Name: primary cultures of rat astrocytes and microglia and their use in the study of amyotrophic lateral sclerosis
Uploaded: 2022-06-23T12:30:00.000+00:00
Duration: 9 min 36 s
Description: Watch this Scientific Journal Video about Primary Cultures of Rat Astrocytes and Microglia and Their Use in the Study of Amyotrophic Lateral Sclerosis at JoVE.com

This work was supported by the Ministry of Education Science and Technological Development Republic of Serbia Contract No. 451-03-9/2021-14/ 200178, the FENS - NENS Education and Training Cluster project &#34;Trilateral Course on Glia in Neuroinflammation&#34;, and the EC H2020 MSCA RISE grant #778405. We thank Marija Ad&#382;i&#263; and Mina Peri&#263; for supplying the immunohistochemistry images and Danijela Batavelji&#263; for help with paper writing.

All experiments were performed in accordance with the EU directives on the protection of animals for scientific purposes and with permission from the Ethical Commission of the Faculty of Biology, University of Belgrade (approval number EK-BF-2016/08). Regarding patient material (sera for IgGs), it was collected for routine clinical examination with informed patient&#39;s consent in accordance with The Code of Ethics of the World Medical Association (Declaration of Helsinki) for experiments involving humans. The protocol was approved by the Ethics committee of the Clinical Center of Serbia (No. 850/6).
1. Primary cell culture preparation
<ol>
	<li>Brain tissue isolation 
	NOTE: Isolation should be performed on ice, using ice cold solutions.
	<ol>
		<li>Use newborn pups 1-3 days old for primary neonatal cell culture33. 
		NOTE: For the study presented here, Sprague Dawley hSOD1G93A and non-transgenic rats were used. For genotyping, rat tails were used for later DNA extraction and PCR.</li>
		<li>Sprinkle the pup&#39;s head with 70% ethanol and immediately decapitate it fast using scissors.</li>
		<li>Cut open the skin using small-angled scissors to expose the skull. Open the skull by making a cut from the foramen magnum toward the orbits. Next, make a perpendicular midline cut. Remove the brain from the skull and place it in a Petri dish containing phosphate-buffered saline (PBS). 
		NOTE: Clean the tools between pups to prevent cross-contamination between transgenic and non-transgenic pups. Isolation of the cortices from the rest of the brain should be done under a stereomicroscope.</li>
		<li>Using the tip of the forceps, tear the connections between both hemispheres. Then, separate the hemispheres with curved forceps by gently pushing a hemisphere from the center to the side.</li>
		<li>Remove the meninges by carefully tearing them with straight and curved forceps.</li>
		<li>Remove the hippocampus by pinching it with a curved forceps. Discard the hippocampus or use it for another cell culture preparation. 
		NOTE: Further steps should be performed under the laminar flow hood to ensure sterile conditions.</li>
	</ol>
	</li>
	<li>Tissue homogenization and dissociation
	<ol>
		<li>Transfer one cortex into a 15 mL tube filled with 3 mL of cold PBS. Pipet the suspension up and down with a 1 mL tip, completing 10-15 strokes until the suspension becomes homogeneous. 
		NOTE: Be very cautious not to produce bubbles in this process.</li>
		<li>Centrifuge at 500 &#215; g for 5 min. Remove the supernatant and resuspend the pellet in 3 mL of cold PBS by pipetting it up and down with a 1 mL tip. Repeat the centrifugation step. 
		NOTE: All solutions used from this point should be prewarmed to 37 &#176;C.</li>
		<li>Discard the supernatant and resuspend the pellet in 2 mL of complete Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and antibiotics (penicillin and streptomycin). Transfer the homogenate to a 2 mL tube.</li>
		<li>Pass the homogenate through 21 G and 23 G needles (three times each) to make a suspension of single cells. 
		NOTE: Be very cautious not to produce bubbles in this process.</li>
	</ol>
	</li>
	<li>Pour the cell suspension prepared from one cortex into a 60 mm diameter Petri dish or a T25 flask (with the surface pretreated with a polypeptide coating for the growth of adherent cells) containing 3 mL of complete DMEM. Lightly shake the Petri dish so that the suspension is uniformly distributed.</li>
	<li>Grow the cells in the incubator in a humidified atmosphere of 5% CO2/95% air at 37 &#176;C.</li>
	<li>Change the medium 48 h after isolation and replace the medium every 3 days.</li>
	<li>To promote astrocyte growth, when cells reach 70%-80% confluency, wash them with prewarmed PBS to remove the loosely attached glial cells and traces of FBS in the medium.
	<ol>
		<li>Trypsinize the underlying layer of astrocytes by adding 1 mL of prewarmed trypsin solution (0.25% trypsin, 0.02% EDTA in PBS, sterile-filtered).</li>
		<li>Place the dish in the incubator at 37 &#176;C for 2-5 min.</li>
		<li>Check the cells under the microscope. When they start to detach, add 4 mL of complete DMEM.</li>
		<li>Collect the cell suspension and transfer it to a 15 mL tube. Centrifuge at 500 &#215; g for 5 min.</li>
		<li>Discard the supernatant and resuspend the pellet in 1 mL of complete medium. Count the cells using a hemocytometer.</li>
		<li>Replate the cells at a density of 104 cells/cm2 in 5 mL of fresh complete DMEM.</li>
	</ol>
	</li>
	<li>Change the medium every 3 days. To minimize the presence of other glial cell types, after astrocytes reach 50% confluence and prior to each media replacement, wash the cells with complete DMEM.
	<ol>
		<li>Aspirate the supernatant medium with a 1 mL pipette and gently dispense it onto the layer of cells several times. Ensure that the whole surface of the cell layer is covered during this washing step.</li>
	</ol>
	</li>
	<li>Once the cells reach 80% confluence (usually after 14 days), repeat the trypsinization and the collection of astrocytes as described in step 1.6.</li>
	<li>Seed 5 &#215; 103 of astrocytes on a 7 mm circular glass coverslip coated with poly-L-lysine (50 &#181;g/mL). Use them in experiments after 48 h.</li>
	<li>To promote microglia, after step 1.7, allow the glial cells to reach a confluent layer34. When microglial cells appear on top of the astrocyte layer (after 10-15 days, recognized by their smaller, more oval bodies and shorter processes), shake the Petri dish on an orbital shaker for 2 h at 220 rpm.</li>
	<li>Dispersing and seeding microglial cells
	<ol>
		<li>Lightly wash the detached and loosely attached cells by aspirating the supernatant medium with a 1 mL pipette tip and gently dispense it onto the layer of cells several times. Be sure to cover the whole surface of the cell layer during this washing step, collect the medium with the detached cells, and transfer it to a 15 mL tube.</li>
		<li>Centrifuge at 500 &#215; g for 5 min. Discard the supernatant and resuspend the pellet in 1 mL of medium.</li>
		<li>Pass the cell suspension through a 21 G needle to obtain a single-cell suspension. 
		NOTE: Most published methods use trypsin or papain to dissociate the tissue; however, 21 G needles have the advantage of preventing the overdigestion of cells, allowing for a gentler dissociation.</li>
		<li>Count the cells using a hemocytometer. Seed 5 &#215; 103 microglial cells on a 7 mm circular glass coverslip coated with poly-L-lysine (50 &#181;g/mL). Use them in experiments after 48 h. 
		&#8203;NOTE: It is difficult to obtain a pure microglial culture by shaking, since the oligodendrocyte precursor cells and astrocytes may still be present in a variable number depending on the experimenter&#39;s experience. The immunocytochemical protocol used to estimate the presence of different cell populations in glial cultures has been described elsewhere35.</li>
	</ol>
	</li>
</ol>
2. Immunocytochemistry
<ol>
	<li>Rinse the cells plated on coverslips in PBS, 2 x 1 min.</li>
	<li>Fix the cells in 4% paraformaldehyde for 20 min at room temperature (RT).</li>
	<li>Wash in PBS 3 x 5 min.</li>
	<li>Incubate in a blocking solution containing 10% normal donkey serum (NDS)/1% bovine serum albumin (BSA)/0.1% Triton X-100 for 45 min at RT.
	<ol>
		<li>Add 50 &#181;L of a blocking solution per 10 mm diameter coverslip.</li>
	</ol>
	</li>
	<li>Prepare primary antibodies in 1% NDS/1% BSA/0.1% TritonX-100 in the following dilutions: mouse anti-GFAP 1:300, goat anti-Iba1 1:500. Incubate the cells with primary antibodies overnight at +4 &#176;C.</li>
	<li>Rinse in PBS, 3 x 10 min.</li>
	<li>Incubate with the secondary antibodies conjugated with a fluorophore: donkey anti-mouse (excitation 488 nm [1:200)) or donkey anti-goat (excited at 647 nm [1:200)). Incubate the cells for 2 h at RT in the dark.</li>
	<li>Wash in PBS, 7 x 5 min.</li>
	<li>Incubate the cells with 4&#39;,6-diamidino-2-phenylindole (DAPI, 1:4,000) for 10 min at RT.</li>
	<li>Wash in PBS 5 x 5 min.</li>
	<li>Mount the coverslips on microscope slides using a mounting solution; use one drop of it per coverslip. 
	NOTE: Antibody dilutions may vary depending on the producer. Users must determine optimal dilutions for their experiments.</li>
</ol>
3. Time-lapse video imaging
NOTE: Solutions containing the fluorescent dye should be protected from direct light. Before starting the imaging experiment, make sure that the glass coverslip does not move when turning on the perfusion.
<ol>
	<li>Preparation of solutions and cells
	<ol>
		<li>Prepare extracellular solution (ECS) for imaging: 140 mM NaCl, 5 mM KCl, 2 mM CaCal2, 2 mM MgCl2, 10 mM D-glucose, and 10 mM HEPES. Adjust the pH to 7.4. Ensure that the osmolarity is 280-300 mOsm/kg and prepare ECS without Ca2+: 140 mM NaCl, 5 mM KCl, 2 mM MgCl2, 10 mM D-glucose, 10 mM HEPES, and 0.1 mM EGTA.</li>
		<li>Prepare the test solutions: 100 &#181;M ATP dissolved in ECS, 1 &#181;M Thg in ECS without Ca2+, and 0.1 mg/mL ALS IgG in ECS8.</li>
		<li>Transfer one coverslip to a dish with ECS to wash out the complete DMEM. Prepare the dye loading solution by diluting 1 mM stock solution of Fluo-4 AM with ECS to the final concentration of 5 &#181;M Fluo-4 AM.
		<ol>
			<li>Place the coverslip with cells in the dye loading solution for 30 min at RT in the dark. Wash the cells in ECS for 20 min.</li>
		</ol>
		</li>
		<li>If cells are not adequately loaded (i.e., if the basal level of fluorescence is too low-&#60;5% of the dynamic range of the camera with exposure time greater than 400 ms), increase the loading time to up to 45 min to 1 h at 37 &#176;C. Alternatively, mix the Fluo-4 AM stock solution with an equal volume of 20% (w/v) detergent (Pluronic) in DMSO before diluting in ECS, making the final Pluronic concentration ~0.02%. 
		NOTE: The experimental examples of this study were performed with human IgG isolated from ALS patients&#39; blood sera as described previously4,5,11. Each experiment was performed with IgG samples of a single patient.</li>
	</ol>
	</li>
	<li>Video imaging 
	NOTE:&#160;In this protocol, the Video Imaging System was combined with a xenon short arc lamp, a polychromator system, and an inverted epifluorescence microscope equipped with water, glycerin, and oil immersion objective. Timelapse imaging was performed using a Digital Camera System.
	<ol>
		<li>Switch on the components of the imaging setup 15 min before the experiment to allow the system to reach the working temperature.</li>
		<li>Place the coverslip in the recording chamber with 1 mL of working solution (ECS or ECS without Ca2+).</li>
		<li>To choose the correct imaging protocol, open the imaging software and in the Acquisition panel, choose the filter pairs for Fluo4-AM with an excitation wavelength at 480 nm, a dichroic mirror at 505 nm, and an emission wavelength at 535 nm.</li>
		<li>Choose the field of view taking care to have a consistent number of cells throughout the experiment. Adjust the exposure time and detector gain appropriately, in such a manner that the signal is not saturated. Choose the pseudocolor mode for acquisition. For more detailed instructions, refer to the manual provided by the manufacturer.</li>
		<li>Adjust the sampling rate to 1 Hz (1 frame per second).</li>
		<li>Start the imaging by acquiring the basal level of fluorescence for 3-5 min for the baseline determination (F0). Ensure a constant flow of the working solution.</li>
		<li>Stop the flow of the working solution and switch to the test solution for the desired length of time (also see time bars in examples of Figure 2 and Figure 3). Between each test solution, wash the cells with a constant flow of the working solution for 3-5 min.</li>
		<li>Keep the solution volume in the recording chamber at ~1 mL by arranging a constant suction from the top of the solution (see Figure 2E). 
		&#8203;NOTE: To apply the treatment directly to the imaged cells, a customized delivery system made of a glass pipette (0.8 mm inner diameter) was positioned ~350 &#181;m away and ~1 mm above the cells, at an angle of 45&#176; combined with the solution exchange system containing pinch valves and an electronic valve controller (see Figure 2E).</li>
	</ol>
	</li>
</ol>
4. Data analysis
<ol>
	<li>Define a region of interest (ROI) that corresponds to the individual cell.
	<ol>
		<li>Choose the frame with the highest signal intensity (i.e., from the application of ATP) and encircle one cell by using the application Polygonal Tool. Repeat for all the cells in the acquired field. Choose five ROIs in the background using the Circle Tool.</li>
		<li>To measure the mean signal intensity of a single cell and the background for each time frame, select all ROIs and use the Multi Measure command in ROI Manager in ImageJ or any equivalent command in commercial software (see the Table of Materials).</li>
	</ol>
	</li>
	<li>Export the mean signal intensity values of the ROIs as a spreadsheet.</li>
	<li>Average five background ROIs and subtract the averaged background of each frame from the mean ROI intensity of the frame acquired at the same time.</li>
	<li>To normalize the obtained data to the baseline signal, use equation (1): 
	&#916;F/F0 = (F - F0)/F0&#160; &#160;(1) 
	where F is the signal intensity of each time frame after background subtraction, and F0 is the baseline Fluo-4 fluorescence. 
	NOTE: A custom-made code was used in this study.</li>
	<li>To analyze calcium activity, determine several parameters4: the amplitude of the calcium peak (&#916;F/F0), the integrated change (time integral, surface under the response recording) of the calcium transient (&#916;F/F0 &#215; s), time-to-peak-elapsed time from the onset of stimulation to the maximum of calcium transient (s), rise time-time elapsed from the onset of the stimulation to the value of &#916;F/F0 that reaches 50%-80% of the maximal amplitude (s), half-width-full width of a transient at half-maximal amplitude (s), frequency of responses if they are repetitive (Hz).</li>
</ol>

The authors have no conflicts of interest to declare.

This paper presents the method of primary cell culturing as a fast and &#34;on the budget&#34; tool for studying different aspects of cell (patho)physiology such as ALS in the rat hSOD1G93A model. The technique is thus suitable for studies at the single-cell level that can be extrapolated and further investigated at a higher level of organization (i.e., in tissue slices or in a live animal). Cell culturing as a technique, however, has a few caveats. It is most critical to do the brain tissue isolation and the ...

Cell culture techniques have given rise to numerous advancements in diverse fields of cellular neurophysiology in health and disease. Particularly, primary cell cultures, freshly isolated from the neuronal tissue of a lab animal, allow the experimenter to closely study the behavior of diverse cells in different biochemical media and physiological setups. Using different fluorescent physiological indicators such as the Ca2+-sensitive dyes in combination with time-lapse video microscopy provides better insight into the cellular biophysical and biochemical processes in real time.
ALS is a devastating neurodegenerative disease that affects upper and lower motor neurons1. The disease has a complex pathogenesis of the familial type but mostly of the sporadic form (90% of cases)2. It is well known that non-cell autonomous mechanisms contribute to ALS pathophysiology, primarily due to the essential role of glial cells3. ALS is also well characterized as a neuroinflammatory disease with involvement of humoral and cellular factors of inflammation.
Immunoglobulin G is widely used as a molecular marker in ALS and other neurodegenerative diseases. Studying the serum level of this marker can indicate the presence and stage of neuroinflammation in the disease4,5,6, while its presence in the cerebrospinal fluid can indicate a breach of the blood brain barrier7. IgGs were also identified as deposits in the spinal cord motor neurons of ALS patients7. Nevertheless, this approach has shown some inconsistencies in the correlation of the level of IgGs with the stage and characteristics of the disease6.
IgG isolated from the sera of ALS patients (ALS IgG) can induce a calcium response in naive astrocytes8 and glutamate release in neurons, pointing to an excitotoxic effect-a hallmark of ALS pathology9. However, studies on the hSOD1G93A ALS rat model (containing multiple copies of the human SOD1 mutation10) showed a number of markers of oxidative stress in cultured neuroglial cells11, tissues12,13,14, or live animals13. It is noteworthy that the astrocytes cultured from the ALS rat model were more prone to oxidative stress induced by peroxide than the astroglia from non-transgenic littermates11.
Microglial cells in culture are affected by ALS IgG in a less apparent way. Namely, a BV-2 microglial cell line displayed a rise in the signal from fluorescent markers of oxidative stress in response to the application of only 4/11 ALS IgG patient samples15. It is well known that microglia participate in many neuroinflammatory pathologies, adding to oxidative stress and late progression phase in the non-cell autonomous mechanism of ALS16,17. Nevertheless, the data with ALS IgGs indicated that these cells may not be as reactive as astrocytes to these humoral factors of ALS inflammation. Several studies have been conducted with primary astrocytes from ALS murine models, not only in pups but also in symptomatic animals, either on the brain or on the spinal cord18,19,20,21. This is also true for microglial primary cultures, although to a lesser extent than astrocytes and mostly from brain regions at the embryonic stage22,23,24.
We use time-lapse video imaging of Ca2+ on cells in culture primarily as a means to follow intracellular transients of this ion as a physiological marker of excitotoxicity. Thus, by biophysical characterization of these transients (amplitude, area under transient, rise-time, frequency) the researcher can obtain experimental diagnostic parameters from diverse cellular models of neurodegeneration. This technique thus offers an advantage of a quantitative physiological assessment of IgGs as disease biomarkers. There is a large body of literature on the role of IgGs and Ca2+ in the induction of ALS. Most of these studies were performed by inducing ALS by injecting patient IgGs into experimental animals25,26,27,28,29, which then showed intracellular Ca2+ elevation and IgG depositions. A line of studies explored the effect of ALS IgGs on the motor synapse in vitro30,31,32. In the above context, the technique presented here puts the focus on the glial cells as important players in the non-cell autonomous mechanism of ALS and quantifies their potential excitotoxic response to IgGs as humoral factors of neuroinflammation. This approach may have a wider application in testing other humoral factors such as whole sera, CSF, or cytokines in different cell culture systems and in cellular models of general inflammation.
This paper describes how to prepare primary cultures of glial cells, astrocytes, and microglia from the cortices of Sprague Dawley rats and how to further use these cells to study ALS pathophysiology with patient sera-derived IgG. Protocols are detailed for the dye-loading of cultured cells (Figure 1) and the recordings of Ca2+ changes in time-lapse video imaging experiments. Examples of video recordings will show how glial cells react to ALS IgG as compared to ATP, the latter activating purinergic membrane receptors. Shown for the first time is an example on how astrocytes isolated from the hSOD1G93A ALS rat brain react with a more pronounced Ca2+ response to ALS IgG compared to non-transgenic controls and how to relate this process to the differences in Ca2+ store operation. Also shown is an example of calcium imaging in microglial cells acutely challenged with ALS IgG, with only a modest response of intracellular calcium.

<table><tbody><tr><td>15 mL tube</td><td>Sarstedt, Germany</td><td>62 554 502</td><td></td></tr><tr><td>2 mL tube</td><td>Sarstedt, Germany</td><td>72.691</td><td></td></tr><tr><td>21 G needle</td><td>Nipro, Japan</td><td>HN-2138-ET</td><td></td></tr><tr><td>23 G needle</td><td>Nipro, Japan</td><td>HN-2338-ET</td><td></td></tr><tr><td>5 mL syringe</td><td>Nipro, Japan</td><td>SY3-5SC-EC</td><td></td></tr><tr><td>6 mm circular glass coverslip</td><td>Menzel Glasser, Germany</td><td>630-2113</td><td></td></tr><tr><td>60 mm Petri dish</td><td>ThermoFisher Sientific, USA</td><td>130181</td><td></td></tr><tr><td>ATP</td><td>Sigma-Aldrich, Germany</td><td>A9062</td><td></td></tr><tr><td>AxioObserver A1</td><td>Carl Zeiss, Germany</td><td></td><td></td></tr><tr><td>Bovine serum albumine</td><td>Sigma-Aldrich, Germany</td><td>B6917</td><td></td></tr><tr><td>Calcium chloride</td><td>Sigma-Aldrich, Germany</td><td>2110</td><td></td></tr><tr><td>Centrifuge</td><td>Eppendorf, Germany</td><td></td><td></td></tr><tr><td>DAPI</td><td>Sigma-Aldrich, Germany</td><td>10236276001</td><td></td></tr><tr><td>D-glucose</td><td>Sigma-Aldrich, Germany</td><td>158968</td><td></td></tr><tr><td>DMEM</td><td>Sigma-Aldrich, Germany</td><td>D5648</td><td></td></tr><tr><td>Donkey-anti goat AlexaFluor 647 IgG antibody</td><td>Invitrogen, USA</td><td>A-21447</td><td></td></tr><tr><td>Donkey-anti mouse AlexaFluor 488 IgG antibody</td><td>Invitrogen, USA</td><td>A-21202</td><td></td></tr><tr><td>EDTA</td><td>Sigma-Aldrich, Germany</td><td>EDS-100G</td><td></td></tr><tr><td>EGTA</td><td>Sigma-Aldrich, Germany</td><td>E4378</td><td></td></tr><tr><td>&amp;rdquo;evolve&amp;rdquo;-EM 512 Digital Camera System</td><td>Photometrics, USA</td><td></td><td></td></tr><tr><td>Fetal bovine serum (FBS)</td><td>Gibco, ThermoFisher Scientific, USA</td><td>10500064</td><td></td></tr><tr><td>Fiji ImageJ Software</td><td>Open source under the GNU General Public Licence</td><td></td><td></td></tr><tr><td>FITC filter set</td><td>Chroma Technology Inc., USA</td><td></td><td></td></tr><tr><td>Fluo-4 AM</td><td>Molecular Probes, USA</td><td>F14201</td><td></td></tr><tr><td>Goat anti-Iba1</td><td>Fujifilm Wako Chemicals, USA</td><td>011-27991</td><td></td></tr><tr><td>HEPES</td><td>Biowest, France</td><td>P5455</td><td></td></tr><tr><td>HighSpeed Solution Exchange System</td><td>ALA Scientific Instruments, USA</td><td></td><td></td></tr><tr><td>Incubator</td><td>Memmert GmbH + Co. KG, Germany</td><td></td><td></td></tr><tr><td>Magnesium chloride</td><td>Sigma-Aldrich, Germany</td><td>M2393</td><td></td></tr><tr><td>Matlab software</td><td>Math Works, USA</td><td></td><td></td></tr><tr><td>Mouse anti-GFAP</td><td>Merck Millipore, USA</td><td>MAB360</td><td></td></tr><tr><td>Mowiol 40-88</td><td>Sigma-Aldrich, Germany</td><td>324590</td><td></td></tr><tr><td>Normal donkey serum</td><td>Sigma-Aldrich, Germany</td><td>D9663</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Sigma-Aldrich, Germany</td><td>158127</td><td></td></tr><tr><td>Penicilin and Streptomycin</td><td>ThermoFisher Sientific, USA</td><td>15140122</td><td></td></tr><tr><td>Poly-L-lysine</td><td>Sigma-Aldrich, Germany</td><td>P5899</td><td></td></tr><tr><td>Potassium chloride</td><td>Sigma-Aldrich, Germany</td><td>P5405</td><td></td></tr><tr><td>Potassium dihydrogen phosphate</td><td>Carlo Erba Reagents, Spain</td><td>471686</td><td></td></tr><tr><td>Shaker DELFIA PlateShake</td><td>PerkinElmer Life Sciencies, USA</td><td></td><td></td></tr><tr><td>Sodium bicarbonate</td><td>Sigma-Aldrich, Germany</td><td>S3817</td><td></td></tr><tr><td>Sodium chloride</td><td>Sigma-Aldrich, Germany</td><td>S5886</td><td></td></tr><tr><td>Sodium phosphate dibasic heptahydrate</td><td>Carl ROTH GmbH</td><td>X987.2</td><td></td></tr><tr><td>Sodium pyruvate</td><td>Sigma-Aldrich, Germany</td><td>P5280</td><td></td></tr><tr><td>Thapsigargine</td><td>Tocris Bioscience, UK</td><td>1138</td><td></td></tr><tr><td>Triton X - 100</td><td>Sigma-Aldrich, Germany</td><td>T8787</td><td></td></tr><tr><td>Trypsin</td><td>Sigma-Aldrich, Germany</td><td>T4799</td><td></td></tr><tr><td>Vapro Vapor Pressure Osmometer 5520</td><td>Wescor, ELITechGroup Inc., USA</td><td></td><td></td></tr><tr><td>ViiFluor Imaging System</td><td>Visitron System Gmbh, Germany</td><td></td><td></td></tr><tr><td>VisiChrome Polychromator System</td><td>Visitron System Gmbh, Germany</td><td></td><td></td></tr><tr><td>VisiView high performance setup</td><td>Visitron System Gmbh, Germany</td><td></td><td></td></tr><tr><td>Xenon Short Arc lamp</td><td>Ushio, Japan</td><td></td><td></td></tr></tbody></table>

primary cultures of rat astrocytes and microglia and their use in the study of amyotrophic lateral sclerosis

This protocol demonstrates how to prepare primary cultures of glial cells, astrocytes, and microglia from the cortices of Sprague Dawley rats and how to use these cells for the purpose of studying the pathophysiology of amyotrophic lateral sclerosis (ALS) in the rat hSOD1G93A model. First, the protocol shows how to isolate and culture astrocytes and microglia from postnatal rat cortices, and then how to characterize and test these cultures for purity by immunocytochemistry using the glial fibrillary acidic protein (GFAP) marker of astrocytes and the ionized calcium-binding adaptor molecule 1 (Iba1) microglial marker. In the next stage, methods are described for dye-loading (calcium-sensitive Fluo 4-AM) of cultured cells and the recordings of Ca2+ changes in video imaging experiments on live cells.
The examples of video recordings consist of: (1) cases of Ca2+ imaging of cultured astrocytes acutely exposed to immunoglobulin G (IgG) isolated from ALS patients, showing a characteristic and specific response compared to the response to ATP as demonstrated in the same experiment. Examples also show a more pronounced transient rise in intracellular calcium concentration evoked by ALS IgG in hSOD1G93A astrocytes compared to non-transgenic controls; (2) Ca2+ imaging of cultured astrocytes during a depletion of calcium stores by thapsigargin (Thg), a non-competitive inhibitor of the endoplasmic reticulum Ca2+ ATPase, followed by store-operated calcium entry elicited by the addition of calcium in the recording solution, which demonstrates the difference between Ca2+ store operation in hSOD1G93A and in non-transgenic astrocytes; (3) Ca2+ imaging of the cultured microglia showing predominantly a lack of response to ALS IgG, whereas ATP application elicited a Ca2+ change. This paper also emphasizes possible caveats and cautions regarding critical cell density and purity of cultures, choosing the correct concentration of the Ca2+ dye and dye-loading techniques.

Characterization of different glial cell types in culture 
It usually takes 15-21 days to produce astrocytes for experiments, while microglial cells take 10-15 days to grow. Immunostaining was performed to assess the cell purity of the culture. Figure 1 shows the expression of double labeling of the astrocytic marker GFAP and the microglial marker Iba1 in respective cultures.
Calcium imaging is known to reveal the differences in cell physiolo...

Watch this Scientific Journal Video about Primary Cultures of Rat Astrocytes and Microglia and Their Use in the Study of Amyotrophic Lateral Sclerosis at JoVE.com

Primary Cultures of Rat Astrocytes and Microglia and Their Use in the Study of Amyotrophic Lateral Sclerosis

We present here a protocol on how to prepare primary cultures of glial cells, astrocytes, and microglia from rat cortices for time-lapse video imaging of intracellular Ca2+ for research on pathophysiology of amyotrophic lateral sclerosis in the hSOD1G93A rat model.

This protocol demonstrates how to prepare primary cultures of glial cells, astrocytes, and microglia from the cortices of Sprague Dawley rats and how to ...

primary-cultures-rat-astrocytes-microglia-their-use-study-amyotrophic

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

3.3K Views.  University of Belgrade Faculty of Biology. We present here a protocol on how to prepare primary cultures of glial cells, astrocytes, and microglia from rat cortices for time-lapse video imaging of intracellular Ca2+ for research on pathophysiology of amyotrophic lateral sclerosis in the hSOD1G93A rat model.

Video: Primary Cultures of Rat Astrocytes and Microglia and Their Use in the Study of Amyotrophic Lateral Sclerosis

Primary Microglia Isolation from Mixed Glial Cell Cultures of Neonatal Rat Brain Tissue

Microglia account for approximately 12% of the total cellular population in the mammalian brain. While neurons and astrocytes are considered the major cell types of the nervous system, microglia play a significant role in normal brain physiology by monitoring tissue for debris and pathogens and maintaining homeostasis in the parenchyma via phagocytic activity 1,2. Microglia are activated during a number of injury and disease conditions, including neurodegenerative disease, traumatic brain injury, and nervous system infection 3. Under these activating conditions, microglia increase their phagocytic activity, undergo morpohological and proliferative change, and actively secrete reactive oxygen and nitrogen species, pro-inflammatory chemokines and cytokines, often activating a paracrine or autocrine loop 4-6. As these microglial responses contribute to disease pathogenesis in neurological conditions, research focused on microglia is warranted.
	Due to the cellular heterogeneity of the brain, it is technically difficult to obtain sufficient microglial sample material with high purity during in vivo experiments. Current research on the neuroprotective and neurotoxic functions of microglia require a routine technical method to consistently generate pure and healthy microglia with sufficient yield for study. We present, in text and video, a protocol to isolate pure primary microglia from mixed glia cultures for a variety of downstream applications. Briefly, this technique utilizes dissociated brain tissue from neonatal rat pups to produce mixed glial cell cultures. After the mixed glial cultures reach confluency, primary microglia are mechanically isolated from the culture by a brief duration of shaking. The microglia are then plated at high purity for experimental study.
	The principle and protocol of this methodology have been described in the literature 7,8. Additionally, alternate methodologies to isolate primary microglia are well described 9-12. Homogenized brain tissue may be separated by density gradient centrifugation to yield primary microglia 12. However, the centrifugation is of moderate length (45 min) and may cause cellular damage and activation, as well as, cause enriched microglia and other cellular populations. Another protocol has been utilized to isolate primary microglia in a variety of organisms by prolonged (16 hr) shaking while in culture 9-11. After shaking, the media supernatant is centrifuged to isolate microglia. This longer two-step isolation method may also perturb microglial function and activation. We chiefly utilize the following microglia isolation protocol in our laboratory for a number of reasons: (1) primary microglia simulate in vivo biology more faithfully than immortalized rodent microglia cell lines, (2) nominal mechanical disruption minimizes potential cellular dysfunction or activation, and (3) sufficient yield can be obtained without passage of the mixed glial cell cultures.
	It is important to note that this protocol uses brain tissue from neonatal rat pups to isolate microglia and that using older rats to isolate microglia can significantly impact the yield, activation status, and functional properties of isolated microglia. There is evidence that aging is linked with microglia dysfunction, increased neuroinflammation and neurodegenerative pathologies, so previous studies have used ex vivo adult microglia to better understand the role of microglia in neurodegenerative diseases where aging is important parameter. However, ex vivo microglia cannot be kept in culture for prolonged periods of time. Therefore, while this protocol extends the life of primary microglia in culture, it should be noted that the microglia behave differently from adult microglia and in vitro studies should be carefully considered when translated to an in vivo setting.

Isolating primary microglia from the cellular heterogeneity of the brain is essential to investigate their role in both physiological and pathological conditions. This protocol describes a mechanical isolation and mixed cell culture technique that provides high yield and high purity, viable primary microglial cells for in vitro study and downstream applications.

Microglia account for approximately 12% of the  total cellular population in the mammalian brain. While neurons and astrocytes are considered  the major ...

Immunology and Infection

Bilaminar Co-culture of Primary Rat Cortical Neurons and Glia

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or co-culture model. This system is suitable for a variety of experimental needs requiring either a glass or plastic growth substrate and can also be used for culture of other types of neurons.
Rat cortical neurons obtained from the late embryonic stage (E17) are plated on glass coverslips or tissue culture dishes facing a feeder layer of glia grown on dishes or plastic coverslips (known as Thermanox), respectively. The choice between the two configurations depends on the specific experimental technique used, which may require, or not, that neurons are grown on glass (e.g. calcium imaging versus Western blot). The glial feeder layer, an astroglia-enriched secondary culture of mixed glia, is separately prepared from the cortices of newborn rat pups (P2-4) prior to the neuronal dissection.
A major advantage of this culture system as compared to a culture of neurons only is the support of neuronal growth, survival, and differentiation provided by trophic factors secreted from the glial feeder layer, which more accurately resembles the brain environment in vivo. Furthermore, the co-culture can be used to study neuronal-glial interactions1.
At the same time, glia contamination in the neuronal layer is prevented by different means (low density culture, addition of mitotic inhibitors, lack of serum and use of optimized culture medium) leading to a virtually pure neuronal layer, comparable to other established methods1-3. Neurons can be easily separated from the glial layer at any time during culture and used for different experimental applications ranging from electrophysiology4, cellular and molecular biology5-8, biochemistry5, imaging and microscopy4,6,7,9,10. The primary neurons extend axons and dendrites to form functional synapses11, a process which is not observed in neuronal cell lines, although some cell lines do extend processes.
A detailed protocol of culturing rat hippocampal neurons using this co-culture system has been described previously4,12,13. Here we detail a modified protocol suited for cortical neurons. As approximately 20x106 cells are recovered from each rat embryo, this method is particularly useful for experiments requiring large numbers of neurons (but not concerned about a highly homogenous neuronal population). The preparation of neurons and glia needs to be planned in a time-specific manner. We will provide the step-by-step protocol for culturing rat cortical neurons as well as culturing glial cells to support the neurons.

Here we provide a protocol for culturing rat cortical neurons in the presence of a glial feeder layer. The cultured neurons establish polarity and create synapses, and can be separated from the glia for use in various applications, such as electrophysiology, calcium imaging, cell survival assays, immunocytochemistry, and RNA/DNA/protein isolation.

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or ...

Isolation and Culture of Mouse Cortical Astrocytes

Astrocytes are an abundant cell type in the mammalian brain, yet much remains to be learned about their molecular and functional characteristics. In vitro astrocyte cell culture systems can be used to study the biological functions of these glial cells in detail. This video protocol shows how to obtain pure astrocytes by isolation and culture of mixed cortical cells of mouse pups. The method is based on the absence of viable neurons and the separation of astrocytes, oligodendrocytes and microglia, the three main glial cell populations of the central nervous system, in culture. Representative images during the first days of culture demonstrate the presence of a mixed cell population and indicate the timepoint, when astrocytes become confluent and should be separated from microglia and oligodendrocytes. Moreover, we demonstrate purity and astrocytic morphology of cultured astrocytes using immunocytochemical stainings for well established and newly described astrocyte markers. This culture system can be easily used to obtain pure mouse astrocytes and astrocyte-conditioned medium for studying various aspects of astrocyte biology.

Astrocytes have been recognized to be versatile cells participating in fundamental biological processes that are essential for normal brain development and function, and central nervous system repair. Here we present a rapid procedure to obtain pure mouse astrocyte cultures to study the biology of this major class of central nervous system cells.

Astrocytes  are an abundant cell type in the mammalian brain, yet much remains to be  learned about their molecular and functional characteristics. In ...

Rapid Genotyping of Animals Followed by Establishing Primary Cultures of Brain Neurons

High-resolution analysis of the morphology and function of mammalian neurons often requires the genotyping of individual animals followed by the analysis of primary cultures of neurons. We describe a set of procedures for: labeling newborn mice to be genotyped, rapid genotyping, and establishing low-density cultures of brain neurons from these mice. Individual mice are labeled by tattooing, which allows for long-term identification lasting into adulthood. Genotyping by the described protocol is fast and efficient, and allows for automated extraction of nucleic acid with good reliability. This is useful under circumstances where sufficient time for conventional genotyping is not available, e.g., in mice that suffer from neonatal lethality. Primary neuronal cultures are generated at low density, which enables imaging experiments at high spatial resolution. This culture method requires the preparation of glial feeder layers prior to neuronal plating. The protocol is applied in its entirety to a mouse model of the movement disorder DYT1 dystonia (&#916;E-torsinA knock-in mice), and neuronal cultures are prepared from the hippocampus, cerebral cortex and striatum of these mice. This protocol can be applied to mice with other genetic mutations, as well as to animals of other species. Furthermore, individual components of the protocol can be used for isolated sub-projects. Thus this protocol will have wide applications, not only in neuroscience but also in other fields of biological and medical sciences.

We describe procedures for labeling and genotyping newborn mice and generating primary neuronal cultures from them. The genotyping is rapid, efficient and reliable, and allows for automated nucleic-acid extraction. This is especially useful for neonatally lethal mice and their cultures that require prior completion of genotyping.

High-resolution analysis of the morphology and function of mammalian neurons often requires the genotyping of individual animals followed by the analysis ...

Preparation of Primary Mixed Glial Cultures from Adult Mouse Spinal Cord Tissue

It has been well-accepted that spinal cord glial responses contribute significantly to the development of neuropathic pain. Tremendous information regarding glial activities at the cellular and molecular levels has been obtained through in vitro cell culture systems. The in vitro systems utilized, mainly include primary glia derived from neonatal brain cortical tissue and immortalized cell lines. However, these systems may not reflect the characteristics of spinal cord glial cells in vivo. In order to further investigate the roles of spinal cord glial cells in the development of peripheral nerve injury-induced neuropathic pain using a culture system that better reflects the in vivo condition, our laboratory has developed a method to establish primary spinal cord mixed glial cultures from adult mice. Briefly, spinal cords are collected from adult mice and processed through papain digestion followed by myelin removal with a density-gradient medium. Single cell suspensions are cultured in complete Dulbecco&#39;s modified Eagle media (cDMEM) supplemented with 2-mercaptoethanol (2-ME) at 35.9 oC. These culture conditions were optimized specifically for the growth of mixed glial cells. Under these conditions, cells are ready to be used for experimentation between 12 - 14 d&#160;(cells are usually in log phase during this time) after the establishment of the culture (D 0) and can be kept in culture conditions up to D 21. This culture system can be used to investigate the responses of spinal cord glial cells upon stimulation with various substances and agents. Besides neuropathic pain, this system can be used to study glial responses in other diseases that involve pathological changes of spinal cord glial cells.

The development of neuropathic pain involves pathological changes of spinal cord glial cells. A reliable glial culture system derived from adult spinal cord tissue and designed for studying these cells in vitro is lacking. Therefore, we show here how&#160;to establish primary mixed glial cultures from adult mouse spinal cord tissue.

It has been well-accepted that spinal cord glial responses contribute significantly to the development of neuropathic pain. Tremendous information ...

Concomitant Isolation of Primary Astrocytes and Microglia for Protozoa Parasite Infection

Astrocytes and microglia are the most abundant glial cells. They are responsible for physiological support and homeostasis maintenance in the central nervous system (CNS). The increasing evidences of their involvement in the control of infectious diseases justify the emerging interest in the improvement of methodologies to isolate primary astrocytes and microglia in order to evaluate their responses to infections that affect the CNS. Considering the impact of Trypanosoma cruzi (T. cruzi) and Toxoplasma gondii (T. gondii) infection in the CNS, here we provide a method to extract, maintain, dissociate and infect murine astrocytes and microglia cells with protozoa parasites. Extracted cells from newborn cortices are maintained in vitro for 14 days with periodic differential media replacement. Astrocytes and microglia are obtained from the same extraction protocol by mechanical dissociation. After phenotyping by flow cytometry, cells are infected with protozoa parasites. The infection rate is determined by fluorescence microscopy at different time points, thus enabling the evaluation of differential ability of glial cells to control protozoan invasion and replication. These techniques represent simple, cheap and efficient methods to study the responses of astrocytes and microglia to infections, opening the field for further neuroimmunology analysis.

The overall goal of this protocol is to instruct how to extract, maintain, and dissociate murine astrocyte and microglia cells from the central nervous system, followed by infection with protozoa parasites.

Astrocytes and microglia are the most abundant glial cells. They are responsible for physiological support and homeostasis maintenance in the central ...

Primary Microglia Isolation from Postnatal Mouse Brains

Microglia are the mononuclear phagocytes in the central nervous system (CNS), which play key roles in maintaining homeostasis and regulating the inflammatory process in the CNS. To study the microglial biology in vitro, primary microglia show great advantages compared to immortalized microglial cell lines. However, microglia isolation from the postnatal mouse brain is relatively less efficient and time-consuming. In this protocol, we provide a quick and easy-to-follow method to isolate primary microglia from the neonatal mouse brain. The overall steps of this protocol include brain dissection, primary brain cell culture, and microglia isolation. Using this approach, researchers can obtain primary microglia with high purity. In addition, the harvested primary microglia were able to respond to the lipopolysaccharides challenge, indicating they retained their immune function. Collectively, we developed a simplified approach to efficiently isolate primary microglia with high purity, which facilitates a wide range of microglial biology investigations&#160;in vitro.

Primary cell culture is one of the primarily used approaches for studying microglial biology in vitro. Here, we developed a method for simple and rapid microglia isolation from the mouse postnatal day 1 (P1) to P4.

Microglia are the mononuclear phagocytes in the central nervous system (CNS), which play key roles in maintaining homeostasis and regulating the ...

Establishment of an Electrophysiological Platform for Modeling ALS with Regionally-Specific Human Pluripotent Stem Cell-Derived Astrocytes and Neurons

Human pluripotent stem cell-derived astrocytes (hiPSC-A) and neurons (hiPSC-N) provide a powerful tool for modeling Amyotrophic Lateral Sclerosis (ALS) pathophysiology in vitro. Multi-electrode array (MEA) recordings are a means to record electrical field potentials from large populations of neurons and analyze network activity over time. It was previously demonstrated that the presence of hiPSC-A that are differentiated using techniques to promote a spinal cord astrocyte phenotype improved maturation and electrophysiological activity of regionally specific spinal cord hiPSC-motor neurons (MN) when compared to those cultured without hiPSC-A or in the presence of rodent astrocytes. Described here is a method to co-culture spinal cord hiPSC-A with hiPSC-MN and record electrophysiological activity using MEA recordings. While the differentiation protocols described here are particular to astrocytes and neurons that are regionally specific to the spinal cord, the co-culturing platform can be applied to astrocytes and neurons differentiated with techniques specific to other fates, including cortical hiPSC-A and hiPSC-N. These protocols aim to provide an electrophysiological assay to inform about glia-neuron interactions and provide a platform for testing drugs with therapeutic potential in ALS.

We describe a method for differentiating spinal cord human induced pluripotent-derived astrocytes and neurons and their co-culture for electrophysiological recording.

Human pluripotent stem cell-derived astrocytes (hiPSC-A) and neurons (hiPSC-N) provide a powerful tool for modeling Amyotrophic Lateral Sclerosis (ALS) ...

Real-Time Fluorescent Measurement of Synaptic Functions in Models of Amyotrophic Lateral Sclerosis

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe dementia (FTLD) is dysfunction of communication between neurons and motor neurons and muscle. The underlying process of synaptic transmission involves membrane depolarization-dependent synaptic vesicle fusion and the release of neurotransmitters into the synapse. This process occurs through localized calcium influx into the presynaptic terminals where synaptic vesicles reside. Here, the protocol describes fluorescence-based live-imaging methodologies that reliably report depolarization-mediated synaptic vesicle exocytosis and presynaptic terminal calcium influx dynamics in cultured neurons.
Using a styryl dye that is incorporated into synaptic vesicle membranes, the synaptic vesicle release is elucidated. On the other hand, to study calcium entry, Gcamp6m is used, a genetically encoded fluorescent reporter. We employ high potassium chloride-mediated depolarization to mimic neuronal activity. To quantify synaptic vesicle exocytosis unambiguously, we measure the loss of normalized styryl dye fluorescence as a function of time. Under similar stimulation conditions, in the case of calcium influx, Gcamp6m fluorescence increases. Normalization and quantification of this fluorescence change are performed in a similar manner to the styryl dye protocol. These methods can be multiplexed with transfection-based overexpression of fluorescently tagged mutant proteins. These protocols have been extensively used to study synaptic dysfunction in models of FUS-ALS and C9ORF72-ALS, utilizing primary rodent cortical and motor neurons. These protocols easily allow for rapid screening of compounds that may improve neuronal communication. As such, these methods are valuable not only for the study of ALS but for all areas of neurodegenerative and developmental neuroscience research.

Two related methods are described to visualize subcellular events required for synaptic transmission. These protocols enable the real-time monitoring of the dynamics of presynaptic calcium influx and synaptic vesicle membrane fusion using live-cell imaging of in vitro cultured neurons.

Before neuronal degeneration, the cause of motor and cognitive deficits in patients with amyotrophic lateral sclerosis (ALS) and/or frontotemporal lobe ...

 Primary Microglia and Neuron Co-Culture Protocol for CNS Interaction Studies

Generating and Co-culturing Murine Primary Microglia and Cortical Neurons

Microglia are tissue-resident macrophages of the central nervous system (CNS), performing numerous functions that support neuronal health and CNS homeostasis. They are a major population of immune cells associated with CNS disease activity, adopting&#160;reactive phenotypes that potentially contribute to neuronal injury during chronic neurodegenerative diseases such as multiple sclerosis (MS). The distinct mechanisms by which microglia regulate neuronal function and survival during health and disease remain limited due to challenges in resolving the complex in vivo interactions between microglia, neurons, and other CNS environmental factors. Thus, the in vitro approach of co-culturing microglia and neurons remains a valuable tool for studying microglia-neuronal interactions. Here, we present a protocol to generate and co-culture primary microglia and neurons from mice. Specifically, microglia were isolated after 9-10 days in vitro from a mixed glia culture established from brain homogenates derived from neonatal mice between post-natal days 0-2. Neuronal cells were isolated from brain cortices of mouse embryos between embryonic days 16-18. After 4-5 days in vitro, neuronal cells were seeded in 96-well plates, followed by the addition of microglia to form the co-culture. Careful timing is critical for this protocol as both cell types need to reach experimental maturity to establish the co-culture. Overall, this co-culture can be useful for studying microglia-neuron interactions and can provide multiple readouts, including immunofluorescence microscopy, live imaging, as well as RNA and protein assays.

Author Spotlight: In Vitro Co-Culture Model for Studying Microglia-Neuronal Interactions in Disease Conditions

This protocol describes a microglia-neuronal co-culture established from primary neuronal cells isolated from mouse embryos at embryonic days 15-16 and primary microglia generated from the brains of neonatal mice at post-natal days 1-2.