Cell culture models to study neurodegeneration

Neurodegenerative diseases are age-dependent pathological disorders characterized by increased cell death, leading to progressive neuronal cell loss [1]. Patients commonly display a wide range of clinical symptoms, including cognitive impairment and motor coordination deficits [2,3]. Since the early 2000s, cell culture has become a standard method in neurobiology for modeling neurodegenerative diseases. This approach offers a stable, controlled environment with minimal systemic variation, advancing cellular and molecular neurobiology by enabling the studies of neuronal development, differentiation, survival, and physiology under genetic or pharmacological manipulation [4,5]. Despite these advantages, cell culture models, ranging from primary neurons to neuronal-like transformed cell lines, have significant limitations. These include the absence of an extracellular matrix, the requirement for anchorage-dependent monolayers, and insufficient gas exchange [6,7]. These inadequacies restrict their applicability and translational relevance, as evidenced by the limited efficacy of existing neurodegenerative therapies in clinical trials [8]. A comparison of the significant differences between cell culture and tissue culture is summarized in Table 1.

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Table 1. Comparative Analysis of Cell Culture and Tissue Culture Techniques. Summary of the advantages and disadvantages of cell culture and tissue culture techniques. Cell Culture refers to monolayer cell cultures, including primary, secondary, or cell lines, whereas Tissue Culture refers to three-dimensional tissue culture. A check mark (✓) indicates an advantage and a cross (×) indicates a disadvantage. Additionally, an asterisk (*) denotes considerations specific to primary cell lines, and a hashtag (#) signifies considerations specific to immortal cell lines.

https://doi.org/10.1371/journal.pone.0342373.t001

The limitations of current models highlight the need for advanced solutions

Preclinical models are currently the most feasible systems for studying age-related neurodegenerative disorders. However, their high costs and limited control over chemical and physical complications present significant drawbacks. Therefore, models that more accurately represent the complexity of in vivo conditions are essential. These models can bridge the gap between preclinical animal models and humans, enabling a better understanding of pathological conditions and potentially leading to more translational outcomes.

Organotypic brain slice culture

Organotypic brain slice culture has emerged as a powerful tool in neurobiology, serving as an alternative to monolayer cell cultures or preclinical models. This method preserves the brain’s complex three-dimensional architecture and cellular composition ex vivo, providing a unique opportunity to investigate intricate neural processes. Various pathophysiological states can be simulated by directly manipulating culture conditions, enabling the study of neurodevelopmental disorders, neurodegenerative diseases, and mechanisms of synaptic plasticity.

Challenges with culturing adult brain tissue

Despite its advantages, organotypic brain slice culture faces challenges, particularly in preparing slices and maintaining their viability [8]. Most studies use rodent brain slices from the first two neonatal weeks, as neonatal tissue demonstrates resistance to ischemic damage, immature metabolic states, and robust dendritic branching [9,10]. However, these characteristics do not accurately mimic age-related neuronal changes. Neonatal brain slices are characterized by resistance to ischemic damage [11], immature metabolic state [12], astrocytic differences [13], amplified synaptic plasticity [14], and robust dendritic branching [15]. To date, studies involving adult brain tissue have shown that slice viability is significantly lower than in neonatal tissue, a trend supported by ordinal logistic regression, which indicates that the odds of longer survival decrease with age. Specifically, juvenile slices (P22–P42) have approximately 16% odds of surviving longer than neonatal slices (P0–P14, OR = 0.155, p = 0.0778). In comparison, adult slices (>P42) have approximately 9% odds (OR = 0.090, p = 0.0176), reflecting a significant reduction in viability with increasing age (Fig 1, Table 2). This corresponds to a median of 7 days ex vivo across all brain regions in adult tissue, compared with 22 days in neonatal tissue [9,16–24]. These findings highlight the need for long-term organotypic brain slice cultures from mature brains to support age-related studies, a gap that our protocol addresses, demonstrating 14-day viability (Fig 1).

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Table 2. Summary of Organotypic Slice Culture Experiments and Outcomes by Brain Region. Data from various studies on organotypic slice cultures detailing the age of the rodents (weeks, w), days ex vivo (DEV), rodent model used, and references with DOIs. The studies are categorized by brain regions: Hippocampus, Cortex, and Cerebellum (Cbl).

https://doi.org/10.1371/journal.pone.0342373.t002

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Fig 1. The lifespan of organotypic brain slice cultures decreases with increasing donor age.

A metadata analysis of 21 published studies reveals that cultured rodent brain slices ex vivo, using 30 mm membrane inserts in 6-well plates, are represented as a dot plot, with days ex vivo (DEV) plotted against donor developmental age (days). Symbols indicate the brain region used for slice culture: (□) Cb = cerebellum, (◇) Ct = cortex, (○) Hc = hippocampus, (◪) Cb^‡ = data from current study as reference and excluded from statistics. Horizontal lines represent median DEV values. To assess the relationship between DEV (ordinal outcome) and donor age group (neonatal, juvenile, adult), independent of brain region, ordinal logistic regression was performed. The model included donor age as the independent variable and DEV as the dependent variable. Model fit was assessed using a likelihood ratio test (Χ² = 6.7, P = 0.0346), and odds ratios were calculated to estimate the effect of donor age on culture longevity. The analysis confirmed that cultures from adult brain tissue exhibit significantly reduced viability, with a median DEV of 7 days across brain regions. In contrast, cultures from neonatal brain tissue have a median DEV of 22 days. The references for the included studies are listed in Table 2.

https://doi.org/10.1371/journal.pone.0342373.g001

Focus on cerebellar tissue

Since their introduction in 1966 [25], organotypic cultures have been developed from various brain regions [8], but the cerebellum remains relatively underexplored. The cerebellum’s unique laminar architecture and specialized circuitry pose significant challenges for culturing organotypic slices [26]. Its cortex comprises three distinct layers: the molecular layer, the purkinje cell layer, and the granule cell layer. Excitatory granule cells receive external inputs and project their axons to connect with the dendrites of inhibitory purkinje cells in the molecular layer. purkinje cells, in turn, relay signals to the cerebellar nuclei, which connect to the brainstem, spinal cord, and other brain regions. Adult cerebellar tissue exhibits reduced neuroplasticity and heightened sensitivity to environmental changes, requiring careful optimization of culture conditions to preserve viability, structural integrity, and functional connectivity [26].

A protocol to enhance cerebellar slice viability from the adult brain

This protocol effectively maintains ex vivo culture of cerebellar brain slices from aged rodents (older than 6 months) for at least 14 days by modifying the medium composition (fig 1, 2a). Key modifications include the use of serum-free media [18] and the anti-inflammatory compound indomethacin [27–31]. These changes improve cerebellar tissue viability and preserve arborization in cerebellar slices, as demonstrated by biochemical and imaging analyses, thus enabling comprehensive analysis of neuronal morphology and function.

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Fig 2. Extended maintenance of OCerSC from aged rodent brain tissue.

(A) Timeline of the culture protocol used to maintain OCerSC from adult rodents for up to 16 days ex vivo (DEV). Slices were initially cultured in Slice Culture Media #1 (with or without serum or indomethacin) for five days under four conditions: horse serum (HS), horse serum plus indomethacin (HS + I), serum-free medium (SF), or serum-free medium with indomethacin (SF + I). On day six, all groups switched to Slice Culture Media #2 (free of indomethacin, with or without serum) and were maintained through day 16, with media changes every 48 hours. The circled points A and B indicate key timepoints, including the first media change at 18–24 hours and the transition to the maintenance medium on day six. Illustration created with BioRender.com. (B) Cytotoxicity, measured by LDH release over time in the SF + I and SF conditions (using the optimized protocol; see Methods), is shown as a spline line plot with individual data points overlaid and summarized as mean ± SD at each time point (N = 3 biological replicates). LDH release was lowest in the SF + I condition. Compared with HS, both HS + I and SF reduced LDH release; however, the greatest reduction was observed with SF + I, which decreased cytotoxicity by 71% relative to HS and 52% relative to SF (based on area under the curve calculations; see Supplementary Table 1). Statistical analysis of LDH time-course data and AUC values using two-way ANOVA (factors: serum and indomethacin) is shown in-panel and detailed in Supplementary Table 1. (C) Densitometry analysis (upper) and representative Western blot (lower) of calbindin expression at 14 DEV across the four initial conditions: SF, SF + I, HS, and HS + I. Densitometry values are shown as individual data points with mean ± SD (N = 4 biological replicates). Calbindin levels were highest in the SF + I condition. Relative calbindin protein levels were analyzed using two-way ANOVA (factors: serum and indomethacin); full statistical results are presented in the panel. Molecular weight markers in kilodaltons (kDa) are indicated for the calbindin Western blot and the total protein on the membrane (Stain-Free) used for normalization. (D) Bright-field images taken at 10 days ex vivo (DEV) to assess cerebellar tissue health under different conditions. A rainbow lookup table (LUT) was applied to the images, where color represents tissue opacity as follows: red = high opacity (less transparent, more degenerated tissue), white = intermediate, dark blue = low opacity (highly transparent, healthier tissue). SF + I slices showed the greatest flattening and transparency, indicated by the largest dark blue regions representing well-preserved central cerebellar lobules. The HS + I condition showed moderate improvement over HS alone, but less than SF + I. These results were confirmed by treating a representative SF + I slice with Triton X-100, which caused delipidation and structural degradation, shifting the color toward higher opacity, serving as a positive control for cytotoxic damage. Overall, initial culture in SF + I provided the best long-term preservation of cerebellar structure.

https://doi.org/10.1371/journal.pone.0342373.g002

Euthanasia

Mice were euthanized if they exhibited critical health deterioration, including head tucking, skin lesions, malocclusions, abnormal breathing, urinary issues, self-mutilation, inability to walk, eat, or drink, agonal breathing, severe ulcerations, or uncontrolled bleeding. In cases of reduced mobility that limited access to food or water, hydrogel and moistened chow were placed on the cage floor. Mice were anesthetized and euthanized using isoflurane (1–3% in oxygen, drop method, as needed) via inhalation, followed by cervical dislocation to confirm death. Death was verified by testing footpad reflexes, in accordance with the recommendations of the American Veterinary Medical Association’s Panel on Euthanasia.

Cerebellar culture

The protocol described in this peer-reviewed article is published on protocols.io (dx.doi.org/10.17504/protocols.io.q26g71428gwz/v1) and included as Supporting Information File 1 at the end of this document. This protocol assessed the effects of media composition on maintaining adult organotypic cerebellar slice cultures (OCerSC) ex vivo for up to 16 days (Fig 2A). Dissection and slicing were performed in ice-cold Hibernate-A medium (Gibco), supplemented with B27 and Glutamax, to minimize excitotoxicity and preserve tissue viability at ambient CO₂ levels without continuous carbogen gassing. Hibernate-A is specifically formulated for short-term storage and handling of postnatal/adult neural tissue, providing stable osmotic and physiological support during benchtop procedures and reducing mechanical stress on fragile, aged cerebellar slices. For the first five days, slices were cultured under four conditions: with or without serum, and with or without the anti-inflammatory agent indomethacin. Starting on day six, all slices were transferred to a common maintenance medium, with or without serum, and maintained through day 16.

Cell cytotoxicity assay

A lactate dehydrogenase (LDH) assay was used to evaluate cytotoxicity in organotypic cerebellar slice cultures (OCerSC). LDH is a stable cytosolic enzyme released when the cell membrane loses integrity, serving as a dependable marker of cytotoxicity [32]. We used the LDH-Glo™ Cytotoxicity Assay (Promega, J2380), a luminescent test that measures LDH activity through linked enzymatic reactions, ultimately producing a luciferin-based bioluminescent signal that correlates with LDH levels.

To ensure precise and specific measurement of tissue-derived LDH release, the following controls and normalizations were applied:

• Background correction: LDH activity was measured in cell-free media samples (identical composition, incubated under the same conditions) and subtracted from experimental values to account for any intrinsic LDH-like activity in the reagents or media components.
• Serum interference control: Baseline LDH levels were determined in fresh serum-free and horse serum-supplemented media formulations before use.
• Normalization to maximal release: To express cytotoxicity as a percentage of total releasable LDH, parallel control wells (triplicate OCerSC at ~14 DIV) were treated with 10 × lysis solution (provided in the kit) or 10% Triton X-100 for 30–60 min to induce complete cell lysis. Media from these wells were collected and processed identically.
• Contamination prevention: All culture media were supplemented with penicillin-streptomycin to minimize bacterial contributions to spurious LDH signals.

Media sampling and processing: At each scheduled full media change (days 1, 2, 4, 8, 12, and 15), 5 µL of conditioned medium was collected from each culture insert and immediately diluted into 95 µL of ice-cold LDH storage buffer (200 mM Tris-HCl pH 7.3, 1% BSA, 10% glycerol) and stored at −20 °C until batch analysis. For the assay, technical triplicates were prepared by mixing 12.5 µL of diluted sample with 12.5 µL of LDH Detection Reagent in white 384-well plates. Reactions were incubated at room temperature for 45 min (protected from light), and luminescence was read on a CLARIOstar plate reader (BMG Labtech). Data are presented as background-corrected relative light units (RLU) or as a percentage of maximal LDH release where indicated.

Western blot and chemifluorescence analysis

Calbindin protein expression in organotypic cerebellar slice cultures (OCerSC) was evaluated using Western blot analysis. Three slices per replicate (N = 4) were lysed in RIPA buffer supplemented with Halt Protease and Phosphatase Inhibitor Cocktail (Thermo Scientific, 78447). Protein lysates were separated on 4–15% Mini-PROTEAN TGX Stain-Free gels (Bio-Rad, 4568084) and transferred to PVDF membranes (Bio-Rad, 1620264). Membranes were blocked in 5% non-fat milk in TBST for 1 hour at room temperature, and incubated overnight at 4°C with calbindin (D1I4Q) XP® Rabbit mAb (1:1000, Cell Signaling Technology, #13176). Subsequently, membranes were incubated with StarBright Blue 700 secondary antibody (Bio-Rad, 12004161) for 1 hour at room temperature. Chemifluorescence signals were detected using a ChemiDoc Imaging System (Bio-Rad). Densitometry analysis was performed using Bio-Rad Image Lab software (version 6.1), with calbindin band intensity normalized to total protein content determined by stain-free membrane quantification.

OCerSC light intensity imaging

To evaluate OCerSC health, bright-field images were captured using the Leica Stereo Microscope MZ 16 with the ORCA-ER CCD camera. Images were processed using Fiji [33]. A thermal lookup table (LUT) was applied during processing to highlight opacity variations across slices. To focus on a specific OCerSC area (designated as the region of interest, ROI), the area outside the ROI was excluded for clearer visualization.

Immunofluorescence staining and imaging

Organotypic cerebellar slice cultures were fixed in 4% paraformaldehyde (PFA) for 20–24 hours at 4 ºC. Slices were then rinsed three times with phosphate-buffered saline (PBS) and either stored in PBS at 4 ºC or processed for immunofluorescence staining. Slices were permeabilized in PBS with 0.5% Triton-X100 for 30 minutes at room temperature, blocked with a 5% BSA solution in PBS with 0.1% Triton-X100 for 90 minutes at room temperature, and incubated with primary antibodies targeting calbindin (D1I4Q; Alexa fluor 488, Cell Signaling Technology #65152S) diluted in a 3% BSA solution containing 0.1% Triton-X100 for 48 hours at 4 ºC. After the primary antibody incubation, slices were rinsed three times with PBS containing 0.1% Triton-X100. During the second wash, NucBlue™ Fixed Cell ReadyProbes™ Reagent (Invitrogen, MA, USA) was added to label nuclei. Slices were mounted onto glass slides and covered with #1.5 coverslips using Prolong™ Diamond Antifade Mountant (Thermo Fisher Scientific, NC, USA), and imaged using an Andor Dragonfly spinning disk confocal with the Zyla Plus 4.2MP sCMOS (Oxford Instruments, Abingdon, United Kingdom), Leica STELLARIS 8 FALCON, or Leica Mica in confocal mode. Images were processed using Imaris Multidimensional Visualization and Analysis software (Schlieren-Zurich, Switzerland) and Fiji, as specified.

Statistical analysis

All statistical analyses were performed using JMP Pro (version 19, SAS Institute, Cary, NC, USA). To examine the relationship between days ex vivo (DEV, ordinal outcome) and donor age group (neonatal, juvenile, adult [34]), independent of brain region, ordinal logistic regression was used (N = 22, 6–9 per donor age group). The model included DEV as the dependent variable and donor age as the independent variable. Model fit was assessed using the likelihood ratio test, and odds ratios were calculated to quantify the effect of age on DEV.

For cytotoxicity and protein expression outcomes, the area under the curve (AUC) of LDH from biological replicates (N = 3 per condition) and relative calbindin expression from Western blots (N = 4 biological replicates, 3 slices per replicate) were analyzed using two-way analysis of variance (ANOVA). The 2x2 factorial design included two main effects-serum (horse serum [HS] vs. serum-free [SF]) and indomethacin (with vs. without)- and their interaction term. Calbindin expression was quantified by densitometry, with band intensity normalized to total protein content. Post hoc comparisons were conducted using Fisher’s Least Significant Difference (LSD) test to identify specific group differences. Assumptions of normality and homoscedasticity were verified using Shapiro-Wilk and Levene’s tests, respectively. All tests were two-tailed with a significance threshold of α = 0.05. Data are reported as means ± standard deviation.

Ethics declarations

All animal work was conducted in accordance with the Guide for the Care and Use of Laboratory Animals under approved IACUC animal use protocols within the AAALAC-accredited program at the University of North Carolina at Chapel Hill. NIH/PHS Animal Welfare Assurance Number: D16-00256 (A3410-01); USDA Animal Research Facility Registration Number: 55-R-0004; and AAALAC Institutional Number: #329. All animals were euthanized in compliance with the governing protocol.