This work was supported by grants from the National Natural Science Foundation of China (32170950, 32371065), the Natural Science Foundation of Guangdong Province, Nos. 2023A1515010899 and 2021A1515010804.

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
<ol>
	<li>Prepare 50 mL of intracellular solution containing 160 mM of KF, 10 mM of HEPES, 10 mM of&#160;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-sucrose13,16. Store at -20 &#176;C.</li>
	<li>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.</li>
	<li>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 &#176;C overnight. Recycle PLL on the second day and dry the coated slips at 37 &#176;C.</li>
</ol>
2. Preparation of single-cell suspension from brain tissue
<ol>
	<li>Anesthetize the mice with 2.5% chloral hydrate (0.15 mL/10 g,&#160;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.
	<ol>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>Resuspend the small pieces of hippocampal tissue in 2 mL of ice-cold Hank&#39;s balanced salt solution (HBSS) and centrifuge at 300 &#215; g for 3 min (at room temperature), then discard the HBSS.</li>
	<li>Add 1950 &#181;L of enzyme mixture 1 to the 15 mL centrifuge tube containing the hippocampal tissue pieces. Shake the centrifuge tube continuously in a 37 &#176;C water bath for 5 min. 
	NOTE: Enzyme mixture 1 is a combination of 50 &#181;L of enzyme P and 1900 &#181;L buffer Y, preheated to 37 &#176;C (present as a part of the commercially available kit, see Table of Materials).</li>
	<li>Remove the centrifuge tube from the water bath. Add 30 &#181;L 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&#176;C. 
	NOTE: Enzyme mixture 2 is a combination of 20 &#181;L of buffer Y and 10 &#181;L 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.</li>
	<li>Remove the centrifuge tube and pipette the suspension up and down multiple times with a 1 mL pipette until no large tissue pieces remain.</li>
	<li>Filter the cell suspension through a 70 &#181;m cell screening filter and collect the filtrate in a clean 50 mL centrifuge tube. 
	NOTE: This step removes any large pieces of undigested tissue.</li>
	<li>Transfer the filtrate to a new 15 mL centrifuge tube and centrifuge at 4 &#176;C, 300 &#215; g for 10 min. Discard the supernatant with a 1 mL pipette.</li>
	<li>Wash the cell suspension by pipetting up and down 20 times with 15 mL HBSS, and centrifuge at 4 &#176;C, 300 &#215; g for 10 min. Discard the supernatant and 
	resuspend the cell pellet in 1 mL of MACS&#160;buffer. 
	NOTE: This step provides the single-cell suspension used in subsequent experiments.</li>
</ol>
3. Acute separation of microglia
<ol>
	<li>Transfer 1 mL of the single-cell suspension into a 2 mL microcentrifuge tube and centrifuge at 4 &#176;C, 300 &#215; g for 3 min. Discard the supernatant.</li>
	<li>Resuspend the single-cell pellet in 90 &#181;L of MACS buffer. Add 10 &#181;L of CD11b/c microbeads and incubate the suspension at 4 &#176;C for 15 min on a horizontal shaker.</li>
	<li>Place the sorting column in the magnetic field of a suitable MACS separator and moisten the column twice with 500 &#181;L of MACS buffer.</li>
	<li>Wash the single-cell suspension by adding 1 mL of MACS buffer directly to the tube and centrifuge at 4 &#176;C, 300 &#215; g for 3 min. Aspirate the supernatant completely.</li>
	<li>Resuspend the cell pellet in 500 &#181;L of MACS buffer.</li>
	<li>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 &#181;L 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.</li>
	<li>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).</li>
	<li>Seed the selected cells into a 24-well plate containing a coverslip coated with PLL. Add 200 &#181;L of culture medium and incubate for 20 min in a 37 &#176;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.</li>
</ol>
4. Immunofluorescence
<ol>
	<li>Aspirate the culture medium from the plate wells and rinse the cells three times with 0.1 M PBS on the shaker.</li>
	<li>Discard the PBS and fix the microglia on the coverslip with 4% paraformaldehyde (PFA) for 20 min at room temperature (RT).</li>
	<li>Remove the fixative and rinse the cover slip three times with 0.1 M PBS.</li>
	<li>Discard the PBS and permeabilize the cells with 200 &#181;L of blocking buffer containing 0.2% Triton X-100 and 5% BSA in PBS for 1 h at RT on the shaker.</li>
	<li>Incubate the microglia with blocking buffer plus anti-Iba1 and anti-CD11b primary antibodies18 overnight at 4 &#176;C.</li>
	<li>The next day, remove the primary antibodies and rinse the cover slip three times with PBS at RT on the shaker.</li>
	<li>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.</li>
	<li>Remove the secondary antibodies and rinse the cover slip three times with PBS. Mount the cover slips on microscope slides for imaging.</li>
</ol>
5. Whole-cell patch-clamp recording of potassium currents in microglia
<ol>
	<li>Thaw the intracellular solution and keep it on ice.</li>
	<li>Pull patch pipettes (electrodes) from thin-walled borosilicate glass capillaries using a vertical puller17. 
	NOTE: The electrode resistance should be less than 10 M&#937;.</li>
	<li>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 use13,16, and adjust the osmotic pressure to 300 mOsm with D-sucrose.</li>
	<li>Locate the spherical microglia under a water microscope using the 40x objective.</li>
	<li>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.
	<ol>
		<li>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.</li>
		<li>Apply a tiny negative pressure to hold the cell. When the resistance reaches 100 M&#937;, switch from bath to patch mode in the membrane test. When the resistance reaches 300 M&#937;, release the negative pressure.</li>
		<li>When the resistance indicates the formation of a gigaohm seal (i.e., &#62;1 G&#937;) between the electrode and the microglial membrane, switch from patch to cell mode in the membrane test.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>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.</li>
	<li>Analyze the obtained data offline using appropriate software.</li>
</ol>

Electrophysiological Recording of Hippocampal Microglia from Adult Mice

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The authors have nothing to disclose.

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 gover...

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 cells1,2. Functionally, in addition to surveilling the local environment, performing immune defense functions, and providing nutrition and support for neurons3,4, 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 tissues11,12,13,14,15. 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.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>100 mm Petri dish</td>
			<td>Corning</td>
			<td>353003</td>
			<td></td>
		</tr>
		<tr>
			<td>15 mL Falcon tubes</td>
			<td>BD</td>
			<td>352096</td>
			<td></td>
		</tr>
		<tr>
			<td>24-well plates</td>
			<td>BD</td>
			<td>353047</td>
			<td></td>
		</tr>
		<tr>
			<td>35 mm Petri dish</td>
			<td>Corning</td>
			<td>353001</td>
			<td>-</td>
		</tr>
		<tr>
			<td>50 mL Falcon tubes</td>
			<td>BD</td>
			<td>352070</td>
			<td></td>
		</tr>
		<tr>
			<td>70 &#956;m cell screening</td>
			<td>Miltenyi</td>
			<td>130-095-823</td>
			<td>To remove cell clumps before cell sorting</td>
		</tr>
		<tr>
			<td>Adult Mouse Brain Dissociation Kit</td>
			<td>Miltenyi</td>
			<td>130- 107-677</td>
			<td></td>
		</tr>
		<tr>
			<td>Anti-CD11b Antibody</td>
			<td>Bio-Rad</td>
			<td>MCA74</td>
			<td>Goat Anti-mouse IgG also available. For blocking endogenous immunoglobulins.</td>
		</tr>
		<tr>
			<td>Anti-Iba 1 Antibody</td>
			<td>SYSY</td>
			<td>234308</td>
			<td>Goat Anti-guinea pig IgG) also available. For blocking endogenous immunoglobulins.</td>
		</tr>
		<tr>
			<td>Axon Digidata 1440A</td>
			<td>USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Axon MultiClamp 700B Amplifier</td>
			<td>USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>BSA Albumin Fraction V</td>
			<td>BioFrox</td>
			<td>4240GR500</td>
			<td>Serum</td>
		</tr>
		<tr>
			<td>C57BL/6 mice&#160;</td>
			<td>Guangdong Medical Laboratory Animal Center</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CD11 b/c MicroBeads</td>
			<td>Miltenyi</td>
			<td>130-105-634</td>
			<td></td>
		</tr>
		<tr>
			<td>Chloral hydrate</td>
			<td>Absin</td>
			<td>abs47051394</td>
			<td>Widely used as anesthesia in mice</td>
		</tr>
		<tr>
			<td>Clampfit 10.6</td>
			<td>USA</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal Microscope</td>
			<td>Zeiss</td>
			<td>&#160;LSM 800</td>
			<td></td>
		</tr>
		<tr>
			<td>Culture medium</td>
			<td>Fisher Scientific</td>
			<td>C11995500BT</td>
			<td>-</td>
		</tr>
		<tr>
			<td>DAPI dye</td>
			<td>Beyotime</td>
			<td>C1002</td>
			<td></td>
		</tr>
		<tr>
			<td>Electrode puller</td>
			<td>Narishige</td>
			<td>PC-10</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS</td>
			<td>Servicebio</td>
			<td>G4203</td>
			<td></td>
		</tr>
		<tr>
			<td>Horizontal shaker</td>
			<td>SCILOGEX</td>
			<td>SLK-O3000-S</td>
			<td></td>
		</tr>
		<tr>
			<td>Image analysis software</td>
			<td>Fiji</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MACS Columns</td>
			<td>Miltenyi</td>
			<td>130-042-201</td>
			<td></td>
		</tr>
		<tr>
			<td>MACS Separators</td>
			<td>Miltenyi</td>
			<td>130-042-102</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Fisher Scientific</td>
			<td>T353-500</td>
			<td>Use fresh 4% solution in 1X PBS, pH 7.2-7.4.&#160;</td>
		</tr>
		<tr>
			<td>Poly-L-lysine</td>
			<td>Beyotime</td>
			<td>&#160;C0313</td>
			<td>Coverslip coating</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma</td>
			<td>X100</td>
			<td></td>
		</tr>
	</tbody>
</table>

isolation and whole-cell patch-clamp recording of hippocampal microglia from adult mice

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.

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 ...

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

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.

Microglia are resident immune cells in the brain that interact with neurons to maintain the homeostasis of the central nervous system (CNS). Studies show ...

isolation-whole-cell-patch-clamp-recording-hippocampal-microglia-from

Research

JoVE Journal

Biology

594 Views.  South China Normal University. 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.

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

Methods for Patch Clamp Capacitance Recordings from the Calyx

We demonstrate the basic techniques for presynaptic patch clamp recording at the calyx of Held, a mammalian central nervous system nerve terminal. Electrical recordings from the presynaptic terminal allow the measurement of action potentials, calcium channel currents, vesicle fusion (exocytosis) and subsequent membrane uptake (endocytosis). The fusion of vesicles containing neurotransmitter causes the vesicle membrane to be added to the cell membrane of the calyx. This increase in the amount of cell membrane is measured as an increase in capacitance. The subsequent reduction in capacitance indicates endocytosis, the process of membrane uptake or removal from the calyx membrane. Endocytosis, is necessary to maintain the structure of the calyx and it is also necessary to form vesicles that will be filled with neurotransmitter for future exocytosis events. Capacitance recordings at the calyx of Held have made it possible to directly and rapidly measure vesicular release and subsequent endocytosis in a mammalian CNS nerve terminal. In addition, the corresponding postsynaptic activity can be simultaneously measured by using paired recordings. Thus a complete picture of the presynaptic and postsynaptic electrical activity at a central nervous system synapse is achievable using this preparation. Here, the methods for slice preparation, morphological features for identification of calyces of Held, basic patch clamping techniques, and examples of capacitance recordings to measure exocytosis and endocytosis are presented.

We demonstrate the basic techniques for presynaptic patch clamp recording at the calyx of Held, a mammalian central nervous system nerve terminal.

We demonstrate the basic techniques for presynaptic patch clamp recording at the calyx of Held, a mammalian central nervous system nerve terminal. ...

Whole Cell Recordings from Brain of Adult Drosophila

In this video, we demonstrate the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons. We begin by describing the dissecting solution and capture of the adult females used in our studies. The procedure for removing the whole brain intact, including both optic lobes, is illustrated. Dissection of the overlying trachea is also shown.  The isolated brain is not only small but needs special care in handling at this stage to prevent damage to the neurons, many of which are close to the outer surface of the tissue. We show how a special holder we developed is used to stabilize the brain in the recording chamber. A standard electrophysiology set up is used for recording from single neurons or pairs of neurons. A fluorescent image, viewed through the recording microscope, from a GAL4 line driving GFP expression (GH146) illustrates how projection neurons (PNs) are identified in the live brain. A high power Nomarski image shows a view of a single neuron that is being targeted for whole cell recording. When the brain is successfully removed without damage, the majority of the neurons are spontaneously active, firing action potentials and/or exhibiting spontaneous synaptic input. This in situ preparation, in which whole cell recording of identified neurons in the whole brain can be combined with genetic and pharmacological manipulations, is a useful model for exploring cellular physiology and plasticity in the adult CNS.

This video demonstrates the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons using standard whole cell technology. It includes images of GFP labeled cells and neurons viewed during recording.

In this video, we demonstrate the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons. We begin by ...

Patch Clamp Recording of Ion Channels Expressed in  Xenopus Oocytes

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological measurement of single or multiple ion channels in cells. This technique can be applied to ion channels in both their native environment and expressed in heterologous cells, such as oocytes harvested from the African clawed frog, Xenopus laevis. Here, we describe the well-established technique of patch clamp recording from Xenopus oocytes. This technique is used to measure the properties of expressed ion channels either in populations (macropatch) or individually (single-channel recording). We focus on techniques to maximize the quality of oocyte preparation and seal generation. With all factors optimized, this technique gives a probability of successful seal generation over 90 percent. The process may be optimized differently by every researcher based on the factors he or she finds most important, and we present the approach that have lead to the greatest success in our hands.

This is intended as an introduction to patch clamp recording from Xenopus laevis oocytes. It covers vitelline membrane removal, formation of a gigaohm seal (gigaseal), and the optional conversion of the patch to the outside-out topology.

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological ...

Pressure-polishing Pipettes for Improved Patch-clamp Recording

Pressure-polishing is a method for shaping glass pipettes for patch-clamp recording. We first developed this method for fabricating pipettes suitable for recording from small (&lt;3 m) neuronal cell bodies. The basic principal is similar to glass-blowing and combines air pressure and heat to modify the shape of patch pipettes prepared by a conventional micropipette puller. It can be applied to so-called soft (soda lime) and hard (borosilicate) glasses. Generally speaking, pressure polishing can reduce pipette resistance by 25% without decreasing the diameter of the tip opening (Goodman and Lockery, 2000). It can be applied to virtually any type of glass and requires only the addition of a high-pressure valve and fitting to a microforge. This technique is essential for recording from ultrasmall cells (&lt;5 m) and can also improve single-channel recording by minimizing pipette resistance. The blunt shape is also useful for perforated-patch clamp recording since this tip shape results in a larger membrane bleb available for perforation.

This is a guide to modifying the shape of glass micropipettes. Specifically, by using heat and air pressure the taper is widened without increasing the tip opening, leading to lower pipette resistance. This is critical to obtain low noise recordings of small cells but is useful in many applications.

Pressure-polishing is a method for shaping glass pipettes for patch-clamp recording.  We first developed this method for fabricating pipettes suitable for ...

Dissection of Hippocampal Dentate Gyrus from Adult Mouse

The hippocampus is one of the most widely studied areas in the brain because of its important functional role in memory processing and learning, its remarkable neuronal cell plasticity, and its involvement in epilepsy, neurodegenerative diseases, and psychiatric disorders. The hippocampus is composed of distinct regions; the dentate gyrus, which comprises mainly granule neurons, and Ammon's horn, which comprises mainly pyramidal neurons, and the two regions are connected by both anatomic and functional circuits. Many different mRNAs and proteins are selectively expressed in the dentate gyrus, and the dentate gyrus is a site of adult neurogenesis; that is, new neurons are continually generated in the adult dentate gyrus. To investigate mRNA and protein expression specific to the dentate gyrus, laser capture microdissection is often used. This method has some limitations, however, such as the need for special apparatuses and complicated handling procedures. In this video-recorded protocol, we demonstrate a dissection technique for removing the dentate gyrus from adult mouse under a stereomicroscope. Dentate gyrus samples prepared using this technique are suitable for any assay, including transcriptomic, proteomic, and cell biology analyses. We confirmed that the dissected tissue is dentate gyrus by conducting real-time PCR of dentate gyrus-specific genes, tryptophan 2,3-dioxygenase (TDO2) and desmoplakin (Dsp), and Ammon's horn enriched genes, Meis-related gene 1b (Mrg1b) and TYRO3 protein tyrosine kinase 3 (Tyro3). The mRNA expressions of TDO2 and Dsp in the dentate gyrus samples were detected at obviously higher levels, whereas Mrg1b and Tyro3 were lower levels, than those in the Ammon's horn samples. To demonstrate the advantage of this method, we performed DNA microarray analysis using samples of whole hippocampus and dentate gyrus. The mRNA expression of TDO2 and Dsp, which are expressed selectively in the dentate gyrus, in the whole hippocampus of alpha-CaMKII+/- mice, exhibited 0.037 and 0.10-fold changes compared to that of wild-type mice, respectively. In the isolated dentate gyrus, however, these expressions exhibited 0.011 and 0.021-fold changes compared to that of wild-type mice, demonstrating that gene expression changes in dentate gyrus can be detected with greater sensitivity. Taken together, this convenient and accurate dissection technique can be reliably used for studies focused on the dentate gyrus.

A dissection technique for removal of the dentate gyrus from adult mouse under a stereomicroscope was demonstrated in this video-recorded protocol.

The hippocampus is one of the most widely studied areas in the brain because of its important functional role in memory processing and learning, its ...

Isolation and Culture of Pulmonary Endothelial Cells from Neonatal Mice

Endothelial cells provide a useful research model in many areas of vascular biology. Since its first isolation 1, human umbilical vein endothelial cells (HUVECs) have shown to be convenient, easy to obtain and culture, and thus are the most widely studied endothelial cells. However, for research focused on processes like angiogenesis, permeability or many others, microvascular endothelial cells (ECs) are a much more physiologically relevant model to study 2. Furthermore, ECs isolated from knockout mice provide a useful tool for analysis of protein function ex vivo. Several approaches to isolate and culture microvascular ECs of different origin have been reported to date 3-7, but consistent isolation and culture of pure ECs is still a major technical problem in many laboratories. Here, we provide a step-by-step protocol on a reliable and relatively simple method of isolating and culturing mouse lung endothelial cells (MLECs). In this approach, lung tissue obtained from 6- to 8-day old pups is first cut into pieces, digested with collagenase/dispase (C/D) solution and dispersed mechanically into single-cell suspension. MLECS are purified from cell suspension using positive selection with anti-PECAM-1 antibody conjugated to Dynabeads using a Magnetic Particle Concentrator (MPC). Such purified cells are cultured on gelatin-coated tissue culture (TC) dishes until they become confluent. At that point, cells are further purified using Dynabeads coupled to anti-ICAM-2 antibody. MLECs obtained with this protocol exhibit a cobblestone phenotype, as visualized by phase-contrast light microscopy, and their endothelial phenotype has been confirmed using FACS analysis with anti-VE-cadherin 8 and anti-VEGFR2 9 antibodies and immunofluorescent staining of VE-cadherin. In our hands, this two-step isolation procedure consistently and reliably yields a pure population of MLECs, which can be further cultured. This method will enable researchers to take advantage of the growing number of knockout and transgenic mice to directly correlate in vivo studies with results of in vitro experiments performed on isolated MLECs and thus help to reveal molecular mechanisms of vascular phenotypes observed in vivo.

Here, we describe a protocol for isolation and culture of murine pulmonary endothelial cells. This method comprises mechanic and enzymatic lung tissue dissociation as well as a 2-step purification process using anti-PECAM-1 and anti-ICAM-2 antibodies conjugated to magnetic beads, which produces a pure endothelial cell population of mostly microvascular origin.

Endothelial cells provide a useful research model in many areas of vascular biology. Since its first isolation 1, human umbilical vein endothelial cells ...

Isolation and Culture of Adult Epithelial Stem Cells from Human Skin

The homeostasis of all self-renewing tissues is dependent on adult stem cells. As undifferentiated stem cells undergo asymmetric divisions, they generate daughter cells that retain the stem cell phenotype and transit-amplifying cells (TA cells) that migrate from the stem cell niche, undergo rapid proliferation and terminally differentiate to repopulate the tissue.
Epithelial stem cells have been identified in the epidermis, hair follicle, and intestine as cells with a high in vitro proliferative potential and as slow-cycling label-retaining cells in vivo 1-3. Adult, tissue-specific stem cells are responsible for the regeneration of the tissues in which they reside during normal physiologic turnover as well as during times of stress 4-5. Moreover, stem cells are generally considered to be multi-potent, possessing the capacity to give rise to multiple cell types within the tissue 6. For example, rodent hair follicle stem cells can generate epidermis, sebaceous glands, and hair follicles 7-9. We have shown that stem cells from the human hair follicle bulge region exhibit multi-potentiality 10.
Stem cells have become a valuable tool in biomedical research, due to their utility as an in vitro system for studying developmental biology, differentiation, tumorigenesis and for their possible therapeutic utility. It is likely that adult epithelial stem cells will be useful in the treatment of diseases such as ectodermal dysplasias, monilethrix, Netherton syndrome, Menkes disease, hereditary epidermolysis bullosa and alopecias 11-13. Additionally, other skin problems such as burn wounds, chronic wounds and ulcers will benefit from stem cell related therapies 14,15. Given the potential for reprogramming of adult cells into a pluripotent state (iPS cells)16,17, the readily accessible and expandable adult stem cells in human skin may provide a valuable source of cells for induction and downstream therapy for a wide range of disease including diabetes and Parkinson's disease.

A rapid, robust way of isolating viable adult epithelial stem cells from human skin is described. The method utilizes enzymatic digestion of skin collagen matrix , followed by plucking of hair follicles and isolation of single cell suspensions or tissue fragments for cell culture.

The homeostasis of all self-renewing tissues is dependent on adult stem cells. As undifferentiated stem cells undergo asymmetric divisions, they generate ...

Isolation and Culture of Individual Myofibers and their Satellite Cells from Adult Skeletal Muscle

Muscle regeneration in the adult is performed by resident stem cells called satellite cells. Satellite cells are defined by their position between the basal lamina and the sarcolemma of each myofiber. Current knowledge of their behavior heavily relies on the use of the single myofiber isolation protocol. In 1985, Bischoff described a protocol to isolate single live fibers from the Flexor Digitorum Brevis (FDB) of adult rats with the goal to create an in vitro system in which the physical association between the myofiber and its stem cells is preserved 1. In 1995, Rosenblattmodified the Bischoff protocol such that myofibers are singly picked and handled separately after collagenase digestion instead of being isolated by gravity sedimentation 2, 3. The Rosenblatt or Bischoff protocol has since been adapted to different muscles, age or conditions 3-6. The single myofiber isolation technique is an indispensable tool due its unique advantages. First, in the single myofiber protocol, satellite cells are maintained beneath the basal lamina. This is a unique feature of the protocol as other techniques such as Fluorescence Activated Cell Sorting require chemical and mechanical tissue dissociation 7. Although the myofiber culture system cannot substitute for in vivo studies, it does offer an excellent platform to address relevant biological properties of muscle stem cells. Single myofibers can be cultured in standard plating conditions or in floating conditions. Satellite cells on floating myofibers are subjected to virtually no other influence than the myofiber environment. Substrate stiffness and coating have been shown to influence satellite cells' ability to regenerate muscles 8, 9 so being able to control each of these factors independently allows discrimination between niche-dependent and -independent responses. Different concentrations of serum have also been shown to have an effect on the transition from quiescence to activation. To preserve the quiescence state of its associated satellite cells, fibers should be kept in low serum medium 1-3. This is particularly useful when studying genes involved in the quiescence state. In serum rich medium, satellite cells quickly activate, proliferate, migrate and differentiate, thus mimicking the in vivo regenerative process 1-3. The system can be used to perform a variety of assays such as the testing of chemical inhibitors; ectopic expression of genes by virus delivery; oligonucleotide based gene knock-down or live imaging. This video article describes the protocol currently used in our laboratory to isolate single myofibers from the Extensor Digitorum Longus (EDL) muscle of adult mice (6-8 weeks old).

Isolation and culture of myofibers is the gold standard in vitro system to study the transition of satellite cells through quiescence, activation and differentiation. Importantly, the single myofiber culture system preserves the myofiber/stem cell association, which is an essential component of the muscle stem cell niche.

Muscle  regeneration in the adult is performed by resident stem cells called satellite  cells. Satellite cells are defined by  their position between the ...

Na&iuml;ve Adult Stem Cells Isolation from Primary Human Fibroblast Cultures

Over the last decade, several adult stem cell populations have been identified in human skin 1-4. The isolation of multipotent adult dermal precursors was first reported by Miller F. D laboratory 5, 6. These early studies described a multipotent precursor cell population from adult mammalian dermis 5. These cells--termed SKPs, for skin-derived precursors-- were isolated and expanded from rodent and human skin and differentiated into both neural and mesodermal progeny, including cell types never found in skin, such as neurons 5. Immunocytochemical studies on cultured SKPs revealed that cells expressed vimentin and nestin, an intermediate filament protein expressed in neural and skeletal muscle precursors, in addition to fibronectin and multipotent stem cell markers 6. Until now, the adult stem cells population SKPs have been isolated from freshly collected mammalian skin biopsies.
Recently, we have established and reported that a population of skin derived precursor cells could remain present in primary fibroblast cultures established from skin biopsies 7. The assumption that a few somatic stem cells might reside in primary fibroblast cultures at early population doublings was based upon the following observations: (1) SKPs and primary fibroblast cultures are derived from the dermis, and therefore a small number of SKP cells could remain present in primary dermal fibroblast cultures and (2) primary fibroblast cultures grown from frozen aliquots that have been subjected to unfavorable temperature during storage or transfer contained a small number of cells that remained viable 7. These rare cells were able to expand and could be passaged several times. This observation suggested that a small number of cells with high proliferation potency and resistance to stress were present in human fibroblast cultures 7.
We took advantage of these findings to establish a protocol for rapid isolation of adult stem cells from primary fibroblast cultures that are readily available from tissue banks around the world (Figure 1). This method has important significance as it allows the isolation of precursor cells when skin samples are not accessible while fibroblast cultures may be available from tissue banks, thus, opening new opportunities to dissect the molecular mechanisms underlying rare genetic diseases as well as modeling diseases in a dish.

Na&#239;ve Adult Stem Cells Isolation from Primary Human Fibroblast Cultures

We report a method to isolate na&#239;ve multipotent skin-derived precursor (SKP) cells from primary human fibroblast cultures. We show that these SKPs derived from fibroblast cultures share similar stem cell properties to the ones derived directly from human skin biopsies. These cells express the neural crest marker, nestin, in addition to the multipotent markers such as OCT4 and Nanog.

Over the last decade, several adult stem cell  populations have been identified in human skin 1-4.  The isolation of multipotent adult dermal precursors ...

Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify and explore the function of these important proteins. However, classical preparations, such as planar bilayers, limit the manipulations and experiments that can be performed on the reconstituted channel and its membrane environment. The more cell-like structure of giant liposomes permits traditional patch-clamp experiments without sacrificing control of the lipid environment.
Electroformation is an efficient mean to produce giant liposomes &gt;10 &mu;m in diameter which relies on the application of alternating voltage to a thin, ordered lipid film deposited on an electrode surface. However, since the classical protocol calls for the lipids to be deposited from organic solvents, it is not compatible with less robust membrane proteins like ion channels and must be modified. Recently, protocols have been developed to electroform giant liposomes from partially dehydrated small liposomes, which we have adapted to protein-containing liposomes in our laboratory.
We present here the background, equipment, techniques, and pitfalls of electroformation of giant liposomes from small liposome dispersions. We begin with the classic protocol, which should be mastered first before attempting the more challenging protocols that follow. We demonstrate the process of controlled partial dehydration of small liposomes using vapor equilibrium with saturated salt solutions. Finally, we demonstrate the process of electroformation itself. We will describe simple, inexpensive equipment that can be made in-house to produce high-quality liposomes, and describe visual inspection of the preparation at each stage to ensure the best results.

Reconstituting functional membrane proteins into giant liposomes of defined composition is a powerful approach when combined with patch-clamp electrophysiology. However, conventional giant liposome production may be incompatible with protein stability. We describe protocols for producing giant liposomes from pure lipids or small liposomes containing ion channels.

The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify ...