Isolation and Whole-Cell Patch-Clamp Recording of Hippocampal Microglia from Adult Mice

Summary

The present protocol describes how to isolate and purify primary hippocampal microglia from adult mice, followed by instructions for conducting whole-cell patch-clamp recordings on these acutely isolated cells.

Abstract

Microglia are resident immune cells in the brain that interact with neurons to maintain the homeostasis of the central nervous system (CNS). Studies show that the microglial surface expresses potassium channels that regulate microglial activation, while abnormalities in these potassium channels can lead to neural diseases. Currently, whole-cell patch-clamp recordings of microglia are mostly performed on cultured primary microglia from fetal or newborn mice due to difficulties in conducting electrophysiological evaluations on acutely isolated microglia. This study introduces an easy-to-follow protocol for isolating hippocampal microglia from adult mice and performing whole-cell patch-clamp recordings on the isolated cells. Briefly, the brain was removed from a mouse after decapitation, the hippocampus was dissected bilaterally, and microglia were isolated using an adult mouse brain dissociation kit. The microglia were then purified using a magnetic-activated cell sorting (MACS) method and seeded onto coverslips. Successful microglial isolation was confirmed by immunofluorescent staining with anti-CD11 and anti-Iba1 antibodies. A cover slip was placed in a recording chamber, and the whole-cell potassium currents of the acutely isolated microglia were recorded under voltage-clamp conditions.

Introduction

Microglia, which derive from myeloid progenitors in the primitive yolk sac, are resident in the CNS and comprise about 10% to 15% of total nerve cells^1,2. Functionally, in addition to surveilling the local environment, performing immune defense functions, and providing nutrition and support for neurons^3,4, microglia can also directly contact neurons to regulate neuronal surface receptors and corresponding ligand binding, and indirectly interact with neurons via the secretion of cytokines⁵. Studies in vitro and in vivo have demonstrated that microglia express various types of potassium channels, which contribute to the maintenance of negative membrane potential⁶ and regulate microglial activation. Given that abnormal microglial potassium channels have been observed in various brain diseases⁷, it is crucial for researchers to identify potential drugs that target these potassium channels and develop strategies to regulate microglial function.

Traditional digestion and purification methods for microglia often use the whole brain, which overlooks the heterogeneity of microglia across different brain areas⁸. In vitro dissociation and long-term culture in serum-containing media place microglia in an active state⁹, which may not accurately reflect their physiological characteristics and true state.

Additionally, while outward rectifier potassium currents have been recorded in primary cultured newborn microglia and cell lines¹⁰, data from long-term cultures of fetal or newborn mouse brain tissue may differ from those of mature microglia. The present protocol aims to acutely isolate microglia and immediately perform whole-cell recordings of microglial potassium currents using adult mouse brain tissue.

Biochemical and potassium channel properties were examined using a method suitable for the isolation and purification of microglia from small tissues^{11,12,13,14,15}. Therefore, this protocol provides guidance for researchers to elucidate the biochemical and functional properties of microglia under physiological and disease conditions. Although this protocol uses the hippocampus and potassium channels as examples, it can also be applied to the study of other brain areas and channel/cell electrophysiological properties.

Protocol

All experiments were approved by the Life Science Animal Care and Use Committee of South China Normal University, and appropriate standards of animal welfare were maintained (ethical approval: SCNU-SLS-2023-048). Male or female 3-5-month-old C57BL/6J mice were used for this study. The details of the reagents and equipment used are listed in the Table of Materials.

1. Preparation of solutions

1. Prepare 50 mL of intracellular solution containing 160 mM of KF, 10 mM of HEPES, 10 mM of EGTA, and 2 mM of MgCl₂. Adjust the pH to 7.2 with 1 M of KOH solution, and adjust the osmotic pressure to 300 mOsm with D-sucrose^13,16. Store at -20 °C.
2. Prepare 50 mL of magnetic-activated cell sorting (MACS) buffer containing 0.5% bovine serum albumin (BSA) and 2 mM of ethylenediaminetetraacetic acid (EDTA) in PBS.
3. Prepare poly-L-lysine (PLL) at a 1:1000 dilution in water.
   NOTE: Coverslips should be coated with PLL the day before the experiment. Incubate the coated slips at 37 °C overnight. Recycle PLL on the second day and dry the coated slips at 37 °C.

2. Preparation of single-cell suspension from brain tissue

1. Anesthetize the mice with 2.5% chloral hydrate (0.15 mL/10 g, i.p.) (following institutionally approved protocols). The disappearance of corneal reflex and pain reflex by pinching the toes or pinching the skin in mice confirm deep anesthesia. Place the animals supine on a dissecting table and pin their limbs.
   1. Expose the chest and perfuse the heart¹⁷ with ice-cold sterile PBS to remove intravascular circulating blood cells. Hold the head and cut the skin along the brain midline with scissors to expose the skull.
   2. Cut the posterior of the skull bilaterally and between the eye sockets with small scissors. Cut the skull along the sagittal suture and remove the skull with tweezers. Extract the brain tissue quickly with tweezers¹⁷.
2. Place the brain tissue in 10 cm culture dishes containing ice-cold PBS. Dissect the cerebral cortex bilaterally with tweezers, isolate the hippocampal tissue, and cut it into smaller pieces. Transfer the pieces into a 15 mL centrifuge tube.
   NOTE: Bilateral dissection of the cerebral cortex reveals the crescent of hippocampal tissue in both hemispheres of the brain.
3. Resuspend the small pieces of hippocampal tissue in 2 mL of ice-cold Hank's balanced salt solution (HBSS) and centrifuge at 300 × g for 3 min (at room temperature), then discard the HBSS.
4. Add 1950 µL of enzyme mixture 1 to the 15 mL centrifuge tube containing the hippocampal tissue pieces. Shake the centrifuge tube continuously in a 37 °C water bath for 5 min.
   NOTE: Enzyme mixture 1 is a combination of 50 µL of enzyme P and 1900 µL buffer Y, preheated to 37 °C (present as a part of the commercially available kit, see Table of Materials).
5. Remove the centrifuge tube from the water bath. Add 30 µL of freshly prepared enzyme mixture 2 to the 15 mL centrifuge tube. Pipette the tissue fragments up and down 20 times with a 1 mL pipette tip, and incubate in a water bath for 5 min with continuous shaking at 37°C.
   NOTE: Enzyme mixture 2 is a combination of 20 µL of buffer Y and 10 µL of enzyme A. The above two digestion steps convert the brain tissue to a cellular suspension. The total digestion time should not exceed 15 min.
6. Remove the centrifuge tube and pipette the suspension up and down multiple times with a 1 mL pipette until no large tissue pieces remain.
7. Filter the cell suspension through a 70 µm cell screening filter and collect the filtrate in a clean 50 mL centrifuge tube.
   NOTE: This step removes any large pieces of undigested tissue.
8. Transfer the filtrate to a new 15 mL centrifuge tube and centrifuge at 4 °C, 300 × g for 10 min. Discard the supernatant with a 1 mL pipette.
9. Wash the cell suspension by pipetting up and down 20 times with 15 mL HBSS, and centrifuge at 4 °C, 300 × g for 10 min. Discard the supernatant and
   resuspend the cell pellet in 1 mL of MACS buffer.
   NOTE: This step provides the single-cell suspension used in subsequent experiments.

3. Acute separation of microglia

1. Transfer 1 mL of the single-cell suspension into a 2 mL microcentrifuge tube and centrifuge at 4 °C, 300 × g for 3 min. Discard the supernatant.
2. Resuspend the single-cell pellet in 90 µL of MACS buffer. Add 10 µL of CD11b/c microbeads and incubate the suspension at 4 °C for 15 min on a horizontal shaker.
3. Place the sorting column in the magnetic field of a suitable MACS separator and moisten the column twice with 500 µL of MACS buffer.
4. Wash the single-cell suspension by adding 1 mL of MACS buffer directly to the tube and centrifuge at 4 °C, 300 × g for 3 min. Aspirate the supernatant completely.
5. Resuspend the cell pellet in 500 µL of MACS buffer.
6. Transfer the resuspended cell suspension to the sorting column in the magnetic field using a 1 mL pipette. Discard the flow-through. Wash the column three times with 500 µL of MACS buffer while it remains in the magnetic field.
   NOTE: The flow-through contains unlabeled cells. The column should be washed without removing it from the magnetic field.
7. Remove the sorting column from the magnetic field. Add 1 mL MACS buffer to the column and slowly push it through with the piston. Collect the flow-through.
   NOTE: The flow-through contains the labeled cells (i.e., the microglia). To improve the purity of the microglia, the process can be repeated with a new sorting column (optional).
8. Seed the selected cells into a 24-well plate containing a coverslip coated with PLL. Add 200 µL of culture medium and incubate for 20 min in a 37 °C incubator with 95% O₂, 5% CO₂, and 60%-80% humidity. This step allows the microglia to attach to the coverslip for subsequent whole-cell patch-clamp and immunofluorescence experiments.

4. Immunofluorescence

1. Aspirate the culture medium from the plate wells and rinse the cells three times with 0.1 M PBS on the shaker.
2. Discard the PBS and fix the microglia on the coverslip with 4% paraformaldehyde (PFA) for 20 min at room temperature (RT).
3. Remove the fixative and rinse the cover slip three times with 0.1 M PBS.
4. Discard the PBS and permeabilize the cells with 200 µL of blocking buffer containing 0.2% Triton X-100 and 5% BSA in PBS for 1 h at RT on the shaker.
5. Incubate the microglia with blocking buffer plus anti-Iba1 and anti-CD11b primary antibodies¹⁸ overnight at 4 °C.
6. The next day, remove the primary antibodies and rinse the cover slip three times with PBS at RT on the shaker.
7. Incubate the microglia with the blocking buffer containing fluorophore-coupled secondary antibodies for 2 h at RT. Add DAPI dye for the last 15 min of incubation.
8. Remove the secondary antibodies and rinse the cover slip three times with PBS. Mount the cover slips on microscope slides for imaging.

5. Whole-cell patch-clamp recording of potassium currents in microglia

1. Thaw the intracellular solution and keep it on ice.
2. Pull patch pipettes (electrodes) from thin-walled borosilicate glass capillaries using a vertical puller¹⁷.
   NOTE: The electrode resistance should be less than 10 MΩ.
3. Transfer a coverslip into the recording chamber and add the extracellular solution containing 160 mM of NaCl, 4.5 mM of KCl, 2 mM of CaCl₂, 1 mM of MgCl₂, and 10 mM of HEPES. Adjust the pH to 7.2 with 1 M NaOH solution before use^13,16, and adjust the osmotic pressure to 300 mOsm with D-sucrose.
4. Locate the spherical microglia under a water microscope using the 40x objective.
5. Fill the electrode with intracellular solution, attach it to the amplifier head-stage of the electrophysiology setup, and apply positive pressure. Switch to the 4x objective and locate the electrode.
   1. Lower the electrode into the bath solution and set the amplifier to voltage-clamp mode to perform the membrane test in bath mode. Switch back to the 40x objective and move the electrode tip closer to the microglia.
      NOTE: When the electrode tip is very close to a microglial cell, the resistance should increase slightly.
   2. Apply a tiny negative pressure to hold the cell. When the resistance reaches 100 MΩ, switch from bath to patch mode in the membrane test. When the resistance reaches 300 MΩ, release the negative pressure.
   3. When the resistance indicates the formation of a gigaohm seal (i.e., >1 GΩ) between the electrode and the microglial membrane, switch from patch to cell mode in the membrane test.
6. Apply a small amount of suction and initiate an electric shock to rupture the membrane. The appearance of large capacitive transients indicates successful membrane rupture.
7. Switch to voltage-clamp mode and open the recording program. Record potassium currents for 200 ms. To test potentials, apply voltage steps in 20 mV increments from -120 mV to +40 mV at 1 Hz.
8. Analyze the obtained data offline using appropriate software.

Results

Briefly, the process involves the isolation of hippocampal microglia from the adult mouse brain followed by whole-cell patch-clamp recording of these cells (Figure 1). The procedure begins with dissecting the hippocampus of 3 to 5-month-old C57BL/6J adult mice. Specifically, the entire brain is removed after perfusion and placed in a culture dish containing ice-cold PBS (Figure 2A). To ensure acute isolation of cells from the hippocampus, the cerebral cortex is removed, and the crescent-shaped hippocampal tissue is isolated intact (Figure 2B).

For optimal cell quality and subsequent high-quality whole-cell patch-clamp recordings, it is crucial to control the digestion time during the preparation of the single-cell suspension. The study consistently manually agitated the hippocampal tissue in the enzyme mixture and strictly limited the digestion period to no more than 15 min (Figure 3). The cells were then isolated using the MACS method, achieving a purity of 86.20% ± 0.68%^19,20. To confirm that these sorted cells were microglia, microglial surface markers were tested by immunofluorescence using anti-Iba1 and anti-CD11b antibodies, which are widely used to detect microglia¹⁸. The results indicated that the isolated cells were CD11b and Iba1 positive, confirming the successful isolation of microglia (Figure 4).

Finally, to assess whether the acutely isolated microglia are of high quality for follow-up experiments, we describe how to perform whole-cell patch-clamp recordings to evaluate microglial potassium currents. The recorded microglia had an average membrane capacitance of 10.42 ± 2.05 (Figure 5). The results demonstrated that microglial potassium currents could be recorded, indicating that the acutely isolated microglia are suitable for neurobiological studies.

Figure 1: Overview of the experimental procedure. There are four main steps: the brain tissues are carefully extracted, and the hippocampus is dissected bilaterally; hippocampus tissues are digested to prepare a single-cell suspension using an enzyme mixture as detailed in the protocol section; the target cells are isolated by a magnetic-activated cell sorting method; and potassium currents in acutely isolated cells are assayed by whole-cell patch-clamp recording. Please click here to view a larger version of this figure.

Figure 2: Dissection of the hippocampus. (A) The whole brain of a 3- to 5-month-old mouse. (B) Isolated bilateral hippocampus. Scale bar: 0.5 cm. Please click here to view a larger version of this figure.

Figure 3: Schematic diagram showing digestion of hippocampal tissues. Illustration of critical experimental steps, ensuring that the hippocampal tissue is digested into a single-cell suspension by adding enzyme mixtures at 37 °C with continuous shaking for up to 15 min. Please click here to view a larger version of this figure.

Figure 4: Identification of acutely isolated hippocampal microglia. Representative confocal images showing CD11b (green) and Iba1 (red) positive cells purified from the hippocampus, confirming that the isolated cells are microglia. Scale bar: 10 µm. Please click here to view a larger version of this figure.

Figure 5: Recording of whole-cell microglial potassium currents. (A) A recording electrode is positioned on the surface of isolated hippocampal microglia. Scale bar: 20 µm. (B) Representative potassium currents of acutely isolated hippocampal microglia under voltage clamp conditions. Please click here to view a larger version of this figure.

Discussion

Cultured microglia from fetal or newborn mice are clearly unsuitable for studying adult microglia. Additionally, given the heterogeneity of microglia across different brain areas²¹, microglia isolated from the whole brain may not accurately represent the characteristics of microglia in a specific brain structure. This protocol provides a method for isolating hippocampal microglia specifically to evaluate the electrical properties of these cells. It includes methods for recording the currents governed by microglial potassium channels.

It is well known that tissue digestion needs to be strictly controlled when preparing single-cell suspensions to avoid compromising the properties of these cells during primary cell culture and subsequent experiments. Typically, a single-cell suspension is obtained from the whole brain through pancreatic enzyme digestion and physical homogenization. However, when isolating microglia from a specific brain area, such as the hippocampus, physical homogenization alone does not yield a sufficient quantity of cells for meaningful experiments. Additionally, pancreatic enzymes have vigorous digestive activity, which can affect cell viability.

To address these issues, the study adopted a relatively mild dissociation procedure and manually controlled the digestion time to no more than 15 min using the adult mouse brain dissociation Kit. This approach maintains the microglia in an active state, which is critical for subsequent whole-cell patch-clamp recordings. The mild dissociation procedure also offers advantages in terms of speed and cost-effectiveness.

Methods for the purification of primary microglia include the stratified shaking method (SSM)²², the density gradient centrifugation method (DGCM)²³, and the fluorescence-activated cell sorting method (FACS)^11,24. Among these, the SSM is widely used for the purification of neurons or glial cells. However, a major limitation of the SSM is its inability to achieve high levels of purity²⁵. Similarly, the DGCM also results in low purity levels. The FACS method requires staining cells for 15 min to 1 h, followed by sorting with a high-speed cell sorter, which can impair cell viability. In contrast, this protocol utilizes the MACS method, which offers a reduced isolation time and avoids the repetitive and time-consuming use of instruments and staining procedures. As a result, high-purity microglia are obtained, though in potentially smaller quantities compared to some alternative methods.

There are limitations to this technique. For instance, the magnetic-activated cell sorting (MACS) method yields relatively few microglia, and acutely isolated microglia cannot be cultured for extended periods. However, this approach is valuable because it allows for the investigation of the electrophysiological properties of microglia that have been freshly prepared from the adult mouse hippocampus, rather than relying on cells that have been cultured for a long time.

The whole-cell patch-clamp technique is a crucial and widely used tool for studying cellular, particularly neuronal, electrophysiological properties in fields such as neuroscience, pharmacology, and cytobiology²⁶. Whole-cell patch-clamp recording has been employed in numerous studies to investigate the properties of acutely dissected brain slices, primary cultures, and cell lines. However, whole-cell patch-clamp recordings of acutely isolated microglia from the adult mouse brain were lacking until our investigations.

This study not only provides a protocol for isolating adult hippocampal microglia but also demonstrates the successful recording of potassium currents in these isolated microglia. Notably, this protocol is likely to apply to the study of microglia in brain structures other than the hippocampus.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
100 mm Petri dish	Corning	353003
15 mL Falcon tubes	BD	352096
24-well plates	BD	353047
35 mm Petri dish	Corning	353001	-
50 mL Falcon tubes	BD	352070
70 μm cell screening	Miltenyi	130-095-823	To remove cell clumps before cell sorting
Adult Mouse Brain Dissociation Kit	Miltenyi	130- 107-677
Anti-CD11b Antibody	Bio-Rad	MCA74	Goat Anti-mouse IgG also available. For blocking endogenous immunoglobulins.
Anti-Iba 1 Antibody	SYSY	234308	Goat Anti-guinea pig IgG) also available. For blocking endogenous immunoglobulins.
Axon Digidata 1440A	USA
Axon MultiClamp 700B Amplifier	USA
BSA Albumin Fraction V	BioFrox	4240GR500	Serum
C57BL/6 mice	Guangdong Medical Laboratory Animal Center
CD11 b/c MicroBeads	Miltenyi	130-105-634
Chloral hydrate	Absin	abs47051394	Widely used as anesthesia in mice
Clampfit 10.6	USA
Confocal Microscope	Zeiss	LSM 800
Culture medium	Fisher Scientific	C11995500BT	-
DAPI dye	Beyotime	C1002
Electrode puller	Narishige	PC-10
HBSS	Servicebio	G4203
Horizontal shaker	SCILOGEX	SLK-O3000-S
Image analysis software	Fiji
MACS Columns	Miltenyi	130-042-201
MACS Separators	Miltenyi	130-042-102
Paraformaldehyde	Fisher Scientific	T353-500	Use fresh 4% solution in 1X PBS, pH 7.2-7.4.
Poly-L-lysine	Beyotime	C0313	Coverslip coating
Triton X-100	Sigma	X100