two protein deposits, the extracellular senile plaques (SP) of amyloid-β (Aβ) peptide, a derivative of the membrane protein amyloid precursor protein and the flame-shaped neurofibrillary tangles (NFTs) of the abnormally phosphorylated protein tau (1, 2). The anatomical distribution of neuritic plaques is somewhat variable and does not coincide with the distribution of NFTs (3). Moreover, Aβ pathology appears to begin in the cortex and spread inward, while tau pathology exhibits an opposite progression (4). Accordingly, while Aβ and tau pathology are anatomically separate in the AD brain, previous evidences indicated that Aβ could still trigger tau pathology and neurodegeneration in regions with minimal fibrillar Aβ pathology (4). According to a report by Rapoport et al., neuronal cells expressing tau protein degenerated in the presence of Aβ aggregates, suggesting a direct link between the lesions (5). Therefore, it is critical to determine which parameters mediate the abnormal aggregation of Aβ and tau. In normal healthy conditions, Aβ is present in low levels (pM), and exerts its physiological functions, including cellular trophic activity with implications in neurodevelopment, memory and learning. In contrast, in pathological conditions, the concentrations of Aβ peptide increase (nM to μM), and this switches it normal functions to pathological effects, causing neurotoxicity and cell death (6). The Aβ peptide has a partially hydrophobic character and shows a high propensity in fibril formation with a cross-β-substructure (7); however, the building principles of the fibres are still under investigation (8). The normal Aβ peptides with predominately α-helical structures convert into insoluble fibrillar aggregates with a high β-sheet content during the early stages of amyloid formation in AD (9, 10). This phenomenon converts the Aβ peptides from non-toxic or less toxic structures to the neurotoxic species (10). The 25–35 fragment of the Alzheimer Aβ peptide, Aβ (25–35), represents the smallest most toxic derivative of the peptide, which retains the aggregation propensities of the full-length molecule, Aβ (1–40) (11, 12). It has been shown that the Aβ peptide promotes tau phosphorylation, which induces self-assembly of tau into NFTs, and initiates multiple pathways resulting in progressive degeneration and/or death of neurons (5, 13). Tau is a neuronal microtubule-associated protein, which is classified as a prototypical natively unfolded or intrinsically disordered protein (IDP). It involves in stabilizing microtubules that is critically important for axonal transport (14). Tau promotes tubulin assembly into microtubules, regulates microtubule dynamics and improves their stability (15). The function of tau is dependent on its phosphorylation state of more than 40 sites including 85 potential serine, threonine and tyrosine (16, 17). Tau inclusions are found in different types of cells in different tauopathies. In AD, tau inclusions are found in neurons (mainly in axons) as NFT, while in many other tauopathies, the inclusions are found both in neurons and glia cells (especially in oligodendrocytes) (18). Moreover, the compositions of the inclusions are different. In many tauopathies, the inclusions are mainly composed by 4R-tau, while in Pick's disease, 3R-tau is the predominant form. In AD, the overall ratio of 3R/4R tau isoforms is equal albeit the expression favours 2N4R (18). The physiological levels of full-length tau are in the range of 1–10 µM (19). Khatoon et al. found that the levels of total tau were about 8-fold higher in AD than in control cases, and this increase was in the form of the abnormally phosphorylated protein (20). In AD, the normal function of tau protein is inefficient to stabilize the organization of cytoskeleton during the process of axonal outgrowth, because the protein loses its microtubule-binding ability (17, 21). The detailed molecular mechanisms by which tau becomes a non-functional protein remain elusive (22). But it is suggested that abnormal metabolism and hyperphosphorylation of tau trigger its detaching from microtubules, aggregating in the cytosol and exhibiting the pathological features (17). Tau also showed a higher affinity to bind DNA compared to microtubules. The functions of tau in nuclei are not completely found, but it seems that tau may have a protective role against oxidative stress conditions (23, 24). Tau has long stretches of positively and negatively charged segments, which do not contribute to intermolecular hydrophobic interactions (17). It contains a hexapeptide motif in the four-repeat domain, which is essential for paired helical filaments (PHF) formation (25, 26) with a high tendency for β structures (27). The β structure in monomeric tau can self-assemble by their own into filaments and co-assemble with heparin as an artificial inducer in vitro (17). The recombinant tau protein may be a good mimic for the native human protein and extensive studies have been carried out to determine the molecular structure and biochemical properties of the protein (28). Therefore, the high yield of pure tau provides a critical advantage for in vitro studies. Nonetheless, due to the high tendency to aggregate and truncating, the expression and purification processes of tau protein are somewhat complicated and time-consuming (29). Non-histidine-tagged (His-tag) recombinant tau is typically purified using heat denaturation and cation-exchange chromatography (30–32), and further purification is performed by gel filtration (33). Nonetheless, the purification process of the protein may not be entirely optimized to achieve >95% of purity or yield a native conformation of the protein without structural modifications. In addition, contaminating nucleic acids is another important factor that has not been fully considered about these methodologies. The main aim of the present work was to investigate the effects of tau protein on aggregation of Aβ25–35 over a long time course (40 days) of co-incubation in vitro, to find a crosstalk between the two molecules. For this purpose, it was necessary to provide an active form of tau protein, which was free of any DNA contaminations as well as protein truncations. Accordingly, we described here three methods necessary for studying the characteristics of tau polymerization to PHFs and the Aβ25–35 peptide fibrillation. These include; (i) a verified purification method for the recombinant tau protein; (ii) the quantification and qualification of polymerization of tau to PHFs by a fluorescence-based assay and microscopic imaging; (iii) co-incubation of tau protein with the Aβ peptide and analysis of the self-assembly of both molecules by microscopic imaging and cytotoxicity assays. Materials and Methods Chemicals Tris, Coomassie Brilliant Blue G250 and sodium dodecyl sulphate (SDS) were obtained from Acros organics (New Jersey, United States). Glycine, imidazole, sodium chloride, dithiotheritol (DTT), agarose, phenylmethylsulfonyl fluoride (PMSF) and isopropyl-β-D-thiogalactopyranoside (IPTG) were obtained from Bio Basic Inc. (Markham, Ontario, Canada). Thioflavin T (Th-T), GTP, heparin, 3-(4,5-dimethylthiazolyl-2)-2,5-diphenyltetrazolium bromide (MTT) and trypsin were purchased from Sigma-Aldrich (Saint Louis, MO, USA). Ampicillin was obtained from Jaber Ebne Hayyan Pharmaceutical Co. (Tehran, Iran). Tryptone and yeast extract were provided by Micromedia Trading House Ltd. (Pest, Hungary). Phosphate-buffered saline (PBS) and Dulbecco's modified Eagle's medium (DMEM) were provided by Gibco (Gibco-BRL, Gaithersburg, MD, USA). Urea, EDTA, EGTA, LiCl, Tween 20, Triton X-100 and all other chemicals were obtained from Merck (Darmstadt, Germany). Nickel-nitrilotriacetic acid agarose (Ni-NTA agarose) and SP Sepharose resins were purchased from Bio Basic Inc. (Markham, Ontario, Canada) and Amersham-Pharmacia (Piscataway, NJ, USA), respectively. DNA ladder (DM4100) was purchased from Fermentas International, Inc. (Burlington, Canada) and protein standard marker was purchased from Thermo Fisher Scientific (Waltham, MA, USA). HRP-conjugated anti-His-tag monoclonal antibody was obtained from Roche Diagnostics (Mannheim, Germany). Plasmids, cell lines and protein expression condition Aβ25–35 lyophilized peptide was a generous gift from Professor Saeed Balalaie, K. N. Toosi University of Technology, Iran. PC12 cell line was obtained from Pasture Institute of IRAN, Tehran, Iran. The cells were maintained in DMEM with HEPES 10 mM, glucose 1.0 g/l, NaHCO3 3.7 g/l, penicillin 100 units/ml, streptomycin 100 μg/ml, 10% foetal bovine serum, incubated at 37°C with 5% CO2. The plasmids pET21a encoding His-tag tau 412 (1N4R isoform) gene (GenBank accession no: P10636) was provided (34) and transformed into Escherichia coli BL21 (DE3) cells. The transformed bacterial cells were incubated overnight in Luria Broth medium with 100 µg/ml ampicillin at 37°C and protein expression was induced by addition of 1 mM IPTG at OD600 of 0.6 and incubation was continued at 37°C for 3 h. All bacterial cultures were then harvested by centrifugation (5,000 × g for 20 min) and stored at −20°C. Purification was performed under native and denaturing conditions as described below. Purification of the recombinant tau protein Method 1. Purification by affinity chromatography under native condition Purification with method 1 was performed based on a previous method with minor modifications (35). The pellet of bacterial cell culture was resuspended in a lysis buffer (30 mM Tris-base, 100 mM NaH2PO4, 100 mM NaCl and 1 mM EDTA, 2 mM PMSF and 3 mM DTT, pH 8.0), disrupted by sonication and clarified by centrifugation (12,000 × g, 20 min, 4°C). The supernatants were loaded onto a Ni-NTA agarose column pre-equilibrated with a washing buffer (30 mM Tris-base, 100 mM NaH2PO4, 100 mM NaCl and 15 mM imidazole, pH 8.0) and re-washed with the buffer. Finally, the protein was eluted with a buffer containing 100 mM NaH2PO4, 100 mM NaCl, 30 mM Tris-base and 100 mM imidazole, pH 8.0. Aliquots of the eluted protein were taken for electrophoresis. Method 2. Purification by cation-exchange and affinity chromatography under native condition Purification with method 2 using cation-exchange chromatography was done based on the method provided by Khalili et al., with some modifications (34). The bacterial cells were disrupted as described in method 1. The supernatant containing the soluble fraction of the crude extract was decanted and filtered through a 0.2 μm filter. The filtered supernatants were loaded onto an SP Sepharose column pre-equilibrated with a buffer containing 50 mM NaH2PO4 and 50 mM NaCl, pH 7.5. The protein fractions were then eluted by a linear gradient of salt (0–100 mM NaCl) in the same buffer. Tau containing fractions were pooled and loaded onto a Ni-NTA agarose column pre-equilibrated with the washing buffer (100 mM NaH2PO4, 30 mM Tris-base, 100 mM NaCl, 15 mM imidazole, 2 mM PMSF, 1 mM EDTA and 1 mM DTT, pH 8.0). The unbound proteins were re-washed out by the same buffer and tau proteins were eluted by the elution buffer (100 mM NaH2PO4, 30 mM Tris-base, 100 mM NaCl, 100 mM imidazole, 2 mM PMSF, 1 mM EDTA and 1 mM DTT, pH 8.0). At each step, the aliquots were taken for electrophoresis and the final purified fraction was used for western blot analysis. Method 3. Purification under denaturing condition by affinity chromatography Purification of tau protein with method 3 was proposed, for the first time, based on some principles provided for expression and purification of IDPs (36). Accordingly, the pellet of bacterial cell culture was disrupted in a lysis buffer containing 10 mM Tris-base, 100 mM NaH2PO4, 300 mM NaCl, 0.3% Triton X-100, 8 M urea and 5 mM β-mercaptoethanol, pH 8.0, followed by sonication and clarification by centrifugation (12,000 × g, 20 min, 4°C). Ni-NTA agarose column was equilibrated with buffer A (10 mM Tris-base, 100 mM NaH2PO4 and 8 M urea, pH 8.0) before addition of the supernatant. The supernatant was loaded onto the pre-equilibrated column, the column was re-washed with buffer A, followed by buffer B (10 mM Tris-base, 100 mM NaH2PO4, 8 M urea, pH 6.3) and the protein fractions were eluted with buffer C (10 mM Tris-base, 100 mM NaH2PO4, 8 M urea, pH 5.9) and buffer D (10 mM Tris-base, 100 mM NaH2PO4, 8 M urea, pH 4.5). The purity of the eluted protein was checked using electrophoresis analysis. Finally, excess salts, imidazole and urea in the collected protein fractions were removed by three times dialyzing against 100 mM sodium phosphate buffer (pH 7.6) containing 80 mM Na2HPO4, 20 mM NaH2PO4, 10 mM Tris-base, by gentle stirring for 12 h at 4°C. Protein concentrations were examined using Lowry assay (37). The dialyzed proteins were stored at −20°C for further use. Electrophoresis and western blots The purity was determined using 12.5% reducing sodium dodecyl sulphate–polyacrylamide gel electrophoresis (SDS–PAGE) by the method of Laemmli (38) followed by Coomassie Brilliant Blue staining. For detection of nucleic acid contamination of the purified samples, electrophoresis was performed on 1% agarose gel followed by staining with ethidium bromide. To identify tau positive bands, western blot was carried out using an anti-His-tag antibody. For western blotting, the final fraction of the purified tau protein with method 2 was used. In vitro formation of PHF PHF formation of tau isoforms 1N4R was carried out in 10 mM PBS-buffer pH 7.4 with 8 μM tau protein and 2 μM heparin. The samples were incubated at 37°C for 16 days with shaking at 250 rpm to promote filament formation. DTT was added every day at a final concentration of 2 mM to prevent intermolecular disulphide bond formation. Assembly was followed either qualitatively by atomic force microscopy or quantitatively by fluorescence assay using Th-T. Fluorescence emission intensity was measured at 485 nm after excitation at 440 nm by the Cytation™ 3 Cell Imaging Multi‐Mode Reader (BioTek Instruments, Winooski, VT, USA). 25 µM Th-T was used for each experiment. Microtubule assembly assay Tubulin polymerization was monitored turbidimetrically at 350 nm with a UV-S2100 spectrophotometer (Scinco, Seoul, South Korea). The final volume of the sample was 250 μl. Experiments were run by incubating 10 μM tubulin (at final concentration) in 250 μl of G-PEM buffer (2 mM MgCl2, 1 mM EGTA, 10 mM PIPES pH 6.9, 1 mM GTP) containing tau at a final concentration of 2.5 μM. Tubulin polymerization is mainly regulated by temperature. At 37°C, tubulin will polymerize into microtubules while at lower temperatures, microtubules will depolymerize to the tubulin subunits. It is critical therefore to pay particular attention to temperature throughout the assay (39). Accordingly, the mixture was prepared at 4°C and the reaction was initiated after addition of tubulin and placing the cuvette in the spectrophotometer at 37°C for 15 min, followed by a decrease in temperature to 15°C for 5 min and then temperature increasing to 37°C for 5 min. In this experiment, the protein purified with method 3 was used. Circular dichroism (CD) spectroscopy CD measurement at far-UV region (190–260 nm) was performed for evaluating the secondary structural content of tau protein. The measurement was done by a JASCO J-715 CD spectropolarimeter (Tokyo, Japan) in a 1 mm path length cell with the protein in 10 mM phosphate buffer, pH 7.4 at 4°C. Tau concentration for CD measurement was 200 µg/ml. In this experiment, the protein purified with method 3 was used. Preparation of Aβ25–35 peptide The lyophilized peptide was dissolved in H2O at a concentration of 4.5 mM and stored at −80°C. This condition has been shown to lead to the predominance of the soluble monomeric form of the Aβ peptide. In any case, in order to verify the non‐aggregated form of the peptide, the size analysis was carried out using dynamic light scattering (DLS) by Zetasizer Nano ZS instrument (Malvern Instruments Ltd., Malvern, Worcestershire, UK). Co-incubation of tau protein with Aβ25–35 Aggregation of Aβ25–35 was investigated in the presence of physiologic (4 µM) and supra-physiologic (20 µM) concentrations of tau under reducing and non-reducing conditions. Each sample contains 450 µM Aβ25–35 (as a supra-physiologic concentration of the peptide), 4 and 20 µM tau protein in 1× PBS, pH 7.4. Samples were incubated at 37°C for 40 days without shaking. Under reducing condition, DTT was added at a final concentration of 1 mM every day in order to avoid intramolecular disulphide cross-linking of tau protein. Under non-reducing condition, DTT was added for the first half of the incubation period, followed by incubation without DTT. Atomic force microscopy The samples were prepared by spotting 5 µl of Aβ25–35 and tau solutions on freshly cleaved muscovite mica disks. The contact time was 1 min; mica disks were then rinsed with 0.22 µm filtered deionized water and gently dried under a nitrogen flow. Morphological analysis was carried out with an NT–MDT atomic force microscope (Di-Veeco, Santa Barbara, CA) in semicontact mode. Analysis of AFM images was performed using Gwyddion software (version 2.31), for height, width and dimension of different species (oligomers, amorphous aggregates and/or fibrils). Additionally, the abundance of different oligomeric species and amorphous aggregates in the samples was estimated by plotting the dimension against the abundance from the AFM images. Cell viability assay The PC12 cells were cultured, trypsinized, washed with PBS, counted and seeded in a 96-well plate. After cells attachment (3–4 h), the medium was replaced with fresh medium containing tau protein at different concentrations (0.75, 1.5 µM in reducing condition and 4, 8 µM in non-reducing condition). After 1 h, Aβ25–35 (20, 40 µM in supra-physiologic doses) was added to the medium and the cells were incubated for 72 h at 37°C. To determine viability, the cells were treated with 1 mg/ml MTT for 3 h at 37°C and then the medium was carefully removed. The resulting formazan crystals were dissolved in 100 µl of DMSO, and absorbance at 570 nm was determined using a microplate reader (µQuant, BioTek, USA). Statistical analysis All of the data were expressed as means ± SE of three independent experiments. Data were analysed for statistical differences by one‐way analysis of variance; a P-value of <0.05 was considered significant. Results Purification of recombinant tau protein According to SDS–PAGE analysis for method 1, the protein band of expected size was detected for the recombinant protein. As observed in Fig. 1A, the fractions of purified protein by Ni-NTA agarose chromatography at higher concentrations contained the other cellular protein truncations (lane 2 in Fig. 1A). In addition, agarose gel electrophoresis analysis clearly detected nucleic acid contaminations of the purified proteins (lane 2 in Fig. 1B). As the proteins purified by method 1 showed truncations and contaminations, the purification was examined with method 2 by using both IMAC and cation-exchange chromatography. The most important feature of the present method was the efficiency of the SP Sepharose resin in isolation of nucleic acid fragments. Therefore, the proteins were firstly passed through an SP Sepharose column (Fig. 1C and D; lane 1) and the extracted fractions containing tau were loaded on a Ni-NTA agarose column (Fig. 1D; lane 2). The levels of tau expression, purification and truncation were checked by SDS–PAGE 12.5% (Fig. 1D) and western blotting (Fig. 1E) analysis. Two bands representing the full length purified protein and the truncated form of tau were detected for the final purified proteins (Fig. 1D and E). According to the agarose gel electrophoresis result, contaminating nucleic acids were not detectable in the samples (Fig. 1F). Moreover, the purified proteins from methods 1 and 2 were incubated with 1 M LiCl at high temperatures to obviate nucleic acid fragments and truncation products, respectively. According to the results, these conditions did not result in any decrease in levels of the remained truncated proteins and nucleic acids (Supplementary Fig. 1). Finally, purification of recombinant tau protein with Ni-NTA affinity chromatography under denaturing condition (method 3) resulted in purifying tau protein via a single chromatography column without any contaminating nucleic acid and truncating product (Fig. 1G and H). This strategy provided several advantages including; (i) in 8 M urea, the electrostatic interactions between tau and ionic macromolecules such as nucleic acids are minimized and this obviates any contaminations and (ii) the proteolysis of tau decreases, because of probable denaturing the protease structures. Fig. 1. Open in new tabDownload slide Analysis of tau purification using SDS–PAGE followed by western blotting and agarose gel. (A, B) Purified protein fractions using method 1; (A) SDS–PAGE 12.5% analysis of the purified proteins at lower (lane 1) and higher concentrations (lane 2). Truncation products were noticeable; (B) agarose 1.5% gel of the purified fractions (lanes 1 and 2 in A) with ethidium bromide staining. Nucleic acid bands were detectable for the higher concentrations of the protein (lane 2). (C–F) Purified protein fractions using method 2; (C) the chromatogram of the gradient elution using SP Sepharose column with 0–100 mM NaCl; (D) tau recovered from SP Sepharose column at first step (lane 1) and second purification step using Ni-NTA agarose column (lane 2). The truncated protein fragments were detectable (lane 1); (E) western blots of the final purified tau (lane 2 in D) showing truncation products; (F) agarose gel electrophoresis did not show any nuclei acid fragments. (G, H) Purified tau using method 3; (G) SDS–PAGE analysis confirmed the purified protein free of truncated products; (H) agarose 1.5% gel with ethidium bromide staining and free of nucleic acids. M: protein size marker and M′: DNA ladder. Fig. 1. Open in new tabDownload slide Analysis of tau purification using SDS–PAGE followed by western blotting and agarose gel. (A, B) Purified protein fractions using method 1; (A) SDS–PAGE 12.5% analysis of the purified proteins at lower (lane 1) and higher concentrations (lane 2). Truncation products were noticeable; (B) agarose 1.5% gel of the purified fractions (lanes 1 and 2 in A) with ethidium bromide staining. Nucleic acid bands were detectable for the higher concentrations of the protein (lane 2). (C–F) Purified protein fractions using method 2; (C) the chromatogram of the gradient elution using SP Sepharose column with 0–100 mM NaCl; (D) tau recovered from SP Sepharose column at first step (lane 1) and second purification step using Ni-NTA agarose column (lane 2). The truncated protein fragments were detectable (lane 1); (E) western blots of the final purified tau (lane 2 in D) showing truncation products; (F) agarose gel electrophoresis did not show any nuclei acid fragments. (G, H) Purified tau using method 3; (G) SDS–PAGE analysis confirmed the purified protein free of truncated products; (H) agarose 1.5% gel with ethidium bromide staining and free of nucleic acids. M: protein size marker and M′: DNA ladder. The absorbance at 280 and 260 nm of the purified samples was determined and the ratio (A280/A260) has been compared (Table I). According to the results, the purification using method 1 showed the lowest efficiency with the maximum nucleic acid contamination. The protein purified by method 2 showed a higher A280/A260 ratio (1.7) compared to method 3 (1.3). It is possible that the induced conformational changes of the cleaved/truncated forms of tau proteins (purified in method 2, Fig. 1D and E) may result in the increased absorbance at 280 nm. Table I. A280/A260 ration of the purified proteins Method . Purification method . A280/A260 . 1 Ni-NTA agarose under native condition 0.6 2 SP Sepharose, Ni-NTA agarose 1.7 3 Ni-NTA agarose under denaturing condition 1.3 Method . Purification method . A280/A260 . 1 Ni-NTA agarose under native condition 0.6 2 SP Sepharose, Ni-NTA agarose 1.7 3 Ni-NTA agarose under denaturing condition 1.3 Open in new tab Table I. A280/A260 ration of the purified proteins Method . Purification method . A280/A260 . 1 Ni-NTA agarose under native condition 0.6 2 SP Sepharose, Ni-NTA agarose 1.7 3 Ni-NTA agarose under denaturing condition 1.3 Method . Purification method . A280/A260 . 1 Ni-NTA agarose under native condition 0.6 2 SP Sepharose, Ni-NTA agarose 1.7 3 Ni-NTA agarose under denaturing condition 1.3 Open in new tab Th-T fluorescence assay The measurement of Th-T fluorescence changes upon PHF formation is a good indicator of filament formation from tau protein. Under this experiment, the protein is unfolded and follows the fibril assembly pathways to form mature PHFs (28). In vitro amyloid fibril formation by tau is extremely slow in the absence of polyanions (40); therefore, heparin is usually used to induce aggregation of tau protein (2). The interaction with polyanions leads to a conformational switch from mostly random coil to a β-sheet structure in some parts of the protein fragment and this conformation is a critical step in formation of filaments (41). Because this structure can bind rapidly to Th-T and led to increasing the fluorescence intensity, Th-T-based assays are among the most popular tools for following tau fibrillation in real time (42). Note that tau purified by method 1 could not increase Th-T fluorescence intensity over the time course of incubation (Fig. 2A), but for the samples obtained by method 2 and method 3, the fluorescence intensity at 480 nm increases about 2.5- (Fig. 2B) and 10-fold (Fig. 2C), respectively, in the presence of induced-tau filaments compared to that with the free dye. Although both samples showed an enhanced fluorescence intensity, the increased intensity was much more considerable for the purified tau by method 3 and followed a sigmoidal kinetic, which is considered as a characteristic of most amyloidogenic proteins. This behaviour suggests a common mechanism for fibrillation models including the assembly of oligomers, the formation of nuclei as well as the growth and the breakage of fibrils (43). Fig. 2. Open in new tabDownload slide Aggregation of purified tau to PHFs. The kinetics of aggregation of tau were measured by Th-T assay. The incubation was carried out with 8 μM tau protein, 2 μM heparin in PBS, pH 7.4 and 2 mM DTT at 37°C for 16 days with shaking. The emission intensity with the excitation wavelength at 440 nm was measured. Fluorescence intensities are relative to the emission of 25 µM Th-T at 480 nm for the purified tau by method 1 (A), method 2 (B) and method 3 (C). Fig. 2. Open in new tabDownload slide Aggregation of purified tau to PHFs. The kinetics of aggregation of tau were measured by Th-T assay. The incubation was carried out with 8 μM tau protein, 2 μM heparin in PBS, pH 7.4 and 2 mM DTT at 37°C for 16 days with shaking. The emission intensity with the excitation wavelength at 440 nm was measured. Fluorescence intensities are relative to the emission of 25 µM Th-T at 480 nm for the purified tau by method 1 (A), method 2 (B) and method 3 (C). Structural and functional characterization of 1N4R human tau purified by method 3 Effect of purified tau protein on tubulin polymerization was investigated. The results from tubulin polymerization showed that the polymerization was induced after 20 min incubation at 37°C confirming the active form of tau protein purified with method 3 (Fig. 3A). The secondary structure of the purified recombinant tau was further characterized using CD spectroscopy (Fig. 3B). The spectra of tau showed minimum peak around 200 nm, characteristic of predominantly random coil structures. Furthermore, the formation of fibrils was monitored using atomic force microscopy (Fig. 3C). AFM imaging analysis showed that after 16 days of incubation, purified tau by method 3 was aggregated into fibrils. The other forms of aggregates and small oligomers were also detectable (Fig. 3C). Fig. 3. Open in new tabDownload slide Characterization of the purified tau protein by method 3. (A) Tubulin polymerization assay. The experiment was performed by incubating 10 μM tubulin and 2.5 μM tau in G-PEM buffer, pH 6.8. The turbidimetric analysis of microtubules assembly was performed at 350 nm. (B) CD spectra within the far-UV region. The spectra for monomers showed negative peaks at about 200 nm. (C) Atomic force microscopy of in vitro PHFs showing the morphological species. Filament formation was induced by 8 μM tau in the presence of 2 μM heparin in PBS-buffer, pH 7.4 at 37°C for 16 days with shaking. Tau globular oligomers with protofibrils and fibrils are detectable. The arrowheads indicate the paths of straight fibrils (green arrowheads) and smooth fibrils (red arrowheads). Protofibril bundles (blue arrowheads) and globular structures (orange arrowheads) are also detectable. Dimension abundance of globular oligomers (inset) of tau was analysed using Gwyddion software version 2.31. Fig. 3. Open in new tabDownload slide Characterization of the purified tau protein by method 3. (A) Tubulin polymerization assay. The experiment was performed by incubating 10 μM tubulin and 2.5 μM tau in G-PEM buffer, pH 6.8. The turbidimetric analysis of microtubules assembly was performed at 350 nm. (B) CD spectra within the far-UV region. The spectra for monomers showed negative peaks at about 200 nm. (C) Atomic force microscopy of in vitro PHFs showing the morphological species. Filament formation was induced by 8 μM tau in the presence of 2 μM heparin in PBS-buffer, pH 7.4 at 37°C for 16 days with shaking. Tau globular oligomers with protofibrils and fibrils are detectable. The arrowheads indicate the paths of straight fibrils (green arrowheads) and smooth fibrils (red arrowheads). Protofibril bundles (blue arrowheads) and globular structures (orange arrowheads) are also detectable. Dimension abundance of globular oligomers (inset) of tau was analysed using Gwyddion software version 2.31. Effect of tau on Aβ25–35 aggregation Aβ25–35 peptide employed in our experimental procedures was in non‐aggregated form i.e. predominately monomeric. The size and morphology of the aggregated structures were visualized by microscopy technique. According to the statistical results, under reducing condition, Aβ25–35 aggregates were mainly in the form of granular structures with the average dimensions and height of 21 and 1.78 nm, respectively (Fig. 4A–C). Tau protein in physiological conditions mainly was granular oligomers with the average dimensions and height of 40.4 and 6.30 nm, respectively (Fig. 4D–F). More interestingly, for the co-incubated samples, the morphology and size of the fragments were different with the individual fragments and the new formed annular protofibrils with the average diameter and height of 48 and 1.6 nm, respectively, were detected (Fig. 4G–I). Fig. 4. Open in new tabDownload slide AFM images showing different morphological species of tau and Aβ25–35 under reducing condition. The images demonstrate the presence of various structures of Aβ25–35 (450 µM) and tau protein (4 µM) in 1× PBS, pH 7.4 at 37°C for 40 days without shaking. DTT was added at a final concentration of 1 mM every day. (A, B) Aβ25–35 granular oligomers (blue arrowheads) and large aggregates (red arrowheads), (C) dimension abundance of granular oligomers of the Aβ peptide; (D, E) granular tau oligomers (blue arrowheads), (F) dimension abundance of granular oligomers of tau; (G, H) Aβ25–35 + tau annular (green arrowheads), spherical (blue arrowheads) and granular structures (orange arrowheads), (I) diameter abundance of annular structures. Fig. 4. Open in new tabDownload slide AFM images showing different morphological species of tau and Aβ25–35 under reducing condition. The images demonstrate the presence of various structures of Aβ25–35 (450 µM) and tau protein (4 µM) in 1× PBS, pH 7.4 at 37°C for 40 days without shaking. DTT was added at a final concentration of 1 mM every day. (A, B) Aβ25–35 granular oligomers (blue arrowheads) and large aggregates (red arrowheads), (C) dimension abundance of granular oligomers of the Aβ peptide; (D, E) granular tau oligomers (blue arrowheads), (F) dimension abundance of granular oligomers of tau; (G, H) Aβ25–35 + tau annular (green arrowheads), spherical (blue arrowheads) and granular structures (orange arrowheads), (I) diameter abundance of annular structures. Under non-reducing condition, the results were also remarkable (Fig. 5). In control sample containing Aβ25–35 fragments, all structures were in granular shapes and the amorphous structures with the average dimensions and height of 205 and 25.1 nm, respectively, were detectable (Fig. 5A–C). For tau aggregates in supra-physiologic concentration (20 µM), spherical and globular oligomers with the average dimensions and height of 49.55 and 6.76 nm, respectively, were obviously detectable (Fig. 5D–F) and these aggregates were structurally different with those observed for tau aggregates in the physiologic concentration (4 µM) under reducing condition (Fig. 4D–F). However, co-incubation of Aβ25–35 with tau for 40 days under similar condition resulted in various structural forms of aggregates in which the particles were as spherical and globular structures with the average dimensions and height of 43.66 and 3.81 nm, respectively (Fig. 5G–I). Fig. 5. Open in new tabDownload slide AFM images showing different morphological species of tau and Aβ25–35 under non-reducing condition. The images demonstrate the presence of various structures of Aβ25–35 (450 µM) and tau protein (20 µM) in 1× PBS, pH 7.4 at 37°C for 40 days without shaking. (A, B) Aβ25–35 amorphous structures (blue arrowheads), (C) dimension abundance of amorphous structures of the Aβ peptide; (D, E) spherical (green arrowheads) and granular (red arrowheads) tau oligomers, (F) dimension abundance of all tau oligomers; (G, H) Aβ25–35 + tau spherical (green arrowheads) and granular structures (red arrowheads), (I) dimension abundance of all structures. Fig. 5. Open in new tabDownload slide AFM images showing different morphological species of tau and Aβ25–35 under non-reducing condition. The images demonstrate the presence of various structures of Aβ25–35 (450 µM) and tau protein (20 µM) in 1× PBS, pH 7.4 at 37°C for 40 days without shaking. (A, B) Aβ25–35 amorphous structures (blue arrowheads), (C) dimension abundance of amorphous structures of the Aβ peptide; (D, E) spherical (green arrowheads) and granular (red arrowheads) tau oligomers, (F) dimension abundance of all tau oligomers; (G, H) Aβ25–35 + tau spherical (green arrowheads) and granular structures (red arrowheads), (I) dimension abundance of all structures. Cytotoxicity of the aggregated fragments using MTT assay Because Aβ25–35 has been shown to induce neuronal apoptosis in the pathogenesis of AD, we tested the cytotoxicity of Aβ25–35 on PC12 cells. Cultures were treated with 20 and 40 µM Aβ25–35, and cell viability was determined 72 h later using MTT assay. To evaluate the effects of tau on Aβ25–35-induced cytotoxicity, we pre-treated PC12 cells with 0.75, 1.5 µM tau (for reducing condition experiments), and 4, 8 µM tau (for non-reducing condition experiments) 1 h prior to the treatment with the Aβ peptide. Results showed that under reducing condition, no toxicity effect was observed among the samples (Fig. 6A). However, under non-reducing condition, PC12 cells treated with 8 µM tau or 40 µM Aβ25–35 alone showed a significant decreased viability (Fig. 6B). More interestingly, tau protein at 8 µM protected cells against the toxic effects of 40 µM Aβ25–35. Fig. 6. Open in new tabDownload slide Tau protects against Aβ25–35-induced cytotoxicity in PC12 cells under reducing (A) and non-reducing (B) condition. PC12 cells were treated with 20, 40 µM Aβ25–35 and 0.75, 1.5 µM (A) and 4 and 8 µM (B) tau for 72 h at 37°C. Cell viability was assessed by MTT assay. All data shown represent the mean ± SD (n = 3). *P < 0.05 versus control group. #P < 0.05 versus 40 µM Aβ25–35-treated cell group. Fig. 6. Open in new tabDownload slide Tau protects against Aβ25–35-induced cytotoxicity in PC12 cells under reducing (A) and non-reducing (B) condition. PC12 cells were treated with 20, 40 µM Aβ25–35 and 0.75, 1.5 µM (A) and 4 and 8 µM (B) tau for 72 h at 37°C. Cell viability was assessed by MTT assay. All data shown represent the mean ± SD (n = 3). *P < 0.05 versus control group. #P < 0.05 versus 40 µM Aβ25–35-treated cell group. Discussion Intracellular deposition of microtubule-associated tau protein as filamentous aggregates is a hallmark pathology of neurodegenerative disorders including AD and other tauopathies (44). Tau is a phosphorylated protein, and the residues phosphorylated on tau are mainly found in its proline-rich flanking domain for tight binding to microtubules (7). Soluble tau contains a little secondary structure, mostly random coils, but this is enough for fibrillization of tau into PHFs, despite their long-range periodicity (2, 45). Tau protein and Aβ peptide can assemble in vitro, yielding filamentous polymers, either alone, or in the presence of inducers like heparin (46). NFTs can be found inside the cell, while SPs are found in the extracellular space (7). There are several studies in which Aβ-induced neurodegeneration with the concomitant tau aggregation as the result of direct interaction of the Aβ peptide with tau protein have been investigated (5, 26, 47). In this study, we have provided some evidences in supporting a key role for tau in Aβ25–35 fibrillation. We investigated the effects of tau protein on Aβ25–35 aggregation over a long time of incubation (40 days), for the first time, to find a crosstalk between the two molecules in an AD-like way. We further tried to consider the physiological and supra-physiological conditions under pH, temperature and ionic potent without any inducer to obtain various types of oligomer structures in vitro. For this purpose, firstly, it was necessary to provide a form of tau protein without any nucleic acid contaminations or protein truncations. Accordingly, we presented a new efficient purification method to provide an active recombinant tau protein suitable for further downstream applications. The His-tag tau construct was used for expression in bacterial cells (34). It has been reported that the assembly characteristics of the recombinant tau are not influenced by the presence of the His-tag (28, 48). The expressed protein was purified using chromatography methods and characterized by SDS–PAGE and dye-based fluorescence spectroscopy. Our findings demonstrated that method 3 led to the high-yield production of an authentic form of tau protein, with the known biochemical and biophysical properties. Notably, this was the first report for purification of tau protein using Ni-NTA affinity chromatography under a denaturing condition by urea. Purification processes In method 1, we used Ni-NTA agarose column to purify His-tag tau protein. The results obtained from SDS–PAGE analysis showed that although this method resulted in a high yield of the protein (Fig. 1A), nucleic acid contaminations significantly occurred during the purification process (Fig. 1B), and the contaminations were not removed after LiCl treatments (Supplementary Fig. 1). In addition, the in vitro assembly assay was performed in order to evaluate the nature of purified tau in the formation of fibrils. According to the results, at an excitation wavelength of 440 nm, the emission intensity of Th-T fluorescence did not increase (Fig. 2A) showing that the purified protein using method 1 could not form any self-assembled structure. Recent studies demonstrated that Th-T can also bind DNA structures, accompanied by a sharp increase in fluorescence emission (49, 50). These observations strongly suggested that Th-T have a high tendency for binding the inner regions of DNA. In fact, molecules of the cyanine family are strong intercalators and Th-T can itself interact with DNA (51). Here, purified tau by method 1 showed large DNA contaminations (Fig. 1B) that can justify the increased Th-T fluorescence intensity at the first time. On the other hand, it has been shown that the presence of sodium ion (Na+) can lead to the removal of the electrostatic binding between Th-T and DNA. Indeed, the interaction of Na+ with the phosphate groups of the nucleic acids blocked the electrostatic interaction of Th-T cations with DNA (50). Moreover, the presence of the truncated proteins, which can result in the measurement of non-real concentrations of the active form of tau protein needed for PHF assay, can be considered as the other reason for the decreased intensity during time. Besides, such truncations may also inhibit tau assembly into PHF (52). Using SP Sepharose resin in method 2 led to the elimination of nucleic acid contaminants and then the isolated protein fractions passed through Ni-NTA agarose column for more purification (Fig. 1C and D). Western blotting analysis also revealed that the final purified protein using method 2 showed proteolytic cleavage probably by bacterial proteases (Fig. 1E). The numbers of digested proteins were minimized by passing the protein through Ni-NTA agarose column, but the truncated fragments even remained after heat shock incubation (Supplementary Fig. 1). Totally, method 2 was efficient for purification of tau in high levels, but the production of truncated proteins seems inevitable. Besides, the purification process with two column-chromatographic steps was time-consuming. It probably arises from the high ionic strength of the protein fractions isolated from SP Sepharose resin, which interferes with the binding of the protein to Ni-NTA agarose resin. According to Th-T assay, the fluorescence intensity slightly increased during the time, but a sigmoidal kinetic was not observed for the purified sample (Fig. 2B). As explained earlier, this result can be due to the presence of the truncated products as well as non-real concentrations of the active tau in PHF assay. Tau contains many potential cleavage sites accessible to proteases (52). The relationship between tau cleavage and its tendency to aggregation is unclear, and according to the studies from different in vitro and in vivo models, the fragments cleaved by caspases preceded the aggregation of tau, while cleavage of tau by calpain partially inhibited tau aggregation (7, 16, 53). Accordingly, the presence of such truncations may inhibit tau assembly into PHF. As the final approach, we used method 3 for purification under denaturing condition using urea. According to SDS–PAGE results, a single band was detected for the purified protein and the samples did not contain any nucleic acid contaminations (Fig. 1G and H). Th-T assay revealed an about 40-fold increase in the fluorescence intensity with a sigmoidal trend confirming the self-assembly of tau into filaments (Fig. 2C). More characterization of the purified protein with method 3, as the most efficient approach, was carried out by tubulin polymerization, CD analysis and AFM imaging (Fig. 3). Analysis of tubulin assembly into microtubules showed that pure tau promoted the microtubules polymerization (Fig. 3A). CD analysis also confirmed a dominant random coil for the purified protein (Fig. 3B), that was in good agreement with previously reported structures for native tau (45, 52, 54). In addition, the microscopic imaging allowed the characterization of different morphological species and distributions of the filaments. Accordingly, the results showed that straight filaments (SF) with average width of 73.33 nm were formed, while protofibrils and oligomer structures were also observed (Fig. 3C). The average width of protofibrils was 134 nm (n = 5) ranging from 70 to 230 nm, while the average dimension of oligomer structures was 46 nm (n = 20), ranging from 30 to 60 nm (Fig. 3C). Tau assembly pathways Protein aggregation has been shown to involve intrinsically disordered systems, such as in Aβ peptide and tau protein in AD. During the amyloidogenic pathway, the formed oligomeric species are mostly assemblies of monomeric units. These aggregates can form highly disordered structures or well-defined fibrils rich in cross-β structures (55). In addition, as aggregation proceeds, oligomer size and β-sheet content increase. Gradual increase in the oligomer size accompanied by conformational transitions within the amyloid oligomers has been proposed to proceed by monomer dissociation from the less stable oligomers followed by association of this monomer with more stable aggregates (56). On the other hand, at least for some proteins and peptides, the conversion of amyloid oligomers from native-like to β-sheet rich forms has been shown to involve internal reorganization rather than dissociation and reassembly. Overall, conformations of amyloid oligomers are highly diverse with a tendency of increased β-sheet propensity with increasing oligomer size (56). There are differences in the tau fibril morphology found within different human tauopathies (57), and the multiple assembly pathways can further occur simultaneously. For example, it has been shown that spherical nucleation units, formed in the early stages of the tau fibrillation reaction, appear to align to form the initial fibrils (58). Moreover, there are others reports indicating that tau fibrillation reactions can be greatly accelerated by the addition of fully mature fibrils (59), and the introduction of these fibrillar nucleation scaffolds may drive further growth by elongation from existing fibril ends in a process that disfavours a colloidal pathway. Consistent with the findings mentioned above, we suggested that various monomer and oligomer structures appear to further assemble to higher-order oligomers to eventually become larger fibrils as observed in AFM results (Fig. 3C). There are various types of tau intermediate structures, which probably differ with each other according to their structural properties and functional roles (30, 60–64). The tau oligomers are also structurally modified. They are rich in β-sheet structures and the formation of fibrils promotes when the size of oligomers reach to 20 nm (45). The granular protofilaments of tau can be identified by an in vitro tau aggregation system. These granular tau oligomers appear to be composed of nearly 40 tau molecules and are able to become filaments in a dose-dependent manner (45). Totally, oligomeric assembly is necessary for tau aggregation, which results in the formation of abnormal filaments (65). Interaction of tau and Aβ25–35 Further, we investigated the effect of purified tau protein using method 3 on Aβ25–35 fibrillation in reducing and non-reducing conditions. The cysteines (C291 and C322) are the basis for tau's property to form dimers via a disulphide bond or an intra-dimer disulphide bridge, which locks the molecule in a folded conformation and greatly enhances the rate of aggregation (66). Since this was necessary to use tau protein in its monomeric state, DTT was added during the incubation period under reducing condition. In non-reducing condition, DTT was only added for the first half of the period (16 days). Accordingly, aggregation of Aβ25–35 was investigated by AFM in the presence of physiologic (4 µM) and supra-physiologic (20 µM) concentrations of tau under reducing and non-reducing conditions. Atomic force microscopy can provide accurate estimations of morphology and filament width and morphology. The results showed that under reducing condition, Aβ25–35 aggregates were mainly consist of granular structures, which the average dimension of these structures was 21 nm (n = 40), ranging from 7 to 45 nm (Fig. 4A–C). The average height of the granular structures was 1.78 nm (n = 21). The structures in some areas were prone to forming protofibrils with the width in the range of 58–82 nm. Besides, the granular tau oligomers in physiologic concentration mainly consisted of a uniform dispersion in terms of morphology with the average dimension of 40.4 nm (n = 25), ranging from 29 to 58 nm (Fig. 4D–F). The average height of the granular tau structures was 6.3 nm (n = 18). More interesting, co-incubation of tau with Aβ25–35 resulted in formation of new annular protofibrils (40.74%) with the average diameter of 48 nm (n = 25), ranging from 31 to 68 nm (Fig. 4G–I). The average height of the annular structures was 1.6 nm (n = 10). In addition, spherical (14.81%) and granular (44.44%) structures with the average dimension of 31.2 and 16.93 nm, respectively, were also detectable. On the other hand, the interaction of tau protein in supra-physiologic doses with the peptide under non-reducing condition revealed a considerable difference between the species of oligomers/aggregates formed in the treatments containing tau or Aβ25–35 alone compared to the co-incubated sample (Fig. 5). The treatment containing Aβ25–35 was virtually devoid of structured fibrils, but it had an amorphous-like consistency, with the average dimension of 205 nm (n = 24), ranging from 140 to 320 nm, and the average height of 25.1 nm (n = 15) (Fig. 5A–C). For the sample incubated with the individual tau protein, the spherical (30%) and granular (70%) oligomers were mainly observed with the average dimension of 49.55 nm (n = 20), ranging from 26 to 116 nm, and the average height of 6.76 nm (n = 10) (Fig. 5D–F), which were considerably different from the structures formed under physiological condition (Fig. 4D–F). More interesting, spherical (26.66%) and granular (73.33%) oligomers with the average dimension of 43.6 nm (n = 30), ranging from 24 to 83 nm, and the average height of 3.81 nm (n = 15) were mainly formed in the co-incubated sample containing tau and Aβ25–35 peptide (Fig. 5G–I). In this regard, it is possible that tau completely inhibited the further self-assembly of Aβ25–35 into large aggregates. Classes of aggregated Aβ25–35 and tau Protein aggregates are generally classified as amyloid fibrils, amorphous aggregates and soluble aggregates (56). However, it is often difficult to discriminate between ordered amyloid fibrils and amorphous aggregates even though they are usually different in their morphologies (67). Amyloid fibrils share a common structural feature forming unique linear self-assemblies with an ordered cross-β structure while insoluble amorphous aggregates include various types of assemblies without ordered intermolecular interactions and a defined shape (68). Inclusion bodies have been classified as insoluble amorphous aggregates (69). In contrast to amyloid fibrillation, there is little about the mechanisms involved in amorphous aggregation. It is suggested that the aggregation of a polypeptide chain into amorphous structures also occurs coupled with the denaturation of proteins. However, the link between nucleation and amorphous aggregation has not been determined to date (67). Similar amorphous aggregates were detected here for the Aβ peptide under non-reducing condition (Fig. 5A and B). Moreover, soluble aggregates, generically described as amyloid oligomers, are defined as soluble non-monomeric structures, which are present as intermediates or final products in the protein aggregation process. Soluble aggregation intermediates include protofibrils, annular protofibrils and oligomers (70). Amyloid oligomers lack one or more of the hallmarks of fibrillar aggregates; however, they can form either en route to insoluble fibril formation or independently to fibrillation (56). In comparison, protofibrils appear to share structural similarity to mature fibrils, including a rope-like fibrillar structure detectable by EM and AFM as well as stable hydrogen bonding (70). Such structures were detected in the present work for the tau protein (Fig. 3C). The formation of pore-like structures, known as annular protofibrils and doughnut-shaped structures, with an outer diameter of 8–12 nm and an inner diameter of 2–2.5 nm has been shown for amyloid proteins, including Aβ and α-synuclein for years. Besides, the formation of tau annular protofibrils has been known in vitro and in vivo. It is suggested that tau oligomers can form annular protofibrils by a specific off-pathway mechanism from fibril formation (56, 70). Here, annular protofibrils were detected only in the co-incubated sample containing the Aβ peptide and tau protein in reducing condition (Fig. 4G). Finally, PHF and SF that make up NFTs are comprised of hyperphosphorylated tau (70). SF and smooth filaments were detected in the present work for the tau protein (Fig. 3C). Toxicity assay The Aβ peptide can have a neurotoxic effect, which is attributed to different aspects such as its relative concentrations, as well as the cellular environment, or even the age of the individuals (6). Small oligomeric species, which are considered as the intermediates in fibril formation, are linked to tissue damage and disease progression. The cross β-sheet structure of amyloid assemblies, especially amyloidogenic intermediates, are inherently toxic to cells (71). The Aβ fraction 25–35 has the ability to enter within the cells, causing mitochondrial damage with an evident trigger of apoptotic signals, and toxicity considerably occur by increased oxidative stress (72, 73). Several mechanisms have been proposed to explain the neurotoxic effect of Aβ; (i) production of reactive oxygen species such as hydrogen peroxide and nitric oxide; (ii) intracellular calcium accumulation; (iii) altered membrane fluidity; (iv) energy depletion; (v) alteration of cytoskeleton components and (vi) inflammatory processes (60). Furthermore, evidences suggest that redox state of methionine-35 significantly affects the solubility of the amyloid peptides and mediates toxic effects (59, 61, 74). Besides, oxidation of this residue in Aβ C-terminal domain greatly influences the Aβ peptide aggregation (75) as we also observed for the individual Aβ peptide in non-reducing condition (Fig. 5A). On the other hand, there has been a debate on whether tau is causal to AD or just a by-product of some disease process. About AD, the case is still open, and changes in tau are mostly viewed as a consequence of Aβ pathology (52). Moreover, results from animal models suggested that tau induces Aβ toxicity in neurons, but this remains a matter of debate (76, 77). Therefore, in the final experiment, we examined the toxicity of the Aβ peptide and protein using MTT assay (Fig. 6). The concentrations of tau protein and Aβ peptide for the induced toxicity were in the range of the concentrations suggested by previous reports (72, 78, 79). According to the results, under reducing condition with 0.75 and 1.5 µM tau and 20 and 40 µM Aβ25–35, there was no any toxicity effect on the cells (Fig. 6A). In fact, the peptide and protein did not show any toxicity. In addition, the cytotoxicity was checked in the presence of 4 and 8 µM tau under non-reducing condition (Fig. 6B). Interestingly, although the cell viability decreased either for 8 µM tau or 40 µM Aβ25–35 alone, the co-incubated sample showed lower toxicity effect in comparison with the individual treatments (Fig. 6B). Accordingly, it was suggested that the presence of tau not only modified the assembly of the Aβ peptide (Fig. 5G–I), but also affected its toxicity (Fig. 6B). This result is not entirely consistent with a number of studies; however, similar result in supporting the protective role of tau on Aβ cytotoxicity have been previously reported (80). In addition, it is assumed that the structure and toxicity of tau may be influenced by the Aβ peptide. Tau aggregates cause cytotoxicity, and the precise mechanism of the induced toxicity remains to be determined (81). The toxic behaviour of tau in AD appears to be solely due to its abnormal hyperphosphorylation, because dephosphorylation of pathological tau changed it to a normal protein (82). For several other amyloid-related disorders, soluble oligomers and protofibrils were proposed to be involved in the toxicity (83). Here, the decreased cell viability under non-reducing condition could be attributed with the soluble oligomers of tau (49.55 nm in dimension), as the primary toxic species, which were also identified by AFM imaging (Fig. 5D). Such observations are important for amyloid diseases in which the oligomeric species is observed and believed to be the most toxic. Previous results from proline mutations showed that the hexapeptide motifs of tau are responsible for aggregation of tau into filaments, which lead to cell death (52). Dysfunction of tau protein, which contributes to its dissociation from microtubules and collapse of cytoskeleton, may also lead to toxicity. Tau aggregates appear to be themselves neurotoxic (15). It is possible that with exposure of β-strands, there is an overburden of accumulated misfolded proteins that stimulates the cell’s chaperone-based defense system, which contributes to detoxification of the exposed β-strands (52). Nonetheless, the toxicity of aggregated tau still remains a subject of debate since neurons can live for years with NFT in humans (84). Not all insoluble tau aggregates are toxic and some tau aggregates are non-toxic, and may even be protective against tau toxicity in vivo (61). In support of this, our AFM results showed that tau protein in reducing condition formed smaller granular tau oligomers (40.4 nm in dimension) (Fig. 4C and D). These structures were supposed to be non-toxic by cell viability assays (Fig. 6A). Here, we could provide an optimal method (method 3) to purify tau protein isoform 1N4R with several advantages, including; (i) a high yield of purified protein using a single chromatographic step, (ii) free of any nucleic acids contamination and (iii) truncated products. Besides, we examined the effect of purified tau on aggregation of Aβ25–35 both in terms of morphology and function. The co-incubation of tau with the Aβ peptide under non-reducing condition resulted in the decreased aggregated forms of Aβ25–35. Moreover, the presence of tau in this condition led to increase in cell viability. It seems that oxidation probably plays an important role on assembly and toxicity control of the Aβ peptide with tau. According to our observations, it can be concluded that the presence of tau protein could influence the aggregation behaviour of the Aβ peptide and reduce its toxicity under non-reducing condition. One of the pathological early events that occurs in the brains of AD affected individuals is the oxidative damage. Oxidative stress promotes Aβ generation, and in these conditions, the formation of amyloid plaques could be a compensatory response to remove reactive oxygen species (6). As a result, deposition and accumulation of Aβ (SPs) and hyperphosphorylated tau (NFTs) in AD occur during oxidative stress (62). Very importantly, this study evaluated for the first time, the interaction of tau and Aβ25–35 peptide upon a non-reducing condition over a long period of fibril formation (40 days). Similar conditions usually occur for individuals during the course of the AD. Our observations revealed that the interaction of tau with Aβ25–35 or their assembly led to formation of the new forms of aggregates, which inhibited the toxicity behaviour of tau or the Aβ peptide aggregates. However, there are still many unanswered questions about their interactions and effects. For example, this is not clear whether the annular protofibrils or granular aggregates are formed as mixed structures of tau and Aβ25–35 or every fragment would organize their individual structures. Further, the possibility that Aβ as well as tau protein may play a protective role in preventing each other to acquire the cytotoxic, aggregated forms can also be suggested. Supplementary Data Supplementary Data are available at JB Online. Acknowledgements This work was supported by the research council of Tarbiat Modares University and Ministry of Sciences, Researches and Technology, Iran. Authors’ responsibility F.M. performed the experiments and interpreted data. Z.T. was involved in manuscript writing and revision, experiments and interpretation of data. H.R. and K.K. contributed to the chromatography methods. M.A.N.K. and G.R. were involved in the formation of the experimental concept of the work. R.H.S. supervised the experiments, provided intellectual input, interpreted data and edited the manuscript. Conflict of Interest None declared. References 1 Miyata Y. , Koren J. , Kiray J. , Dickey C.A. , Gestwicki J.E. ( 2011 ) Molecular chaperones and regulation of tau quality control: strategies for drug discovery in tauopathies . Future Med. Chem . 3 , 1523 – 1537 Google Scholar Crossref Search ADS PubMed WorldCat 2 Friedhoff P. , Schneider A. , Mandelkow E.-M. , Mandelkow E. 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Neurosci . 4 , 49 Google Scholar Crossref Search ADS PubMed WorldCat Abbreviations Aβ amyloid-β AD Alzheimer's disease CD circular dichroism DLS dynamic light scattering DMEM Dulbecco's modified Eagle's medium DTT dithiotheritol His-tag histidine-tagged IDP intrinsically disordered protein IPTG isopropyl-β-D-thiogalactopyranoside NFTs neurofibrillary tangles Ni-NTA agarose nickel-nitrilotriacetic acid agarose PBS phosphate-buffered saline PHF paired helical filaments PMSF phenylmethylsulfonyl fluoride SDS–PAGE sodium dodecyl sulphate–polyacrylamide gel electrophoresis SF straight filaments SP senile plaques Th-T thioflavin T. © The Author(s) 2020. Published by Oxford University Press on behalf of the Japanese Biochemical Society. All rights reserved This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)
TI - Interplay of isoform 1N4R tau protein and amyloid-β peptide fragment 25–35 in reducing and non-reducing conditions
JF - The Journal of Biochemistry
DO - 10.1093/jb/mvaa101
DA - 2021-02-06
UR - https://www.deepdyve.com/lp/oxford-university-press/interplay-of-isoform-1n4r-tau-protein-and-amyloid-peptide-fragment-25-0jz9bCVvDF
SP - 119
EP - 134
VL - 169
IS - 1
DP - DeepDyve
ER -