Visual abstract

Introduction

Primary cultured neurons, directly isolated and cultured from specific regions of the rat nervous system, closely mimic the in vivo environment to provide physiologically relevant data [1,2]. As such they have broad applicability as both a murine in vitro model for human neurodegenerative disorders such as Alzheimer’s and Parkinson’s disease, and in research on pathological mechanisms and potential therapeutic strategies [3–5].

Neurological functions depend on complex intercellular interactions and therefore, primary cultured neurons constitute a valuable tool for experimental observation and analysis of neuron-neuron interactions, neuronglial cell relationships, and synapse formation [6–9]. In addition, murine models enable physiological evaluation of drug efficacy and cell toxicity, thereby facilitating preclinical verification of the effectiveness and safety of drug candidate compounds [10–12]. However, the process of isolating and culturing neurons from rat neural tissues poses diverse technical challenges. These include appropriate tissue dissociation, optimization of culture conditions, prevention of contamination with other cells, and guided neuronal growth and maturation. In particular, the precise isolation of the requisite neurons and their maintenance necessitate skilled dissection technique, and this poses technical challenges to the use of murine models of primary cultured neurons. Therefore, optimized protocols, which are standardized and reproducible, are essential. This will ensure high cell yield, viability, and neuron-specific purity that will directly affect the culture success rate. Even minor variations in enzyme concentration, dissociation methods, and culture conditions can significantly affect the neuronal culture quality and contribute to interlaboratory inconsistencies in results. Thus, the development of reliable protocols with tissue-specific customization is crucial for establishing robust and functionally relevant in vitro neural murine models. Herein, we present optimized protocols for primary neuron isolation and culture of rat cortex, hippocampus, spinal cord, and dorsal root ganglion (DRG).

Materials and Methods

1. Animals

1.1. Source and handling of rats for cortical neuron isolation

Cortical neurons were isolated from rat embryos on embryonic Days 17–18 (E17–E18). Pregnant Sprague-Dawley rats (Samtako Bio, Republic of Korea; approval no.: JSR-2023-09-001-A, Jaseng Animal Care and Use Committee) were maintained under controlled conditions (12-hour light/dark cycle, constant temperature of 22°C, with ad libitum food and water).

1.2. Source and handling of rats for hippocampal neuron isolation

Hippocampal neurons were isolated on postnatal Days 1–2 (P1–P2) from rat pups (Samtako Bio, Republic of Korea; approval no.: JSR-2024-08-001-A, Jaseng Animal Care and Use Committee). Rat pups were kept with dams for natural feeding and housed under identical controlled conditions as above.

1.3. Source and handling of rats for spinal cord neuron isolation

Spinal cord neurons were isolated from rat embryos on Day 15 (E15). Pregnant Sprague-Dawley rats (Samtako Bio, Republic of Korea; approval no.: JSR-2023-02-003-A-001, Jaseng Animal Care and Use Committee) were maintained under the same controlled conditions.

1.4. Source and handling of rats for DRG neuron isolation

DRG neurons were isolated from 6-week-old young adult rats (Samtako Bio, Republic of Korea; approval no.: JSR-2021-07-003-A-001, Jaseng Animal Care and Use Committee). All animals were housed under the same controlled environment.

2. Reagents

2.1. Reagents for rat cortical neuron culture

Please refer to Table 1.

2.2. Reagents for rat hippocampal neuron culture

Please refer to Table 2.

2.3. Reagents for rat spinal cord neuron culture

Please refer to Table 3.

2.4. Reagents for rat DRG neuron culture

Please refer to Table 4.

2.5. Reagents for immunocytochemistry

Please refer to Table 5.

2.6. Reagent setup

Neuronal culture medium (cortical, spinal cord, and hippocampal neuron): the medium comprises Neurobasal plus medium, 1× P/S, 1× GlutaMAX supplement, and 1× B-27 supplement.

Neuronal culture medium (DRG neuron): the medium comprises F-12 medium, 1× P/S, 10% fetal bovine serum (FBS), and 20 ng/mL nerve growth factor.

Permeabilization solution: this solution comprises phosphate-buffered saline (PBS) and 0.2% (vol/vol) Triton X-100 (to prepare 50 mL solution, add 0.1 mL Triton X-100 in 49.9 mL PBS. Store the solution at 4°C for at least 1 week).

Blocking solution: This solution comprises PBS and 2% (vol/vol) normal goat serum (to prepare 40 mL solution, combine 0.8 mL normal goat serum and PBS to a final volume of 40 mL. Store the solution at 4°C for at least).

3. Materials

3.1. Materials for rat primary neuronal cultures (cortex, hippocampus, spinal cord, and DRG)

Please refer to Table 6.

4. Equipment

4.1. Equipment for rat primary neuron cultures (cortex, hippocampus, spinal cord, and DRG)

Please refer to Table 7.

5. Procedures

5.1. Coating of cell culture plate

The detailed procedure for the coating of the cell culture plate can be found in Table 8 and the Supplementary Materials.

6. Isolation and dissection

6.1. (A) Embryonic rat cortex (duration 2–3 min per embryo)

An ice tray was prepared and a 100-mm cell culture dish was filled with cold Hanks’ balanced salt solution (HBSS) and was placed on top of the ice. Apparatus for the experiment were collected including ethanol-sterilized coating trays, sterile large scissors, tooth forceps, and the euthanasia system/CO₂ chamber (for the sacrifice and dissection of the dam). The dam (E17) was placed in the CO₂ chamber until respiration ceased and the absence of cardiac activity was verified. Subsequently, death was confirmed by applying mechanical stimuli to the foot using tooth forceps at 2–3-second intervals and checking for nociceptive responses. With the dam in a supine position, dissection was performed to separate the embryos which were placed in a 100-mm cell culture dish filled with cold HBSS and kept on ice. Holding #5 fine forceps in each hand, the amniotic membrane was carefully torn to extract the embryos which were transfer to a fresh 60-mm cell culture dish containing cold HBSS. The embryo was placed in a prone position. Using #5 fine forceps, with one hand to immobilize, the neck of the embryo was gently pressed and the skin and skull were carefully removed with the other hand to expose the brain (a caution would be to take extra care to avoid damaging the brain’s morphology whilst removing the skin and skull. Avoid applying direct pressure or puncturing the brain with the forceps, and carefully lift only the skull to maintain brain integrity). The brain was positioned into a dorsal view and, #5 fine forceps were used to carefully divide the cerebrum into hemispheres whilst ensuring that the surrounding tissues, such as the cerebellum, were not included (a caution would be that the positioning the brain in a dorsal view is essential for accurately dividing the cerebral hemispheres without contamination with other tissues. If the brain is positioned in a ventral view, precise division becomes difficult, and this increases the risk of including unwanted tissues). Whilst holding #5 fine forceps in each hand, the meninges surrounding the brain were carefully removed (a caution would be to take extra care to grasp only the meninges to avoid puncturing or damaging the brain. This step requires a high level of skill, as completely removing the meninges whilst preserving the brain’s morphology is crucial. Incomplete meninges removal can reduce neuron-specific purity). The hemispheres were positioned with the inner surface facing up and the C-shaped darker hippocampal structure located in the posterior 1/3 of the hemisphere was identified. Whilst using #5 fine forceps, the hippocampus was carefully isolated and precisely removed. The abovementioned procedure was repeated to obtain the desired number of cells. The cortical tissues were collected in a 15-mL tube containing cold HBSS. Please refer to Figure 1 and Supplementary Video 1 as an aid for the successful isolation of the embryonic rat cortex (a caution would be that the dissection time per embryo should be limited to 2–3 minutes. Considering that each dam typically yields 8–12 embryos, the total dissection time should be kept within 1 hour to maintain a healthy condition for the neurons. Of note, the embryo at TP17 measures approximately 18 mm in width and 10 mm in length, and during the brain isolation process the olfactory bulb may detach automatically; if not, use #5 fine forceps to remove it).

6.2. (B) Postnatal rat hippocampus (duration 2–3 minutes per one pup)

Cell culture dishes (60-mm) were filled cold Dulbecco’s phosphate-buffered saline (DPBS) and placed on ice. The P1–P2 day-old pups were place on an ice pad to induce hypothermia and received isoflurane anesthesia. The rat pup was placed in a prone position and by using #5 fine forceps in both hands, the skin was grasped and pulled to the sides to tear it open. After removing the skin to expose the skull, incisions along the lambdoid and sagittal suture lines were made using #5 fine forceps. Then, the skull was carefully lifted and removed whilst ensuring the brain remained undamaged. The brain was carefully isolated and placed in a 60-mm cell culture dish filled with cold DPBS. The brain was then positioned in a dorsal view and using #5 fine forceps, the cerebrum was carefully divided into hemispheres whilst ensuring that the surrounding tissues, such as the cerebellum, were not included. With #5 fine forceps in each hand, the meninges surrounding the hippocampal tissue were carefully removed (a critical tip is that although the entire meninges covering the brain can be removed, it is time-efficient to selectively remove only the meninges surrounding the hippocampal tissue. However, this requires a high level of skill, and selective removal can be more challenging for some individuals. Additionally, it may be difficult to accurately identify the position of the hippocampus when enclosed by the meninges. Therefore, depending on the researcher’s preference, either selective removal around the hippocampal tissue or complete removal of the meninges can be chosen. The most crucial aspect is to perfectly remove the meninges without causing any damage to the brain tissue). The hemispheres were positioned with the inner surface facing up, and then the C-shaped darker hippocampal structure located in the posterior 1/3 of the hemisphere was identified. Using #5 fine forceps, the hippocampus was carefully isolated and removed. The abovementioned procedure was repeated as necessary to obtain the desired number of cells. The hippocampal tissues were collected in a 15-mL tube containing cold DPBS. Please refer to Figure 2 and Supplementary Video 2 for the isolation process of the embryonic rat cortex (caution lies in the dissection time for each pup which should be limited to 2–3 minutes. To maintain a healthy condition for the neurons, the total dissection time should be less than 1 hour).

6.3. (C) Embryonic rat spinal cord (duration 2–3 minutes per one pup)

An ice bath was prepared and a 100-mm cell culture dish was filled with cold L-15 medium. Following the same protocol used for the isolation of the rat cortex, the E15 pregnant rat was euthanized. The E15 embryo was placed in a supine position. Using #5 fine forceps in one hand, gently pressure was placed around the spine to stabilize the position, and with the other hand, another pair of #5 fine forceps were used to carefully remove the abdominal organs until the vertebrae were exposed. The embryo was laid on its side, and using #5 fine forceps, the head and tail were carefully removed leaving only the torso. The isolated torso was positioned with the ventral side facing upwards. The #5 fine forceps were carefully inserted into the space between the spinal cord and vertebrae to sever the vertebrae. Vannas-Tübingen sprung scissors were used to carefully cut along both sides of the vertebra centered on the spine, and then the vertebral column was isolated (the step of cutting both sides of the spine may be omitted if necessary. A critical tip would be that if the ventral vertebral cut is not performed before isolating the vertebrae, the lack of surrounding structures may cause difficulties in stabilizing the tissue, and thereby pose challenges to vertebral removal. Therefore, following the instructions in order is important). The #5 fine forceps were inserted into the ventrally cut vertebra, the vertebra and spinal meninges were gripped together, and then carefully peeled downwards along the spinal cord. The DRG tissue connected to the dorsal root was also removed (a caution would be that experts may be able to remove the vertebrae and meninges in a single step, however, if they are not fully removed, carefully ensure that any remaining vertebrae and meninges are completely cleared to leave only the spinal cord tissue). The abovementioned procedure was repeated as necessary to obtain the desired number of cells. The spinal tissues were collected in a 15-mL tube containing cold L-15. Please refer to Figure 3 and Supplementary Video 3 for the isolation process of the embryonic rat cortex (a caution would be that the dissection time per embryo should be limited to 2–3 minutes. Considering that each dam typically yields 8–12 embryos, the total dissection time should be less than 1 hour to maintain a healthy condition for the neurons).

6.4. (D) DRG in young adult rats (duration 1 hour for L1–L7 DRG isolation)

An ice bath was prepared and an empty 100-mm cell culture dish was placed on top. Ethanol-sterilized coating trays, sterile large scissors, toothed forceps, a 1-mm rongeur, and a tabletop anesthesia system filled with isoflurane gas were collected. After anesthetizing the rat, a bilateral rib incision was used to expose the heart for cardiac perfusion. Using a 1-mL syringe, 200 μL heparin (1,000 IU/mL) was directly injected into the heart, following 1 minute, the right atrium was cut, and 30 mL HBSS was slowly injected into the left ventricle to flush out the blood (a caution would be that blood contains various immune cells, proteins, and other factors that can interfere with cell culture and potentially affect the health and growth of DRG neurons. Therefore, to reduce the presence of unwanted elements that may compromise the quality and viability of DRG tissue, thorough cardiac perfusion was performed whilst the heart was still functioning. This meant that the animal remained anesthetized because if the solution was injected after the heart stopped, it could not effectively reach the tissue, resulting in insufficient blood removal and poor fixation of the tissue). A laminectomy was performed to remove the vertebrae and expose the DRG. A caution would be that this step should be performed quickly whilst maintaining spinal cord integrity. Neural tissue, primarily composed of fatty material, is very delicate, has a soft fragile texture, and lacks structural support which can cause it to easily collapse or become deformed. Therefore, it is essential to swiftly remove just the vertebrae whilst preserving the shape of the spinal cord. This will allow easy identification of the DRG which is connected to the dorsal root of the spinal cord. If the spinal tissue is extensively damaged or dispersed, or if the dorsal root is severed, accurately locating the DRG becomes challenging owing to difficulties in distinguishing it from the surrounding tissue. Under a dissecting microscope, #5 fine forceps and Vannas-Tübingen sprung scissors were used to isolate and collect the DRG which were then placed in a 60-mm cell culture dish filled with HBSS (a critical tip would be that the primary focus is to accurately locate the DRG without applying pressure or causing any damage with the forceps. When handling the DRG, carefully hold the roots attached to either side of the DRG using #5 fine forceps, and use Vannas-Tübingen sprung scissors to cut the roots to separate the DRG. The isolated DRG tissue will have a candy-like appearance). Using #5 fine forceps, each attached ventral and dorsal root of the DRG tissue were grasped and carefully separated. The meninges surrounding the DRG tissue were peeled and both roots were cut with Vannas-Tübingen sprung scissors. The de-sheathed DRGs were collected in a fresh 60-mm cell culture dish filled with HBSS (a caution would be that this step is crucial for achieving high-purity DRG neuron isolation and requires the highest level of technical skill. The success of primary DRG neuron cultures largely depends on the researcher’s proficiency in this process. A critical tip would be that the DRG is attached to both the dorsal and ventral roots). Using #5 fine forceps, the dorsal and ventral roots were carefully separated to easily remove the surrounding meninges. Subsequently, the meninges surrounding the DRG were removed and the roots were trimmed as close to the DRG tissue as possible. Owing to the lack of surrounding structures to stabilize the tissue (as pressing or applying direct pressure to the DRG tissue with forceps is contraindicated), trimming the root before removing the meninges increases the difficulty in removal. Therefore, the meninges would always be removed first, then the roots would be trimmed, and the isolated DRG tissue would be left behind. The abovementioned procedure was repeated as necessary to obtain the desired number of cells. The DRG tissues were collected in a 15-mL tube containing cold HBSS. Please refer to Figure 4 and Supplementary Video 4 for the successful isolation process of the embryonic rat cortex (a caution would be that the de-sheathing time for each DRG should be less than 2 minutes. Completing the entire process as quickly as possible, ideally within 1 hour, to collect 8–12 trimmed DRGs per sample is recommended).

6.5. Tissue dissociation for single-cell culture

Effective tissue dissociation methods for isolating single cells from cortical, hippocampal, spinal cord, and DRG tissues are provided in the supplementary material.

6.6. Optimized cell seeding for each cell culture dish

Table 8 shows optimized cell seeding:

1. Embryonic rat cortex;

2. Postnatal rat hippocampus;

3. Embryonic rat spinal cord;

4. DRG in young adult rats.

6.7. Maintenance of neurons

The detailed procedure for the maintenance of neurons is provided in the Supplementary Materials.

6.8. Immunocytochemistry of primary neurons

The immunocytochemistry protocol used for primary neurons, including antibody details and staining conditions, is provided in the Table 5 and Supplementary Materials.

Results

Immunocytochemistry was performed to visualize neurons isolated from the cortex, hippocampus, spinal cord, and DRG at Days 3, 7, and 14 of cell culture. Neuronal markers Class 3 beta-tubulin (which is expressed in immature neurons < Day 5), and microtubule-associated protein 2 (expressed more prominently in mature neurons > Day 5) were used for staining. Over time, all neuron types exhibited a progressive increase in neurite branching, neurite length, and the formation of neuronal networks (Figure 5A–5D). DRG neurons displayed a multipolar growth pattern in vitro. For compound efficacy testing, DRG neurons at Day 3 were assessed for neurite outgrowth, and DRG neurons at Day 10 days were tested for the expression of pain-related markers [such as isolectin B4 (IB4), transient receptor potential vanilloid receptor 1 (TRPV1), and calcitonin gene-related peptide (CGRP)]. Cortical neurons ranged in size from 6 to 9 μm, hippocampal neurons from 5 to 9 μm, and spinal cord neurons from 7 to 10 μm. DRG neurons, being the largest, had cell body sizes ranging from 12 to 34 μm. On average, cortical neurons measured 7.9 μm, hippocampal neurons 8.1 μm, spinal cord neurons 9.1 μm, and DRG neurons 21.9 μm (Figure 5E). For DRG neurons, which are representative sensory neurons, expression of pain-related markers (such as IB4, TRPV1, and CGRP) indicated a significant increase in expression after treatment with hydrogen peroxide compared with the untreated control. This suggested a heightened pain signal related response under oxidative stress conditions (Figures 6A–6C).

1. Comparison with existing protocols

1.1. Improved viability

Previous studies have commonly employed single-enzyme treatments such as trypsin, papain, dispase II, or collagenase for tissue dissociation to achieve single-cell suspensions [13–19]. While effective to some extent, these methods often result in lower viability and yield due to the stress exerted on neurons during the dissociation process. In contrast, our protocol employs a more advanced approach, particularly for rat cortical and hippocampal neurons, by utilizing the Neural Tissue Dissociation Kit and Cell Dissociation. This kit ensures a gentler, more effective, and controlled dissociation process compared with traditional methods. Enzymatic degradation is performed twice with different enzymes, and gentle MACS Dissociators (magnetic beads) are used for the mechanical dissociation to produce a single-cell culture. The Neural Tissue Dissociation Kit, comprising proprietary Enzyme Mix 1 and Enzyme Mix 2, offers a multi-step enzymatic digestion tailored for specific neural tissues. For hippocampal neurons, our protocol optimizes the enzymatic process by pre-warming Enzyme Mix 1 (3 mL Earle’s Balanced Salt Solution, 20 U/mL papain, and 100 U/mL DNase 1; total volume 5 mL) and Enzyme Mix 2 (5 mL Earle’s Balanced Salt Solution and 1 μg/mL ovomucoid protease inhibitor with BSA). The sequential application of these Enzyme Mix 1 and 2 effectively minimizes tissue damage while maintaining high viability and reducing cellular debris. Furthermore, by using this kit and associated equipment, our method achieved single-cell suspensions without variability, and these results were reproducible.

1.2. Reduced glial contamination

Previous studies have primarily relied on antimitotic agents such as cytosine arabinoside (AraC) or 5-fluoro-2′-deoxyuridine (FUdR) to limit glial cell proliferation in neuronal cultures [14,20]. While these agents can effectively suppress glial cell growth, they exhibit significant limitations. For instance, AraC has been reported to possess neurotoxic effects, necessitating its use at low concentrations (up to 50 μM), and FUdR requires additional uridine supplementation due to its mechanism of action [14]. In addition, these treatments do not eliminate glial cells entirely, and a residual population remains which can affect neuronal culture purity and functionality. Furthermore, incomplete removal of meninges during the tissue preparation process can exacerbate glial cell proliferation, which AraC and FUdR may partially restrict but, at the cost of neuronal health and imaging quality. In contrast, our protocol incorporates a pre-plating method, particularly for DRG cultures, to enhance neuron-specific purity without relying heavily on potentially neurotoxic agents. In this method, dissociated DRG cells are first seeded onto uncoated cell culture dishes and incubated at 37°C under 5% CO₂ for 1 hour. This step allows non-neuronal cells, including glial cells, to adhere to the dish surface while neurons remain suspended in the medium. After this pre-plating incubation, the neuron-rich cell suspension is collected, gently resuspended, and seeded onto coated dishes containing F12 medium supplemented with 10% FBS, 1% penicillin-streptomycin, and 20 ng/mL nerve growth factor. This approach significantly enhances neuronal purity by physically separating neurons from non-neuronal cells during the pre-plating step.

1.3. Limitations

Whilst this protocol offers significant advantages in terms of neuronal viability, purity, and reproducibility, it is not without its limitations. Firstly, the protocol is optimized specifically for embryonic and neonatal neurons of rat cortical, spinal cord, and hippocampus, making its direct applicability to rat adult tissues limited. Adult neuronal tissues often require different enzymatic and mechanical dissociation approaches due to increased extracellular matrix density and the distinct physiological characteristics of mature neurons. Future studies will need to explore modifications to adapt this protocol for adult neuronal cultures. Secondly, although the protocol demonstrates improved outcomes in terms of neuronal yield and viability as compared to published studies, a direct quantitative side by side comparison with other established dissociation methods such as single-enzyme treatments or alternative dissociation kits has not been conducted. Such comparative analyses would provide a more comprehensive evaluation of the relative advantages of this current method and further substantiate its effectiveness. Addressing these limitations in subsequent investigations will help broaden the applicability and robustness of the protocol.

Supplementary Materials

Supplementary Video 1.

Supplementary Video 2.

Supplementary Video 3.

Supplementary Video 4.

Article information

Author Contributions

Conceptualization: GD. Methodology: GD and DIK. Validation: GD and DIK. Formal analysis: GD, DIK, and JOB. Investigation: GD and JYH. Data curation: GD, DIK, and JOB. Writing - original draft: GD and JYH. Writing - review & editing: JYH. Visualization: DIK. Supervision: JYH. Project administration: JYH.

Conflicts of Interest

The authors have no conflicts of interest to declare.

Ethical Statement

The authors are accountable for all aspects of the work in ensuring that questions related to the accuracy or integrity of any part of the work are appropriately investigated and resolved. All work with animals were approved by Samtako Bio, Republic of Korea; approval no.: JSR-2023-09-001-A, JSR-2024-08-001-A, JSR-2023-02-003-A-001, JSR-2021-07-003-A-001 Jaseng Animal Care and Use Committee.

Data Availability

Figure 1

Sequential images showing the isolation of the brain, hemispheric division, and meninges removal to obtain cerebral cortex tissue for cortical neuron culture.

Figure 2

Sequential images showing the isolation of the brain, hemispheric division, and meninges removal to obtain hippocampal tissue for hippocampal neuron culture.

Figure 3

Sequential images showing the isolation of the spinal cord from the vertebrae and meninges, as well as the removal of DRGs, to obtain spinal cord tissue for spinal cord neuron culture.

DRG = dorsal root ganglion.

Figure 4

Sequential images showing laminectomy, DRG isolation, and meninges removal to obtain DRG tissue for DRG neuron culture.

DRG = dorsal root ganglion.

Figure 5

Immunocytochemistry images and neuron cell size (A) cortical; (B) hippocampal; (C) spinal cord; (D) DRG neurons stained with the neuronal-specific markers [Tuj1 and MAP2 (green)]; and (E) Comparison of cell body sizes among different neuronal types. The bar graph represents the average cell body size (μm) for cortical, hippocampal, spinal cord, and DRG (dorsal root ganglion) neurons. Each bar indicates the mean ± SE of the mean (SEM), with individual data points shown as circles. Images were captured at 10× magnification (white scale bar = 200 μm) and 40× magnification (yellow scale bar = 50 μm).

DRG = dorsal root ganglion.

Figure 6

Representative immunocytochemistry images showing pain-related markers including: (A) IB4 (red); (B) CGRP or TRPV1 (red); and (C) MAP2-positive (green) in 14-day matured DRG neurons treated with H2O2 or without (blank).

Images were captured at 40× magnification (white scale bar = 50 μm).

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Figure & Data

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