Blood samples were collected from clinically diagnosed sporadic (sALS) and familial (fALS) ALS patients and age-matched disease and healthy controls at the Institute of Neurology, Clinical Center of Serbia. Human sera from 20 ALS patients (3 fALS and 17 sALS; 13 males and 7 females) of age 59.6 ± 2.5 years (mean ± SEM) with the average score of 38.7 ± 1.0 on the ALS functional rating scale according to the El Escorial revised criteria for the diagnostics of ALS (33) and 11 controls (four healthy and seven disease controls; details in Table 1), age 57.4 ± 3.0 years, were collected for routine clinical examination with informed patient’s consent in accordance with The Code of Ethics of the World Medical Association (Declaration of Helsinki) for experiments involving humans. The protocol was approved by the Ethics committee of the Clinical Center of Serbia (No. 1985/5). A part of biological material was used for IgG isolation at the Institute of Virology, Vaccines and Sera-Torlak, Belgrade, Serbia as described previously (20).

In this study, we used microglial BV-2 cell line, frequently used as substitute for primary microglia, derived from raf/myc-immortalized murine neonatal microglia (34). Cells were used in passages 5–15. BV-2 cell line was maintained in growth medium consisting of RPMI with l-glutamine, and sodium pyruvate (Sigma Aldrich, Germany), supplemented with 25 mM HEPES, 10% fetal bovine serum (Gibco, Invitrogen, USA), and penicillin/streptomycin (Gibco, Invitrogen, USA) at 37°C in humidified incubator with 5% CO₂. After the cells reached ~90% confluence, they were washed with PBS, trypsinized (0.25% trypsin and 0.02% EDTA), centrifuged (500 g, 5 min), and then passaged and/or plated for the experiments. For biochemical analysis, cells were plated on 24-well plates (6 × 10⁴ cells/well) in 0.5 ml of growth media, while for the expression analysis, BV-2 microglial cells were plated on 6-well plates (3 × 10⁵ cells/well) in 2 ml of growth media. On the following day, the culture medium was replaced with the culture medium containing ALS IgG (0.1 mg/ml) or control IgG (0.1 mg/ml) and cells were kept for the indicated period of time (4 or 24 h). The IgG concentration tested was selected from our previous studies on neuronal and glial cells in culture (20, 21, 35). Cells kept in medium without IgG were used as a control.

BV-2 cells (6 × 10⁴ cells/well) were plated and treated for 24 h (here and in Sections “Malondialdehyde (MDA) Determination” to “Total Glutathione Determination,” at least in duplicate for each IgG) as described. After the indicated time supernatants were discarded, cells were washed in ice-cold PBS then collected with plastic scraper, lysed by sonication and centrifuged at 15,000 g for 5 min at 4°C for NO determination. NO is a highly unstable molecule and therefore the production was determined indirectly by measuring nitrite concentration spectrophotometrically at 492 nm using Griess method, after reducing nitrates to nitrites by cadmium reduction (36). Results are expressed as mean nitrite concentration (μM) ± SEM for each group.

BV-2 cells (6 × 10⁴ cells/well) were plated and treated for 24 h and collected for measurements as described. MDA content was determined by the spectrophotometric method of Vilicarra et al. (37). First, thiobarbituric acid (TBA) reagent (15% trichloracetic acid and 0.375% TBA water solution, Merck-Darmstadt, Germany) was mixed with cell lysate, heated to 95°C, centrifuged, and then the absorbance was measured at 532 nm. Results are expressed as mean MDA concentration (nmol/ml) ± SEM for each group.

BV-2 cells (6 × 10⁴ cells/well) were plated and treated for 24 h, then collected for measurements as described. Total SOD activity, which combines the activity of two isoforms, MnSOD and cytoplasmic SOD (Cu/ZnSOD), was determined by epinephrine method. SOD activity was determined by spectrophotometric measurement of a decrease in the rate of the spontaneous epinephrine autooxidation at 480 nm. The kinetics of enzyme activity was followed in 50 mM carbonate buffer (pH 10.2) with 1 mM EDTA after the addition of 10 mM epinephrine and 5 mM KCN for MnSOD (38). Cu/ZnSOD activity was determined as a difference between total SOD and MnSOD activity. Results are presented as units per milligram of total protein (U/mg). One unit is described as an amount of protein (enzyme) required for 50% inhibition of autooxidation of epinephrine. Total protein concentration was determined by Lowry method (39).

Catalase activity was assayed by a method based on spectrophotometric determination of colored complex formed between ammonium molibdate and H₂O₂ at 405 nm (40). The results are expressed as units per milligram of total protein (U/mg), whereas the unit represents an amount of H₂O₂ reduced per min (μM H₂O₂/min).

Glutathione peroxidase activity was assayed by indirect method that is based on spectrophotometric measurement of NADPH oxidation by glutathione reductase (GR) at 340 nm (41, 42). Briefly, GPx catalyzes the oxidation of GSH to GSSG which is then recycled back to GSH by GR. In that process GR reduces NADPH coenzyme, as a donor of reducing equivalents, to NADP⁺. The results are expressed as units per milligram of proteins (U/mg), whereas one unit represents an amount of reduced NADPH per min (μM NADPH/min).

Activity of GR was determined using method that is based on fluorimetric measurement of GR-mediated NAPDH oxidation to NADP⁺ (43). Decrease in NADPH fluorescence was measured with excitation/emission of 360/460 nm. We used 100 mM NAD⁺ as a standard in the reaction. Results are expressed as units per milligram of proteins (U/mg). One unit is described as an amount of oxidized NAPDH per min (μM NADPH/min).

BV-2 cells (6 × 10⁴ cells/well) were plated and treated for 24 h, then collected for measurements as described. Intracellular glutathione content was assessed by DTNB-GSSG reductase recycling assay (44). The formation of 5-thio-2-nitrobenzoic acid (TNB), which is proportional to total glutathione content in the sample, was measured spectrophotometrically at 412 nm for 6 min. Total glutathione concentration in samples was determined from standard curve constructed with known GSSG concentrations. Results are expressed as mean total glutathione concentration (nmol/ml) ± SEM for each group.

BV-2 cells (3 × 10⁵ cells/well) were plated, treated for 4 h with ALS IgG or control IgG (in duplicate for each IgG) and then collected. Total mRNA isolation was performed by using TRIzol reagent (Invitrogen, USA). The RNA content was quantified spectrophotometrically at 260 nm and 1 µg of RNA was used for reverse transcription with High Capacity cDNA Reverse Transcription Kit (Applied Biosystems, Foster City, CA, USA). Real-time PCR amplifications were performed with SensiFAST™ SYBR^® Hi-ROX Kit (Bioline, United Kingdom) and specific primers (sequences and annealing temperatures are given in Table 2, Invitrogen, USA), with QuantStudio TM Real-Time PCR System (Applied Systems, USA). The results were analyzed by using 2^−ΔΔCt method. In previous experiments on BV-2 cells it was shown that there is no change in GAPDH, ACTB, and HPRT gene expression following LPS stimulation of these cells (Bozic, unpublished), therefore we used GAPDH and ACTB genes expression as internal controls. Since there were no differences, only results with GAPDH gene expression as an internal control are presented.

BV-2 cells (6 × 10⁴ cells/well) were seeded and treated with ALS IgG or control IgG (in triplicate for each IgG) for 24 h as previously described. After the indicated time supernatants were collected and TNF-α concentration was determined by ELISA by using detection and capture antibody (eBioscience, Germany) according to the manufacturer’s protocol. After applying biotinylated capture antibody, avidine-HRP complex was added, and subsequently chromogenic substrate 3,3′,5,5′-tetramethylbenzedine (eBioscience, Germany). The reaction was stopped by adding 1 M H₃PO₄ and absorbance was measured at 450 nm. TNF-α concentration was determined from standard curve constructed with known murine recombinant TNF-α concentrations. Results are expressed as mean concentration (pg/ml) ± SEM for each group.

BV-2 cells were plated onto PLL-coated 10 mm circular glass coverslips (1.5 × 10⁴ per coverslip). On the following day, cells were treated with human IgG (0.1 mg/ml, ALS or control, in growth media; in duplicate for each IgG) for additional 24 h. After treatment, cells were briefly washed with PBS, fixed in 4% PFA for 15 min, and washed in PBS. Then, the cell membrane was labeled with Wheat Germ Agglutinin, Tetramethylrhodamine Conjugate (WGA, 1:2,000; Molecular Probes, Thermo Fisher Scientific) for 10 min. Next, cells were permeabilized with 0.05% Triton X-100 for 15 min, following washing in PBS. Cells were then labeled with secondary antibody, goat anti-human IgG AlexaFluor 633 (1:200, Molecular Probes, Thermo Fisher Scientific) for 1 h in the dark at 37°C. After several washing steps with PBS, coverslips were incubated for 10 min with a nuclear counterstain (DAPI, 1:4,000), washed again thoroughly, and mounted on microscopic slides using Mowiol 4-88 mounting medium (Sigma Aldrich, Germany). Control staining was done by omitting IgG from the treatment.

Stained cells were visualized on a confocal laser-scanning microscope (LSM 510, Carl Zeiss GmbH, Jena, Germany) equipped with HeNe (543 and 633 nm) lasers. Oil-immersion objective 63×/NA 1.4 was used, and pinhole was set to 2.2 µm for both detection channels.

For live cell imaging BV-2 cells were seeded onto 7 mm (6 × 10³ cells) or 10 mm (1.5 × 10⁴ cells) round glass coverslips (Menzel Glasser, Germany), coated with poly-l-lysine (50 µg/ml, Sigma Aldrich, Germany). For each IgG sample 1–4 coverslips were prepared. The number of analyzed cells [region of interest (ROI)] is indicated in figures. In order to evaluate acute hydrogen peroxide production in response to IgG treatment, BV-2 cells were transfected with HyPer, a genetically encoded sensor for H₂O₂ with expanded dynamic range (45), pC1-HyPer-2 (Addgene plasmid # 42211). Since HyPer shows pH dependence, a set of control experiments were done with BV-2 cells transfected with SypHer, peroxide-insensitive version of HyPer (46), pC1-HyPer-C199S (Addgene plasmid # 42213). Transfection was done with Lipofectamine (LTX, Invitrogen, USA), or Turbofect (Thermo Fisher Scientific, USA), 24 h after plating, and cells were imaged on the following day. In addition, 6-carboxy-2′,7′-dichlorodihydrofluorescein diacetate (carboxy-H2DCFDA, Molecular Probes, Thermo Fisher Scientific, USA) a general oxidative stress indicator was used to detect acute generation of ROS following treatment with IgG. BV-2 cells (24–48 h after plating) were loaded with 50 µM carboxy-H2DCFDA in working solution for 30 min. Cells were briefly rinsed three times and imaged immediately.

Transfected or dye-loaded coverslips were transferred into the recording chamber supplied with 1 ml of working solution, placed on an inverted epifluorescent microscope (AxioObserver A1, Carl Zeiss, Germany) equipped with water, glycerin, and oil immersion objective LD LCI Plan-Apochromat 25x/0.8 (Carl Zeiss, Germany) and combined with VisiFluor Imaging System. The excitation light source was Xenon Short Arc lamp (Ushio, Japan) combined with high speed polychromator system (VisiChrome, Visitron Systems, Germany). The excitation light (480 nm) and the emission light passed through the FITC filter set (Chroma Technology Inc., USA). Time-lapse images were obtained by “evolve”-EM 512 Digital Camera System (Photometrics, USA) via VisiView high performance imaging software (Visitron Systems, Germany). Initially, fluorescence intensities were recorded for 2–5 min to determine the baseline fluorescence (F₀). Thereafter, human IgG (0.1 mg/ml) in working solution (1 ml) were applied directly to the imaged cells by customized delivery system, via glass pipettes (0.8 mm inner diameter, positioned ~350 μm away and ~1 mm above the cells, at the angle of 45°) connected to High Speed Solution Exchange System (ALA Scientific Instruments, USA) with pinch valves and VC3 electronic valve controller. The volume in the recording chamber was kept at ~1 ml by suction from the top of solution. The response of cells to IgG was recorded for additional 5–10 min, followed by several minutes of constant perfusion of working solution (washing step), after which the cells were stimulated by H₂O₂ (100 µM or 4 mM) for 3–10 min, and washed again with working solution. In another set of experiments after recording fluorescence intensities for 2–5 min, 30 mM NH₄Cl in working solution was applied for 4 min and then washed with working solution for several minutes.

The working solution consisted of NaCl (152 mM), KCl (2.5 mM), CaCl₂ (2 mM), MgCl₂ (1 mM), d-glucose (10 mM), and HEPES (10 mM), pH 7.4, adjusted with NaOH. The osmolality of solution was ~300 mOsM, measured by vapor pressure osmometer (Vapro 5520, Wescor, USA).

After the experiment, the analysis began with extracting average fluorescence intensities from ROI that were drawn around each cell, in each time point of the experiment. Additionally, several ROIs were extracted from the background. The background correction was done by subtracting averaged background fluorescence from each ROI in every time point. Time sets of data from each ROI, corrected for background were further normalized to the baseline fluorescence (F₀). Normalized fluorescence data, were expressed as ΔF/F₀, where ΔF represents the change in fluorescence emission from the baseline fluorescence.

Effects of ALS IgG and control IgG on NO, MDA and GSH levels, antioxidative enzymes gene expression and activities, NADPH oxidase 2 (NOX2) and Na⁺/H⁺ exchanger 1 (NHE1) gene expression and TNF-α gene expression and release were analyzed by comparing the means (absolute values) by one-way ANOVA, followed by Bonferroni post hoc test, with a significance level of p < 0.05. Acute effects of ALS and control IgG on ROS, H₂O₂ and pH were analyzed by comparing the amplitudes in certain time points (normalized fluorescence − ΔF/F₀) by two-tailed Student’s t-test, with a significance level of p < 0.05. All mean values were presented as ±SEM.

Production of NO and the intracellular level of MDA were analyzed as they are relevant oxidative stress indicators. MDA is an end product of cell membrane phospholipids degradation and therefore is used as an index of lipid peroxidation (47). In this set of experiments, we treated BV-2 cells with ALS IgG isolated from nine different patients (seven sALS and two fALS) with an average ALSFRSr 36.6 ± 1.7, aged 54.7 ± 4.0 years and with IgG from age-matched control subjects (three healthy and two disease controls, Table 1), and 24 h after the treatments cell lysates were collected for further examination. As shown in Figure 1A both ALS IgG and control IgG treatments resulted in a significant increase of NO production relative to untreated control (87.13 ± 4.52 and 36.07 ± 4.86 relative to 11.69 ± 0.53 µM; p < 0.001 and p < 0.05, respectively). Furthermore, ALS IgG caused 2.4-fold higher NO production than control IgG (p < 0.001). Similarly, treatment with ALS IgG induced a higher level of MDA compared with untreated cells and cells treated with control IgG (0.64 ± 0.03 relative to 0.31 ± 0.01 and 0.39 ± 0.01 nmol/ml, both p < 0.001; Figure 1B).

To examine the effect of IgG on the antioxidative system in BV-2 cells, we measured the activity of antioxidative enzymes (MnSOD, Cu/ZnSOD, CAT, GR, GPx) as well as the total intracellular glutathione content. BV-2 cells were treated for 24 h with ALS IgG from nine different patients and age matched controls (all same as above, Table 1). Results show that MnSOD and Cu/ZnSOD activities were significantly elevated, in ALS IgG-treated cells compared with untreated control (102.18 ± 3.98 vs. 52.53 ± 2.27 U/mg and 556 ± 15.12 vs. 279.64 ± 3.92 U/mg, respectively, both p < 0.001). In addition, both enzymes’ activities were significantly higher in ALS IgG-treated compared with control IgG-treated cells (1.4-fold, p < 0.001 and 1.2-fold, p < 0.01, respectively; Figures 2A,B). Similarly, CAT activity was increased in BV-2 cells treated with ALS IgG (121.72 ± 6.14 U/mg, that was 1.5-fold higher as compared with the control group, 82.78 ± 1.18 U/mg, p < 0.01, and 1.2-fold higher as compared to cells treated with control IgG, 100.85 ± 5.41 U/mg, but with no statistical significance; Figure 2C). On the other hand, we observed a decreased GPx activity after ALS IgG treatment in comparison with the untreated group (88.42 ± 1.79 vs. 99.28 ± 2.46 U/mg, p < 0.05), however, we did not observe a significant difference from the activity in control IgG (93.35 ± 2.11 U/mg), all shown in Figure 2D. IgG treatments (ALS and control) also increased GR activity compared with the control group (33.33 ± 1.03 U/mg, p < 0.001 and p < 0.05, respectively), while ALS IgG induced a 1.5-fold higher activity (70.44 ± 2.04 U/mg) than control IgG (46.58 ± 1.8 U/mg, p < 0.0001), all shown in Figure 2E. Finally, we measured total intracellular glutathione content and found that it was decreased after both ALS IgG (4.18 ± 0.25 nmol/ml) and control IgG treatments (5.09 ± 0.29 nmol/ml) compared with the untreated group (6.61 ± 0.21 nmol/ml, p < 0.001, p < 0.05, respectively). Although there was no statistical significance, there was a trend of a decrease in total glutathione content after ALS IgG treatment compared with control IgG treatment (Figure 2F).

To further examine the effect of ALS IgG on the antioxidative system of BV-2 cells, we determined the gene expression levels for MnSOD, CAT, and GPx by qRT-PCR. BV-2 cells were treated with human IgG and the mRNA content was determined 4 h after the treatment (Figure 3). In this set of experiments, we used IgG from 6 different ALS patients (ALSFRSr 40.3 ± 1.5, aged 65.3 ± 2.8 years) and IgG from three disease controls (aged 65.3 ± 4.7 years, Table 1). Analysis of the results shown in Figure 3A indicates that neither ALS IgG nor control IgG induced significant changes in MnSOD expression compared with untreated control. Interestingly, MnSOD gene expression was only upregulated (approx. 3.5-fold) after treatment with ALS IgG from a patient having fALS with ALSFRSr 45 (sample #3 in Table 1). Following a 4 h ALS IgG treatment, there were also no significant changes in expression of CAT and GPx genes (Figures 3B,C).

Furthermore, we wanted to examine the effect of ALS IgG on the gene expression for NOX2, an enzyme that produces O₂⁻ and therefore is a potential oxidative stress inducer. We did not observe any notable difference in gene expression after ALS IgG treatment when compared to either control IgG treated or untreated control group. However, ALS IgG from the fALS patient with ALSFRSr 45 (sample #3 in Table 1) induced an approx. threefold increase in NOX2 gene expression as compared to control IgG (Figure 3D), resembling the effect on MnSOD gene expression (Figure 3A). Finally, we analyzed the effect of ALS IgG on gene expression of NHE1which may be a regulator of NOX2 activity (48), however, we did not find any significant alterations when compared either with control IgG or the untreated control.

The inflammatory potential of ALS IgG was evaluated by determining their effect on the TNF-α release and gene expression following 24 and 4 h treatments, respectively, using the same samples as in evaluating activity of antioxidative enzymes. ALS IgG-treatment induced a rise in TNF-α release (454.31 ± 42.75 pg/ml) compared with control IgG-treated group (160.88 ± 10.37 pg/ml, p < 0.05) as well as with the untreated group (15.15 ± 9.76 pg/ml, p < 0.01), as seen in Figure 4A. The results of the TNF-α gene expression experiments showed no difference between the groups. Interestingly, ALS IgG from the fALS patient with ALSFRSr 45 (sample # 3 in Table 1) caused a marked increase in TNF-α gene expression (Figure 4B).

Previous results have shown that ALS IgG increase the markers of oxidative stress, but also enhance the antioxidative system in BV-2 cells after 24 h treatment. Although control IgGs differ in effect from ALS IgG, the treatment with control IgG increased NO production, as well as the activity of Mn and Cu/ZnSOD and GR, and decreased total glutathione content. Since both ALS and control IgGs affected BV-2 cells after 24 h treatment, we were interested to evaluate the localization of human IgG in BV-2 cells following 24 h incubation. For this purpose, cells were fixed after treatment with human IgG (three ALS, two control IgG, details in the Table 1), and stained with WGA, the membrane marker, while IgGs were visualized with fluorescent secondary antibody against human IgG. Interestingly, both ALS and control IgGs showed similar staining pattern on BV-2 cells, and these humoral immune factors could be visualized mostly on the cellular membrane (Figure 5). We did not detect any substantial changes in the shape of BV-2 cells in any of the treatments compared to the untreated control (data not shown). The cells were mostly of round morphology in the middle focal plane (as represented in the micrographs of Figure 5).

All the effects of ALS IgGs so far were evaluated after 4 or 24 h treatments. Hence, we were interested to see if there are any acute effects of human IgG on BV-2 cells. For this purpose, we used an epifluorescent imaging system, and plasmids coding for probes sensitive to peroxide (HyPer) and pH (both HyPer and SypHer). In this set of experiments, we used samples from 11 ALS patients (9 sALS and 2 fALS) with an average ALSFRSr 38.0 ± 1.6, aged 58.1 ± 3.6 years and IgG from age-matched control subjects (3 healthy and 2 disease controls, Table 1). Four ALS IgG (36.36%; three sALS and one fALS) induced acute rise in HyPer fluorescence intensity immediately after the application of IgG that continued to increase throughout the 5 min period, decreased slowly during the wash period, and rose again during the application of 100 µM H₂O₂ (Figures 6A–D, red traces). HyPer is a very sensitive indicator of peroxide, but since it is also sensitive to pH changes, control experiments were done with the same ALS samples on BV-2 cells transfected with SypHer that reacts only to the pH changes (peroxide sensing part in the plasmid is mutated). Interestingly, all four ALS samples induced an increase in SypHer intensity in the similar manner (although with the lower averaged amplitude), the intensity decreased during the washing period, and as expected, no change in fluorescence intensity could be detected in response to 100 µM peroxide (Figures 6A–D, blue traces). None of the control IgGs induced changes in HyPer intensity (example shown in Figure 6E). Since both plasmids have the same spectral characteristics, their response could be evaluated only on separate coverslips with cells, so we wanted first to evaluate the similarity of response of these two sensors to the same pH perturbation, induced by 30 mM NH₄Cl. As expected, ammonium-chloride induced a very sharp increase in the fluorescence intensity (indicating increase in the cytosolic pH) followed by a slow decay during the stimulation with 30 mM NH₄Cl (Figure 6F). The removal of ammonium-chloride was accompanied by sharp decay in fluorescence intensity (indicating decrease in cytosolic pH) that slowly returned to the baseline (Figure 6F). The amplitudes of maximal normalized fluorescence intensity in response to alkalization were measured in both cases of transfected cells, with HyPer (0.48 ± 0.02; n = 4) and SypHer (0.55 ± 0.03; n = 6), and the analysis showed no difference between the two groups (two-tailed t-test, p = 0.145; Figure 6G). Since the two sensors react in the same manner to pH changes, the attribution of peroxide to HyPer intensity changes could be determined by subtraction of SypHer from HyPer intensity. By comparing the intensities in the fifth minute after ALS IgG application (maximal measured intensity), the difference between HyPer and SypHer was greater than 10% only in the case of sample #7 (Figure 6B, 17% difference).

In addition, several experiments were conducted with carboxy-H2DCFDA, a general ROS indicator. In this set of data, we used three ALS samples and one control (see Table 1 for details). One ALS and one control IgG induced slow rise in the fluorescence intensity of the ROS probe (Figure 6H). Note that the increase in fluorescence intensity started later for the control IgG (Figure 6H, orange trace), and remained on the same level during the washing step, while the generation of ROS continued even during washing in the case of ALS IgG (Figure 6H, green trace). The amplitudes of ALS- and control IgG-induced normalized fluorescence intensities (reflecting ROS generation) are statistically different throughout the application and wash period (two-tailed t-test on values in the same time points). Linear regression analysis showed also the difference in slopes during the IgG application (p < 0.0001; F = 347.42, DFn = 1, DFd = 19598). Two of the ALS samples did not induce any changes (data not shown).

TNF-α is a proinflammatory cytokine that is released in large amounts by microglia in pathological conditions, and this de novo production is considered to be an important feature of neuroinflammatory response associated with neurodegenerative diseases (65). While an earlier study has shown that intraperitoneal injection of ALS IgG in mice resulted in the activation of microglia (66), a more recent study detected an increased release of TNF-α in the spinal cords of animals inoculated with ALS IgG (24). Our experiments are in line with previous finding, as we demonstrated a significant release of TNF-α after 24 h treatment of BV-2 cells with ALS IgG (Figure 4A), and stress out the contribution of microglia to the ALS IgG-induced inflammatory response. Elevated levels of TNF-α could potentiate glutamate excitotoxicity, a well-established fact in ALS, either influencing the balance of excitatory and inhibitory receptors on neurons, or indirectly by inhibiting glutamate uptake by astrocytes (65). Our immunocytochemistry experiments showed that both ALS IgG and control IgG have similar binding patterns on the cell surface of the BV-2 cells (Figure 5). Therefore, we excluded the possibility that effects on TNF-α release, as well as the other results presented in this study, were due to direct intracellular effects of these humoral factors. However, further experiments have to be carried out in order to reveal the differences in the signaling pathways of ALS IgG and control IgG. In our study, no difference was detected in gene expression level of TNF-α (Figure 4B), which is probably due to a relatively short treatment with IgG (4 h). Our previous study on BV-2 cells has shown that LPS induced changes in TNF-α gene expression (67), which is in disagreement with this finding. However, this difference may be due to different types of activator molecules and therefore possible differences in signaling pathways. Nevertheless, we were able to detect a dramatic increase in TNF-α gene expression of BV-2 cells even after 4 h of treatment with IgG from a fALS patient (sample #3, Table 1) with mutation in SOD1 and ALSFRSr 45 (red dot, Figure 4B). This outlier result may indicate a potential functional difference in IgGs from sALS and fALS patients that prompts a future study toward a differential disease marker. Notably, it was found that a variant of exogenous mutant SOD1 (G93A) protein could also induce a release of TNF-α in microglia that could induce toxicity to motor neurons via TLR and CD14 pathways (68). Taken together, further investigations are necessary in order to determine the signal cascade(s) that ALS IgG initiates in microglial cells, and potential differences or similarities in the mechanism(s) of action of sALS and fALS IgG.

As a further proof of microglial humoral activation the results of this study show that the NO production was significantly increased in BV-2 cells by ALS IgG (Figure 1B). These results are in agreement with a previous study on the ALS animal model showing that the disease progression is followed by an increase in the number of microglial cells expressing inducible nitrate oxide synthase (69). While the effect of NO might be overestimated in vitro where there is no drain for this molecule, NO diffuses to red blood cells in tissue within seconds, where interacts with oxyhemoglobine (70). Nevertheless, while NO is moderately toxic, high concentrations may out-compensate SOD, and it may react with the superoxide anion to form peroxynitrite, that can selectively oxidize certain biological macromolecules, a mechanism pointing to a common cause of in vitro motor neuron death (71). In addition, we observed a significant increase of the index of lipid peroxidation (MDA production; Figure 1B), another confirmation of our hypothesis that ALS IgG can induce oxidative stress, that was in agreement with results obtained in vivo and ex vivo from brainstem and hippocampus of hSOD1(G93A) rats (32). Wu et al. (61) showed that NOX is activated in spinal cords of ALS patients and animal model of the disease, where NOX driven oxidant products from activated microglial cells impair the survival pathways of motor neurons, linking oxidative stress with neuronal death in ALS. Interestingly, our study has found increased NOX2 expression even after 4 h of treatment with one fALS (SOD1D90A) IgG (red dot Figure 3D), stressing out the need for additional experiments that might define the mechanism(s) underlying potential functional differences between familial and sporadic form of the disease.

This study shows that the activity of MnSOD and Cu/ZnSOD was elevated after the treatment with ALS IgG (Figures 2A,B), as well as CAT (Figure 2C), but without a change in gene expression after 4 h treatment (Figure 3B). As a consequence of oxidative stress in an in vitro model of microglial activation with LPS, protein expression of MnSOD and Cu/ZnSOD was also increased (72). Therefore, we can assume that the antioxidative defense in BV-2 cells is upregulated as a result of the microglial activation and/or oxidative stress induced by ALS IgG. Analysis of gene expression of MnSOD after 4 h treatment did not show any change, although again the same outlier sample from the fALS patient (score 45) induced an increase in gene expression. Interestingly, as this fALS patient has mutated SOD1, these data are in agreement with elevated activity of MnSOD in the brainstem and hippocampus of hSOD1G93A rats, already in the presymptomatic stage (32). This could be explained as a compensatory mechanism of antioxidative defense when SOD1 activity is reduced due to mutation, and oxidative stress is most probably increased. In addition, we found that the total glutathione level was significantly decreased due to ALS IgG treatment compared with untreated cells (Figure 2F). A characteristic of in vitro microglial stimulation with LPS and IFNγ, as well as in oxidative stress is a decrease in total glutathione level (73, 74). These data underline that ALS IgG induces oxidative stress in BV-2 cells. Regarding the antioxidative defense an upregulation was observed as a rise in GR activity following the ALS IgG treatment (Figure 2E). On the other hand, we observed a significantly decreased GPx activity (Figure 2D) compared with untreated cells, and there was no change in the gene expression level compared with both groups (Figure 3C). One might speculate that while the increase in the GR activity reflects upregulation of antioxidative system, the decrease in GPx activity might be the consequence of the decrease in substrate, as the total glutathione levels decrease. The role of GPx in ALS, however, is controversial because different studies showed that its activation and expression in spinal cord and cerebral cortex of patients can be decreased (75), without change compared with the control group (76) or not detectable at all (77).

We were unable to detect a change in gene expression for NOX2 after the treatment with ALS IgG (Figure 3D), which is in disagreement with results that show an upregulation of NOX2 in an animal model with the disease progression (61). The possible explanation of this discrepancy could be that it might take more than 4 h of ALS IgG treatment for the change in NOX2 expression or any other gene considered in this study. It is also worth mentioning that the sample from the fALS patient (with mutation in SOD1D90A, and ALSFRSr of 45) induced a rise in gene expression of NOX2, as well as of MnSOD and TNF-α, again indicating a possible differential marker for sALS and fALS that needs to be further studied.

Our imaging experiments on single cell responses to acute application of ALS IgGs revealed an expected heterogeneity of reactions, having in mind the heterogeneity of sALS patients in terms of, e.g., clinical course, disease duration and response to pharmacological treatment. The common denominator of all experiments is that roughly one third of ALS samples show a well-defined rise in the signal for pH-sensitive (SypHer, HyPer) and peroxide sensitive (HyPer) genetically encoded indicators, as well as for ROS-sensitive probe (Carboxy-H2DCFDA). Although not significantly different from pH change in all studied cases, the peroxide response with HyPer was always on top of pH-specific SypHer signal. The ALS IgG-induced ROS generation was significantly higher than control IgGs-induced, as confirmed with carboxy-H2DCFDA. The difference in ROS generation during the washout period between those two types of IgG (Figure 6G, indicated by arrows) might indicate stronger or more specific binding of ALS IgG to the still unknown BV-2 membrane antigen. Nevertheless, alkalization of the cytoplasm induced by ALS IgGs is also a sign of antioxidative defense by way of proton extrusion (78) where NHE1 was suspected as a key transporter (48). We have thus, checked the gene expression for NHE1 after 4 h ALS IgG-treatment, but were unable to detect a change in mRNA (Figure 3E), possibly due to the short treating period. Hence, additional experiments would be needed to confirm or exclude the role of NHE1 in the activation of microglia by ALS IgG.

The present study in an in vitro setup provides a missing link between the oxidative stress in ALS and many different actions of humoral factors in the underlining neuroinflammation. It focuses on microglial cells as key players of the inflammatory response and highlights ALS IgG-induced ROS generation as the trigger of activation of initially healthy microglia. Revealing the ALS IgG signaling cascade in microglial cells could offer valuable molecular biomarkers and/or potential therapeutic targets. Namely, such biomarkers could be followed in microglial cells from patients (i.e., derived form inducible pluripotent stem cells) under no or standard stimulation (e.g., with ATP or H₂O₂) or as shown here, healthy cells (preferably of human origin) could be used to test ALS IgGs for better disease profiling.