Excitotoxicity triggered by the over activation of NMDA receptors always induces abnormal [Ca2⁺]_i levels. Nowadays, there are varieties of methods to detect it. Here, some methods based on indirect measurements of free [Ca²⁺]_i levels are described.

An accessible strategy to quantify [Ca²⁺]_i concentrations within the cytosol is to use some [Ca²⁺]_i indicators that upon binding to the ion can emit fluorescence or shift their emission spectrum wavelength. In this case, the sensors are based on the Ca²⁺ chelating properties of ethylene glycol bis (2-aminoethyl) tetraacetic acid (EGTA) [91] and detection using a charge coupled device (CCD) camera. Some sensors can be expressed by the cells using genetic engineering, such as GCaMP or loaded into the cells through membrane permeabilization like Fura-2 and Fluo-4 acetoxymethyl (AM), they vary in their affinity to Ca²⁺, fluorescence emission intensity and stability. Russell in 2011 provided a seminal overview about these techniques [91].

Fluorescent probes, which change their excitation or emission spectra upon Ca²⁺ binding, allow ratiometric measurements. Consequently, measurements are not affected by differences in dye loading between cells. The calculation of the dynamic range is the ratio of the fluorescence intensities recorded in Ca²⁺ saturated and depleted conditions. Although the most frequently used estimation is the signal-to-baseline ratio (SBR), which is the ratio of the change in peak-rest fluorescence (F-F0) over the baseline fluorescence. In order to define the absolute [Ca²⁺]_i concentrations, calibrated values of Ca²⁺ standard levels are plotted against fluorescence intensities and ratios. As mentioned in the previous topics, it is important to use culture medium that does not have high concentrations of glutamate, and in the case of NMDA studies, it is important to supplement with glycine. We recommend recording Ca²⁺ for at least 10 min, because in conditions of excitotoxicity, [Ca²⁺]_i levels do not return to the basal state.

Further to increase experimental data quality, an optional experimental control is the addition of divalent cation ionophores. Ca²⁺ ionophores are lipophilic molecules capable of permeabilizing Ca²⁺ through the plasma membrane. The most common options are ionomycin and Br-A23187 [92]. Both ionophore molecules have distinct kinetic properties that are linked to their affinity in different medium pH [92]. The use of Ca²⁺ ionophores allows the estimation of maximal [Ca²⁺]_i within the cell, and consequently work as an experimental positive control for both fluorescence and luminescence assay, as well as mimicking the overt inflow of Ca²⁺ through NMDAR during excitotoxicity [93].

Besides Ca²⁺ imaging, researchers can use patch-clamping electrophysiology to evaluate excitotoxicity. More specifically, the whole-cell patch-clamp technique can estimate the activity of glutamate-gated ion channels and changes in the cell membrane potential. The whole-cell mode allows the evaluation, in excitable cells, of transmembrane voltage and current dynamics. In voltage-clamp mode, cells are “clamped” at a specific voltage, allowing the recording of currents that flow through ion channels. In current-clamp mode, cells have spontaneous or depolarizing current-induced action potentials recorded [94].

In excitotoxicity studies, the most common parameter analyzed with whole-cell patch clamp is the function of NMDA receptors. Through this assay, studies can explore what the effect of receptors, gene expression, neurotrophic factors modulation has on NMDA receptors [66]. In this assay researchers use for stimulation NMDA combined with glycine, which plays a co-agonist role [95]. Upon agonist binding to the NMDAR, an inward current of Na⁺ and Ca²⁺ flux occurs. For the specific evaluation of NMDAR currents, glycine, GABA subtype A (GABA_A), AMPA, and kainate receptors plus voltage-gated Na⁺ channels are blocked through the application of specific antagonists. Glycine itself is an inhibitory neurotransmitter, acting via specific glycine receptors, and strychnine is applied as an antagonist to block this effect. Bicuculline is used to inhibit GABA_A receptors, 2,3-dihydroxy-6-nitro-7-sulphamoyl-benzo(F)quinoxaline (NBQX) for AMPA and kainate receptors, and tetrodotoxin (TTX) for voltage-gated Na⁺ channels. Although NMDA receptors are physiologically activated by glutamate, they are also coupled to a voltage-dependent condition, carried out by Mg²⁺ blockage of current flow [96]. For this reason, whole-cell patch-clamping experiments aiming to evaluate NMDA currents must have a Mg²⁺ free bath solution.

When studying excitotoxicity, it is also possible to analyze miniature excitatory postsynaptic currents (mEPSC). The mEPSC assay evaluates spontaneous currents, rather than inducing the activity of specific receptors as mentioned before. In this whole-cell patch clamp analysis, it is possible to observe the spontaneous release of glutamate and the excitatory response on the neuron recorded. TTX and bicuculline are necessary to block voltage-dependent excitatory currents and GABA_A receptor-mediated inhibitory currents [97].

One of the major hallmarks of excitotoxicity is mitochondrial dysfunction. Mitochondrial membrane depolarization, ATP quantification and mitochondrial membrane permeability or rupture are the most of them observed parameters when studying excitotoxicity [98]. As mentioned before, the substantial increase of intracellular Ca²⁺ is the key factor for cell death induced by excitotoxicity. Among several pathways that are activated in this process, many are involved with mitochondrial bioenergetics. Besides being responsible for cell respiration and ATP production, mitochondria are considered a major hub for Ca²⁺ homeostasis [99]. Whenever [Ca²⁺]_i rises above the set-point (50–100 nmol/L in neurons), mitochondria promote Ca²⁺ buffering through its uptake, conducted by a specific MCU [100]. Later, concentrations are re-equilibrated between mitochondria and cytosol, through a slow Ca²⁺-Na⁺ exchange [101]. During excitotoxicity conditions, this Ca²⁺ overload induces depolarization of the organelle, triggering cell death mechanisms. Neuronal cells exposed to glutamate that show an ability to overcome this depolarization, recovering mitochondrial potential and energy balance, tend to survive early necrosis stages [102]. The partial inhibition of MCU is considered a possible neuroprotective treatment, but only when mitochondrial Ca²⁺ uptake reaches concentrations between 100 nmol/L and 5 μmol/L [61]. Indeed, the partial knockdown of MCU expression in hippocampal neurons impaired NMDA-induced Ca²⁺ overload in the mitochondria, leading to a neuroprotective effect [103]. On the other hand, a full inhibition of MCU would result in a large impairment of mitochondrial activity, which can lead to a bioenergetic collapse, causing the opposite effect and enhancing excitotoxicity [104]. Thus, mitochondrial depolarization is one of the major indicators of excitotoxicity by the continuous entry of Ca²⁺ in the cell.

Measurement of mitochondrial membrane potential in intact neurons is challenging. Most methodologies rely on fluorescent membrane-permeant cations, although this methodology can have an interference of plasma membrane potential [105]. The most frequently used indicators tetramethylrhodamine methyl ester (TMRM) [64, 106, 107] and rhodamine 123 [60, 61, 63, 64, 66] are accepted as the most reliable fluorescent indicators, with low interference rates in mitochondrial functions [105]. The fluorescent probe JC-1, also used to measure mitochondrial membrane potential in excitotoxicity studies [40, 43, 48], is the least indicated dye to evaluate these dynamics. The JC-1 dye emits fluorescence in the green light spectrum, and when aggregated within the mitochondrial matrix, shifts to a red fluorescence. First, it is inaccurate to calculate a ratio of red aggregated JC-1 and green JC-1 monomers. In addition, when mitochondrial membrane potential decreases, JC-1 aggregates may not dissipate completely back to the green monomer. This combined with a slow permeation compared to hydrophilic dyes, can give incorrect results [105]. There are also commercial kits available that use tetramethylrhodamine ethyl ester (TMRE), an ethyl ester closely related to TMRM [66], or the range of Mitotracker^TM dyes [48, 58]. All these methods do not specifically determine the mitochondrial membrane potential. Yet they provide a semiquantitative estimative when comparing basal vs. treated values. For further reading, Nicholls [105] provided a seminal overview of mitochondrial membrane potential fluorescence methods in neuronal cell culture.

Upon depolarization, the loss of mitochondrial membrane potential can be dangerous for the cell metabolism, mainly due to ATP synthesis impairment. The evaluation of mitochondrial parameters is important to estimate damage extent, as it may lead to necrosis or apoptosis. Cell necrosis occurs when there is a sharp ATP synthesis decrease, resulting in a bioenergetic collapse. On the other hand, apoptosis is facilitated due to membrane rupture by mitochondrial swelling or by outer membrane permeabilization, causing the release of cytochrome c to the cytosol and activating the apoptotic caspase cascade pathway [108]. Mitochondrial activity in intact cells can be assessed by measuring oxygen consumption through high resolution respirometry [40, 61]. The use of specific inhibitors allows us to collect important information about mitochondrial function during these assays. Initially, basal respiration values are detected. After addition of the ATP synthase inhibitor oligomycin, the integrity of mitochondrial inner membrane can be observed, once respiration in this step occurs only due to proton leakage. Next, trifluoromethoxy carbonylcyanide phenylhydrazone (FCCP) or carbonyl cyanide m-chlorophenyl hydrazone (CCCP) addition uncouples the electron transport chain to ATP synthase, in which we see maximal respiration capacity and can observe the function of enzymatic complexes I to IV. And finally, by the inhibition of complexes I (rotenone) and III (antimycin A), oxygen consumption reflects mitochondria-independent oxidative processes [61].

While high resolution respirometry is crucial for analyzing mitochondria activity, it is important to combine it with the measurement of intracellular ATP concentration to provide reliable results. Luciferase-dependent assays are the most common for ATP quantification [40, 48, 51]. It is noteworthy to mention that prolonged mitochondrial depolarization can reverse ATP synthase activity, which then starts hydrolyzing ATP, contributing to bioenergetic collapse. In addition to high resolution respirometry and ATP quantification, excitotoxicity-induced cell death can be detected by mitochondrial cytochrome c, which is released through the permeabilization or rupture of mitochondrial outer membrane. Cytochrome c can be detected by commercial ELISA kits, normally followed by the detection of caspase-3 cleavage to confirm if this release was sufficient to activate caspase cascades [109]. Lastly, although mitochondrial membrane potential, oxygen consumption and ATP quantification are the most used methods to measure mitochondrial impairment in excitotoxicity, there are other important mitochondria-related pathways to consider, such as the activation of nitric oxide synthase and nicotinamide adenine dinucleotide phosphate (NADPH) oxidase [98].

ROS participate in diverse biological events, including excitotoxicity. PSD-95 is a scaffold protein that interacts with c-terminal NR2A or NR2B subunits of the NMDAR and to the residues 108–111 of nNOS. Activation of NMDAR triggers nNOS to produce nitric oxide. During excitotoxicity, over-activation of nNOS promotes production of nitric oxide, ROS and peroxynitrites (ONOO^–). ROS triggers apoptosis by facilitating cytochrome c release through expression and activity upregulation of pore-stabilizing proteins (Bcl-2 and Bcl-xL). ROS detection can be challenging, since ROS concentrations are often in nanomolar range [110].

The commonly used method is the application of fluorescent dyes like fluorescein and rhodamine that were previously modified to “dihydro” colorless, nonfluorescent leuco dyes. Upon oxidation by ROS they turn back to a fluorescent state, serving as an indicator. 2’,7’-dichlorodihydrofluorescein diacetate (H₂DCFDA) is the most common one, and similarly to calcium sensitive dyes they are trapped in the intracellular space after lipophilic blocking groups removal by esterases [111]. Carboxy-H₂DCFDA shows improved cellular retention while the fluorinated molecule (H₂DFFDA) is more photostable, producing less artifacts. Although this is a simple and cheap technique, it is noteworthy to remember that any oxidative factor can increase the signal [112]. Another way to overcome this limitation, is the use of MitoSOX™ Red, which detects superoxides in mitochondria, emitting red light upon oxidation.

Additionally, some genetically encoded probes are available. These can detect ROS changes in real time, like redox sensitive green fluorescent protein (roGFP) [113]. The roGFP is a redox sensor due to the two cysteines added to the original GFP sequence. The oxidized state is a disulfide while dithiol is the reduced. The redox state determines the fluorescence spectrum of the sensor, 488 nm wavelength excites the oxidized form and 405 nm for the reduced form [113]. A good experimental condition for comparison of a positive signal is the addition of H₂O₂ to the sample.

NMDA-induced excitotoxicity triggers the apoptosis pathway [114]. This process, known as “programmed cell death”, is crucial for organism development and homeostasis [115, 116]. Many circumstances and signals regulate apoptosis through some canonical pathways, such as uncontrolled cell proliferation, DNA damage, cell stress and alteration of metabolic pathways [117]. Upon activation, apoptosis pathway leads to DNA fragmentation, chromatin condensation, cytoplasm compaction with plasma membrane bubbles, cellular swelling and apoptotic body formation. Later, immune cells phagocyte these bodies through recognition of the cell surface markers [118].

Furthermore, the three main apoptosis pathways are: (a) the intrinsic pathway begins with intracellular stimuli, such as viral infection and toxic agents, causing permeabilization of the mitochondrial intermembrane space. This event is followed by the release of pro-apoptotic elements to the cytosol, where cytochrome c can form a complex with Apaf-1 and caspase-9, known as apoptosome, that activates a cascade of caspases, resulting in cell death [119]; (b) another intrinsic pathway starts in the ER, where the accumulation of unfolded or misfolded proteins activates ER-proteins sensors, as protein kinase R-like ER kinase (PERK), inositol-requiring enzyme 1 (IRE1a) and activating transcription factor 6 (ATF6) that lead to transcription attenuation, decreased protein synthesis and proteolytic degradation [117, 120], causing cell death; and (c) the extrinsic pathway consists of the interaction between death ligands in specific immune cells and death receptors in the cell destined to die. This interaction leads to a recruitment of adaptor proteins, producing a protein structure called death inducing signaling complex (DISC) that mobilizes and activates procaspases, promoting apoptosis [121]. The published studies in the field of excitotoxicity usually evaluate the outcome, which is cell death. However, the field still lacks a standard method that reliably assesses cell death induced through NMDA excitotoxicity. Among the various methods, we highlight the most frequent ones, such as TUNEL staining, LDH, MTT, and WB for Bax and Bcl. Although all of them are suitable as a cell death detection assay, most powerful methods are here suggested to reliably identify and quantify excitotoxicity-induced apoptotic cell death, as well as the disadvantages of each technique.

Even though cell death can be qualitatively obvious by simple morphological microscopic observation, such as shrinkages, nuclear blebbing and apoptotic bodies [122, 123], this is not enough to measure the extension of the damage in the cell population. There are many quantitative methods to detect apoptosis and they are based on different characteristics or stages of the process. Dye exclusion assays are often used for apoptosis study due to their simplicity and inexpensiveness, and are based on membrane integrity. The most common dye is trypan blue, which enters and stains dead cells, but not the viable ones [124]. However, plasma membrane disruption is a late event of apoptosis; therefore, dye exclusion assays are restricted to later time points [125]. The scientist should account for some technical flaws that occur when cells absorb imperceptible amounts of dye, or cell loss during the process [124].

Since apoptosis frequently involves DNA fragmentation, a common strategy is to examine the different sizes of DNA fragments by agarose gel electrophoresis. In this method, the total genomic DNA samples containing the fragmented DNA are exposed to electric tension and separated by their sizes in agarose gel. The result is a DNA ladder pattern through the gel [126]. Nevertheless, this method is not very accurate, once the DNA fragmentation occurs later in the apoptotic process. In this case, DNA fragmentation should be evaluated at different time points to avoid false negative results [127].

Another technique to determine DNA fragments is the TUNEL. Basically, the method quantifies DNA strand breaks within a cell by the detection of the in situ insertion at free 3’-OH ends with fluorescent labeled nucleotides, such as BrdU and biotinylated deoxyuridine triphosphate (dUTP), mediated by terminal deoxynucleotidyl transferase enzyme activity [128]. Besides the modernity of the method, the TUNEL assay is costly and results in significant number of false positive results [129]. These unreliable outcomes emanate from the fact that the TUNEL assay is not specific for apoptosis, but for DNA damage in general, which can occur in necrosis cell death, highly proliferative cells and cells in specific processes, such as DNA repair and active gene transcription [130, 131].

During healthy and non-stressful conditions, phosphatidylserine (PS) location is within the cytoplasmic cell membrane leaflet surface. Upon apoptosis, two main events are responsible for PS translocation to the outer plasma membrane leaflet: active PS externalization and inhibition of PS internalization. Scramblases translocate PS to the external membrane leaflet surface in order to be engulfed by macrophages [132, 133], followed by inhibition of flippase enzymes responsible for homeostatic maintenance of PS in inner plasma membrane leaflet. A common approach to quantify apoptosis signal is the detection of fluorescent labeled annexin V [134]. Annexin V is a Ca²⁺-dependent phospholipid-binding protein present in humans that shows high affinity to PS. Incubation of fluorescence- or biotin-labeled annexin V with the intact membrane cells, followed by its detection through flow cytometer allows the quantification of the percentage of apoptotic cells [135]. As for the TUNEL assay, the cost of this technique is above the average. In addition, to ensure accuracy, some comparative controls that access basal PS externalization should be conducted in parallel.

The caspase activity assay for cell death analysis focuses on detection of executioner caspase activity, such as caspase-3 and caspase-7. The strategy is to use caspase substrates linked to fluorochromes. Upon cleavage of these substrate, the fluorescence emission rates are recorded and caspase activity is measured by flow cytometry. This technique is very accurate but more expensive than the simpler ones [136, 137]. The immune detection of cleaved caspases through WB is also possible, as well as the expression of other important proteins involved in the apoptosis process, such as Bcl-2, Apaf-1 and cytochrome c [138, 139]. Normally, this apoptosis WB assay is performed simultaneously with the application of a cocktail of apoptosis induction proteins such as p53, cytochrome c, Bcl-2, Apaf-1, PARP 1, CD95, CD261, CD262, CD120b, BAX, BAD, receptor-interacting protein kinase 3 (RIPK3), RIPK1, caspase-3, 7 and 9 and with DAPI staining to identify apoptotic nuclei.

Apart from this, another widely used method to measure neuronal cell death in excitotoxicity experiments is the quantification of cellular metabolic activity. This high throughput assay uses cellular metabolic activity as a proxy for cellular viability, proliferation and cell death. It relies on tetrazolium salt reduction to formazan crystals through mitochondrial dehydrogenases. Compounds for high throughput assays are available as MTT or other reduction salts as shown in Table 1 [140]. Despite their wide application throughout the literature, the use of oxidation-reduction (redox) reagents should be carefully evaluated, because MTT reduction is cytotoxic and its assay alone does not distinguish between proliferation/cell death and metabolic changes. Cells undergoing differentiation programs or under drug treatment may trigger “metabolic” shifts. In view of that, additional assays for verification of cell death induction are needed, like the ones previously mentioned here [141, 142]. Moreover, some drug interventions could potentially introduce experimental bias in the MTT assay, as demonstrated for ionizing radiation, which evoked mitochondrial proliferation and consequently increased MTT reduction, while it also increased the cellular death observed in other methods [142]. Similar effects were observed following drug administration with cholesterol, polyphenols and some protein inhibitors [142–145].

In the excitotoxicity field, the leakage detection of LDH is consistently used as an approach to detect cell death [146]. LDH is a soluble intracellular enzyme released by cells into the extracellular medium when the plasma membrane is disrupted. As in the MTT assay, the LDH method involves redox reactions. However, in this case, the reaction of converting tetrazolium salt into the colored formazan form is provided by the nicotinamide adenine dinucleotide (NADH) that is produced when LDH catalyzes the oxidation of lactate to pyruvate [147]. The extent of formazan formation MTT is colorimetrically measured. The essential point of this assay is detection of necrosis, once this depends on acute membrane permeability, but does not discriminate between primary and secondary necrosis. The difference between primary and secondary necrosis is that the last one occurs as a consequence of the apoptosis process. Considering all this, the LDH assay is capable of detecting apoptosis only in late stages. As for the MTT assay, reagents interfering with the redox reactions or with colorimetric patterns should be averted, such as compounds include oxidizing and reducing agents and even products that can modify the color of the cell culture medium. Common reagents used in cell culture, such as serum and medium, may contain factors, interfering with the MTT assay. Another limitation is that quantities of LDH formation differ between cell types. Therefore, cell death comparison between lineages based on LDH formation is a convoluted concern.

Taken together, the here discussed methods explore distinct mechanisms for assessing viability and quality of living or quantifying dead cells. Every protocol has considerable advantages and disadvantages that should be taken into account for each experiment. For this reason, combining methods based on different detection mechanisms may result in improved results.