JP acknowledges funding support from the Natural Sciences and Engineering Research Council of Canada and the University of Saskatchewan College of Medicine. YD acknowledges funding support from the University of Saskatchewan College of Medicine Startup Fund, the Natural Sciences and Engineering Research Council of Canada Discovery Grant (RGPIN-2023-03659), MS Canada Catalyst Grant (1019973), Saskatchewan Health Research Foundation Establishment Grant (6368), and Brain Canada Foundation Future Leaders in Canadian Brain Research Grant. Figure 1A, Figure 2A, and Figure 3A were created with BioRender.com.

All animals used in this study were housed and handled with approval from the University Animal Care Committee (UACC) of the University of Saskatchewan and the Canadian Council on Animal Care (CCAC). Post-natal days 0-2 CD1 male and female mice and&#160;embryonic days&#160;16-18 (E16-18) embryos from pregnant CD1 mice were used for this study. The details of the reagents and the equipment used are listed in the Table of Materials.
1. Primary microglia culture
NOTE: It is crucial to time the mixed glia and neuronal cultures so that microglia are mature and ready for harvest within 2 days after neurons are seeded into a 96-well plate.
<ol>
	<li>Preparation
	<ol>
		<li>Pre-warm complete glia culture media, which includes Dulbecco&#39;s Modified Eagle Medium (DMEM) with high glucose supplemented with 10% heat-inactivated fetal bovine serum, 50 U/mL Penicillin/Streptomycin with 2.92 mg/mL of L-glutamine, 1mM sodium pyruvate, 1x non-essential amino acids, and 20 mM of HEPES in a 37 &#176;C water bath.</li>
		<li>Add 5 mL of 10 &#181;g/mL poly-L-ornithine (PO) hydrobromide to each T-75 flask. Gently swirl the flask to ensure that PO completely covers the bottom of the flask. Incubate at 37 &#176;C in a 5% CO2 incubator for at least 1 h. 
		NOTE: This coating step is required to facilitate cell adhesion to the T-75 flasks.</li>
	</ol>
	</li>
	<li>Isolation of brains from P0-P2 neonatal mice 
	NOTE: Sterilize all dissection tools and maintain the sterility of reagents. Ensure adherence to aseptic techniques throughout.
	<ol>
		<li>Before starting dissection, spray down the countertop with 70% ethanol or disinfecting spray where dissection will take place. Set up the dissection microscope and place the Petri dish under the microscope. Pour 20 mL of Leibovitz&#39;s L-15 media into the petri dish. Keep the lid of the Petri dish on while the microscope is not in use to reduce contamination.</li>
		<li>Place down several layers of paper towel and thoroughly spray the paper towels with 70% ethanol. Prepare the dissection tools (dissection scissors, a pair of micro-forceps, and tissue forceps) by thoroughly spraying them with 70% ethanol and laying the tools down on the paper towels.</li>
		<li>Transfer post-natal days 0-2 mouse onto the paper towels and thoroughly spray the mouse with 70% ethanol. Decapitate the mouse using dissection scissors (following institutionally approved protocols).</li>
		<li>While gently holding the head, create a midline incision in the cranium starting from the neck to the nose. Peel the skull back with tissue forceps to reveal the brain.</li>
		<li>Using tissue forceps, gently scoop the brain out by inserting the forceps under the brain and lift the brain out from the head. Immediately place the brain into the&#160;Petri dish containing 20 mL of Leibovitz&#39;s L-15 media.</li>
		<li>Under a dissection microscope, carefully remove the brain stem and the meninges using a pair of micro-forceps. Collect the brains in a 50 mL conical tube containing 5 mL of Leibovitz&#39;s L-15 media.</li>
	</ol>
	</li>
	<li>Brain dissociation, seeding and maintaining mixed glia culture 
	NOTE: Perform all subsequent steps in a biosafety cabinet.
	<ol>
		<li>Mince the brains into approximately 1 mm2 pieces with a sterile disposable scalpel blade. Carefully transfer the minced brains into a new sterile 50 mL tube.</li>
		<li>To the minced brains, add a final concentration of 0.25% trypsin and incubate the tissue in a 37 &#176;C water bath for 20 min. Every 2-3 min, mix the solution by gently inverting the tube to aid tissue dissociation.</li>
		<li>From the 5% CO2 incubator, transfer the T-75 flasks containing 5 mL of PO into the biosafety cabinet. Discard the PO and rinse the flasks 3 times with sterile 1x phosphate buffer solution (PBS). Ensure to thoroughly rinse the flasks as excess PO is toxic to cells.</li>
		<li>Transfer the digested tissue from the 37 &#176;C water bath into the biosafety cabinet. Add 5 mL of complete glia culture media to neutralize the trypsin. Gently mix the tissue suspension using a 10 mL pipette.</li>
		<li>Place a sterile 70 &#181;m nylon mesh cell strainer onto a sterile 50 mL conical tube and wet the mesh by pouring roughly 5 mL of L-15. Carefully decant the cell suspension through the cell strainer. Remove the plunger from a sterile 1 mL syringe, grind the tissue into the mesh using the plunger to get rid of tissue clumps, and generate a homogenized cell suspension.</li>
		<li>Periodically, pour approximately 5 mL of Leibovitz&#39;s L-15 media into the 70 &#181;m nylon mesh and continue grinding the tissue on the mesh with the 1 mL syringe plunger. When little to no visible tissue remains on the mesh, discard the mesh and set the cell suspension aside.</li>
		<li>Mix the cell suspension by gently pipetting up and down. To each flask, add equal volumes of cell suspension containing 1-2 brains with glia culture media to a final volume of 15 mL.</li>
		<li>Incubate the cells at 37 &#176;C in a 5% CO2 incubator overnight. The following day, wash the flasks twice with sterile 1x PBS to remove unattached cells and debris.</li>
		<li>Add 15 mL of fresh complete glia culture media to the culture and incubate at 37 &#176;C in a 5% CO2 incubator. After 3 days, change half the media with 10 mL of fresh glia culture media supplemented with 40 ng/mL of macrophage colony-stimulating factor (MCSF). 
		NOTE: Thereafter, half the media is replaced with 10 mL of fresh glia culture media supplemented with 40 ng/mL of MCSF every 3 days. The culture is maintained for 8-10 days before microglia are harvested.</li>
	</ol>
	</li>
</ol>
2. Primary neuron culture
<ol>
	<li>Preparation
	<ol>
		<li>Thaw B-27 plus supplement from B-27 plus neuronal culture system. Pre-warm neurobasal plus medium from B-27 plus neuronal culture system, 1x Hank&#39;s Balanced Salt Solution (HBSS), and heat-inactivated fetal bovine serum (FBS) at 37 &#176;C in a water bath.</li>
		<li>Coat T-25 flasks with 5 mL of 10 &#181;g/mL PO and incubate the flasks for at least 1 hour at 37 &#176;C. Prepare instruments and supplies for dissection. 
		NOTE: This coating step facilitates cell adhesion to the T-25 flasks.</li>
	</ol>
	</li>
	<li>Brain isolation from E16-E18 mouse embryos 
	NOTE: Sterilize all dissection tools and maintain the sterility of reagents. Ensure adherence to aseptic techniques throughout.
	<ol>
		<li>Before starting dissection, spray down the countertop with 70% ethanol or disinfecting spray where dissection will take place. Set up the dissection microscope and place the Petri dish under the microscope. Pour 20 mL of 1x HBSS into the Petri dish. Keep the lid of the Petri dish on while the microscope is not in use to reduce contamination.</li>
		<li>Place down several layers of paper towel and thoroughly spray the paper towels with 70% ethanol. Prepare the dissection tools (spring scissors, a pair of micro-forceps, and tissue forceps) by thoroughly spraying them with 70% ethanol and laying the tools down on the paper towels.</li>
		<li>Anesthetize a pregnant mouse between gestation days 16-18 using isoflurane and euthanize the mouse by cervical dislocation (following institutionally approved protocols).</li>
		<li>Lay the mouse on its back on a piece of paper towel soaked in 70% ethanol. Disinfect the abdominal area with 70% ethanol. Lift the skin of the lower abdomen with tissue forceps and perform a V-shaped incision from this point to the lower rib cage with dissection scissors.</li>
		<li>Grab the uterine horns containing the embryos with tissue forceps and gently remove the embryos from the abdominal cavity. Briefly disinfect the uterine horns containing embryos with 70% ethanol.</li>
		<li>Dissect one embryo from its individual sac at a time. Using spring scissors, create a midline incision at the top of the skull, starting from the back of the neck to the nose. Then, lift the skull away from the incision site to reveal the brain. Gently scoop out the brain with tissue forceps and transfer the brain to a 10 cm Petri dish containing 15 mL of 1x HBSS.</li>
		<li>Under a dissection microscope, carefully remove the brainstem and the meninges on both sides of the brain cortices with a pair of micro-forceps. Transfer the brain cortices into a 50 mL conical tube containing 10 mL of 1x HBSS and place on ice.</li>
	</ol>
	</li>
	<li>Brain dissociation and seeding of cortical neurons in cell culture 
	NOTE: Perform all subsequent steps in a biosafety cabinet.
	<ol>
		<li>Mince the brains into approximately 1 mm2 pieces in the 50 mL conical tube with a sterile disposable scalpel blade. Transfer the minced brains into a new sterile 50 mL conical tube.</li>
		<li>Add trypsin to the tube with a final concentration of 0.25%. Immerse the tube in a 35-38 &#176;C hot water bath for 15 min and gently mix the tube to homogenize the tissue every 2-3 min.</li>
		<li>Take the tube out of the hot water bath. Gently pipette the tissue suspension up and down to further dissociate cells, then add 0.5 mL of heat-inactivated FBS to the homogenized suspension to neutralize trypsin activity. Avoid air bubbles to minimize toxicity to neuronal cells.</li>
		<li>Place a sterile 70 &#181;m nylon mesh cell strainer onto a sterile 50 mL conical tube and wet the mesh by adding roughly 5 mL of neurobasal media. Filter the cell suspension through the 70 &#181;m nylon mesh cell strainer. Using a 1 mL syringe plunger, gently grind the tissue into the cell strainer. Periodically pour approximately 5 mL of 1x HBSS while continuing grinding to wash the cell strainer.</li>
		<li>Centrifuge the conical tube at 300 x g for 5 min at 4 &#176;C. During this time, dilute B-27 plus supplement in neurobasal plus media to 1x final concentration to use as complete neurobasal media.</li>
		<li>Discard the PO in the T-25 flasks and rinse the flasks three times with 5 mL of 1x PBS.</li>
		<li>Carefully decant to remove the supernatant and resuspend the cell pellet by gently pipetting up and down 2-3 mL of complete neurobasal media. Determine the cell concentration and total cell numbers with a hemocytometer and trypan blue. Expect up to 1.0 x 107 cells per embryo.</li>
		<li>Plate the cells in the PO-coated T-25 flasks with approximately 2.0 x 107 cells per flask in 5 ml of complete neurobasal media. Incubate the cell culture at 37 &#176;C in a 5% CO2 incubator.</li>
		<li>Perform a half media change every 2-3 days by replacing 2.5 mL of media from the culture flask with freshly made complete neurobasal media.</li>
	</ol>
	</li>
</ol>
3. Co-culture of primary neurons and microglia
NOTE: All subsequent steps are to be done in a sterile biosafety cabinet.
<ol>
	<li>Seeding neurons in 96-well plates
	<ol>
		<li>Add 100 &#181;L of 10 &#181;g/mL PO per well and incubate the flasks for at least 1 h at 37 &#176;C to coat the wells with PO. Wash the plate with 1x PBS 3 times before seeding cells and aspirate any remaining liquid after the last wash.</li>
		<li>Neurons grown in T-25 flasks are harvested for co-culture after 2-5 days in culture. Replace the media with 5 mL of 1x Versene solution with 0.25% trypsin and 1 mg/mL DNase I to detach neurons from the flask. Place the flasks inside the incubator for 5-6 min for digestion and gently swirl the flask every 2-3 minutes to aid cell detachment.</li>
		<li>Take the flask out of the incubator and check if cells are detached under a microscope. Add 0.5 mL of heat-inactivated FBS to neutralize the trypsin and DNase I. Transfer cells to a 15 mL conical tube. Wash the flask with 5 mL of complete neurobasal media once to maximize collected neurons. Collect all cells in the same 15 mL conical tube.</li>
		<li>Centrifuge the cell suspension at 300 x g for 5 min at 4 &#176;C. Discard the supernatant and gently resuspend the cell pellet in 2-3 mL of complete neurobasal media.</li>
		<li>Count cells with a hemocytometer and add complete neurobasal media to achieve a final concentration of 7.5 x 105 neurons/mL. Seed 100 &#181;L of the cell suspension to each well in the PO-coated 96-well plate to obtain 7.5 x 104 neurons/well. A yield of 5-6 million neurons per flask is expected.</li>
		<li>Incubate the neurons in a 96-well plate overnight. Add 100 &#181;L of complete neurobasal media to each well the next day. 
		&#8203;NOTE: Once neurons display appropriate neurite process growth (around day 3-4 after seeding in 96-well plates), they are ready for co-culture with microglia.</li>
	</ol>
	</li>
	<li>Isolation and microglia-neuronal co-culture
	<ol>
		<li>Between 8-10 days of mixed glia cultivation, collect microglia by gently washing the T-75 flasks with glia culture media in each flask with a 10 mL pipette, and transfer the cells and media into a sterile 50 mL conical tube. Wash the flasks again with 10 mL fresh glia culture media to detach additional microglia from the flasks and collect the media. A yield of 7.5 x 105 microglia per flask is expected.</li>
		<li>Add 20 mL of fresh glia culture media supplemented with 20 ng/mL MCSF to each flask. Microglia may be harvested once or twice more if the mixed glia culture is maintained.</li>
		<li>Centrifuge the cell collection at 300 x g for 5 min at 4 &#176;C. Carefully remove the supernatant and gently resuspend the pellet in 2 mL of complete neurobasal media.</li>
		<li>Count the cells with a hemocytometer and trypan blue. Add complete neurobasal media to a final concentration of 2.5 x 105 cells/mL.</li>
		<li>From the 96-well neuronal culture plate, remove 100 &#181;L of neurobasal media and add 100 &#181;L of 2.5 x 105 cells/mL cell suspension to seed 2.5 x 104 microglia per well of 7.5 x 104 neurons. Incubate the culture overnight at 37 &#176;C in a 5% CO2 incubator to allow microglia to settle. The co-culture may now be used for downstream experiments.</li>
	</ol>
	</li>
</ol>

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

The authors declare no conflicts of interest.

This article describes a protocol for isolating and culturing mouse primary neurons and primary microglia, which are subsequently used to establish a microglia-neuronal co-culture that can be used to study how microglia and neuron interactions regulate their cellular health and function. This relatively simple and accessible approach can provide critical insights into the mechanisms and functional outcomes of microglia neuron interactions in the CNS.
To achieve an optimal co-culture, several c...

Microglia are tissue-resident macrophages that facilitate immunosurveillance and homeostasis in the central nervous system (CNS)1,2,3. They originate from yolk sac erythromyeloid progenitor cells that colonize the brain during embryonic development4,5,6&#160;and are maintained throughout the organism&#39;s life span through self-renewal, which involves proliferation and apoptosis7. At steady-state, resting microglia have ramified morphology and engage in tissue surveillance8,9,10.
Microglia express numerous cell-surface receptors, which enables them to rapidly respond to changes in the CNS11,12 and to promote inflammatory responses in the event of infections or tissue injury12,13,14, as well as during neurodegenerative diseases9,15, such as&#160;multiple sclerosis (MS)16,17. Microglia also express receptors to various neurotransmitters and neuropeptides18,19,20, which suggests they may also respond to and regulate neuronal activity21,22. Indeed, microglia and neurons interact in various forms of bidirectional communication8,23 such as direct interactions mediated by membrane proteins or indirect interactions through soluble factors or intermediate cells23,24.
For instance, various neurotransmitters secreted by neurons can modulate the neuroprotective or inflammatory activity of microglia25,26,27. Additionally, direct interactions between neurons and microglia help to maintain microglia in a homeostatic state28. Conversely, direct interactions of microglia with neurons can shape neuronal circuitry29 and influence neuronal signaling30,31,32. As disruptions of these interactions induce hyperexcitability of neurons30 and&#160;microglia reactivity33,34, dysregulated microglia-neuronal interactions&#160;are implicated&#160;as a contributing factor to&#160;neurological diseases33,35. Indeed, psychotic23,26 and neurodegenerative diseases have been described to exhibit dysfunctional microglia-neuronal interactions33. While these observations highlight the importance of microglia-neuronal communication in the CNS, specific mechanisms of how these interactions regulate microglial and neuronal functions in health and disease are relatively unknown.
Within a complex milieu such as the CNS, multiple environmental factors can influence microglia-neuronal interactions, which limits the ability to study transient cellular interactions in vivo. Here, we present an in vitro microglia-neuronal co-culture system that can be used to study direct cellular interactions between microglia and neurons. This protocol describes the generation of primary microglia and neurons from the cortices of neonatal mice between post-natal days 0-2 and embryonic mice days 16-18, respectively. Neurons and microglia are then co-cultured in 96-well plates for downstream high-throughput experiments. We previously used this approach to demonstrate that microglia phagocytosis protects neurons from oxidized phosphatidylcholine mediated cell death37, suggesting that this method can help to understand the roles of microglia in the context of neurodegeneration and MS. Similarly, microglia-neuronal co-cultures may also be useful for investigating the impact of microglia-neuronal crosstalk in other contexts such as viral infections38 or neuronal injury and repair39. Overall, in vitro microglia-neuronal co-culture systems&#160;enable researchers to study microglia-neuronal interactions in a manipulatable and controlled environment, which complements in vivo models.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10 cm Petri dish&#160;</td>
			<td>Fisher&#160;</td>
			<td>07-202-011</td>
			<td>Sterile</td>
		</tr>
		<tr>
			<td>1x Versene</td>
			<td>Gibco</td>
			<td>15040-066</td>
			<td></td>
		</tr>
		<tr>
			<td>B-27 Plus Neuronal Culture System&#160;</td>
			<td>Gibco&#160;</td>
			<td>A3653401</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection microscope</td>
			<td>VWR</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>DNase I</td>
			<td>Roche</td>
			<td>11284932001</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#8217;s Modified Eagle Medium (DMEM)</td>
			<td>Gibco</td>
			<td>11960-044</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum&#160;</td>
			<td>ThermoFisher Sci</td>
			<td>12483-020</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS (10x)</td>
			<td>Gibco</td>
			<td>14065-056</td>
			<td></td>
		</tr>
		<tr>
			<td>Hemacytometer</td>
			<td>Hausser Scientific</td>
			<td>1475</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES&#160;</td>
			<td>ThermoFisher Sci</td>
			<td>15630080</td>
			<td></td>
		</tr>
		<tr>
			<td>Leibovitz&#8217;s L-15 Medium (1x)</td>
			<td>Fisher Scientific&#160;</td>
			<td>21083027</td>
			<td></td>
		</tr>
		<tr>
			<td>Macrophage colony stimulating factor&#160;</td>
			<td>Peprotech</td>
			<td>315-02</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro-Forceps</td>
			<td>RWD</td>
			<td>F11020-11</td>
			<td>Autoclaved/Sterile</td>
		</tr>
		<tr>
			<td>Non-essential amino acids</td>
			<td>Cytiva</td>
			<td>SH3023801</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS (10x)</td>
			<td>ThermoFisher Sci</td>
			<td>AM9625</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin Streptomycin Glutamine (100x)</td>
			<td>Gibco</td>
			<td>103780-16</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly-L-ornithine hydrobromide&#160;</td>
			<td>Sigma</td>
			<td>P3655-100MG</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium pyruvate (100 mM)</td>
			<td>Gibco</td>
			<td>11360-070</td>
			<td></td>
		</tr>
		<tr>
			<td>Spring scissors</td>
			<td>RWD</td>
			<td>S11008-42</td>
			<td>Autoclaved/Sterile</td>
		</tr>
		<tr>
			<td>Surgical blade</td>
			<td>Feather</td>
			<td>08-916-5D</td>
			<td>Sterile</td>
		</tr>
		<tr>
			<td>T-25 flasks</td>
			<td>Fisher</td>
			<td>10-126-9</td>
			<td></td>
		</tr>
		<tr>
			<td>T-75 flasks&#160;</td>
			<td>Fisher</td>
			<td>13-680-65</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue forceps</td>
			<td>Codman</td>
			<td>30-4218</td>
			<td>Autoclaved/Sterile</td>
		</tr>
		<tr>
			<td>Tissue scissors</td>
			<td>RWD</td>
			<td>S12052-10</td>
			<td>Autoclaved/Sterile</td>
		</tr>
		<tr>
			<td>Trypan Blue&#160;</td>
			<td>Thermofisher Sci&#160;</td>
			<td>15250-061</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin (2.5%)</td>
			<td>ThermoFisher Sci</td>
			<td>15090046</td>
			<td></td>
		</tr>
		<tr>
			<td>Widefield Immunofluorescence Microscope</td>
			<td>Zeiss</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

A flowchart showing the key steps of the mixed glia&#160;culture for microglia is shown in Figure 1A. Overall, sparse cells and excessive cellular debris are expected on day 1 (Figure 1B). By day 4, increased cell number should be observed, especially with the generation of adherent astrocytes, as indicated by their elongated morphology (Figure 1C). A few microglia may be observed on top of the astrocytes or as small round cells flo...

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.

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 ...

generating-co-culturing-murine-primary-microglia-cortical

Research

JoVE Journal

Neuroscience

1.7K Views.  University of Saskatchewan. 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.

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

In utero Electroporation followed by Primary Neuronal Culture for Studying Gene Function in Subset of Cortical Neurons

In vitro study of primary neuronal cultures allows for quantitative analyses of neurite outgrowth. In order to study how genetic alterations affect neuronal process outgrowth, shRNA or cDNA constructs can be introduced into primary neurons via chemical transfection or viral transduction. However, with primary cortical cells, a heterogeneous pool of cell types (glutamatergic neurons from different layers, inhibitory neurons, glial cells) are transfected using these methods. The use of in utero electroporation to introduce DNA constructs in the embryonic rodent cortex allows for certain subsets of cells to be targeted: while electroporation of early embryonic cortex targets deep layers of the cortex, electroporation at late embryonic timepoints targets more superficial layers. Further, differential placement of electrodes across the heads of individual embryos results in the targeting of dorsal-medial versus ventral-lateral regions of the cortex. Following electroporation, transfected cells can be dissected out, dissociated, and plated in vitro for quantitative analysis of neurite outgrowth. Here, we provide a step-by-step method to quantitatively measure neuronal process outgrowth in subsets of cortical cells.
The basic protocol for in utero electroporation has been described in detail in two other JoVE articles from the Kriegstein lab 1, 2. We will provide an overview of our protocol for in utero electroporation, focusing on the most important details, followed by a description of our protocol that applies in utero electroporation to the study of gene function in neuronal process outgrowth.

In utero Electroporation followed by Primary Neuronal Culture for Studying Gene Function in Subset of Cortical Neurons

In utero electroporation is a valuable method for transfecting neuronal progenitor cells in vivo. Depending upon the placement of the electrodes and the developmental timepoint of electroporation, certain subsets of cortical cells can be targeted. Targeted cells can then be analyzed in vivo or in vitro for effects of genetic alteration.

In vitro study of primary neuronal cultures allows for quantitative analyses of neurite outgrowth. In order to study how genetic alterations affect ...

Nucleofection and Primary Culture of Embryonic Mouse Hippocampal and Cortical Neurons

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and synapse formation and function. An advantage of culturing these neurons is that they readily polarize, forming distinctive axons and dendrites, on a two dimensional substrate at very low densities. This property has made them extremely useful for determining many aspects of neuronal development. Furthermore, by providing glial conditioning for these neurons they will continue to develop, forming functional synaptic connections and surviving for several months in culture. In this protocol we outline a technique to dissect, culture and transfect embryonic mouse hippocampal and cortical neurons. Transfection is accomplished by electroporating DNA into the neurons before plating via nucleofection. This protocol has the advantage of expressing fluorescently-tagged fusion proteins early in development (~4-8hrs after plating) to study the dynamics and function of proteins during polarization, axon outgrowth and branching. We have also discovered that this single transfection before plating maintains fluorescently-tagged fusion protein expression at levels appropriate for imaging throughout the lifetime of the neuron (&gt; 2 months in culture). Thus, this methodology is useful for studying protein localization and function throughout CNS development with little or no disruption of neuronal function.

This protocol outlines the steps required to dissect, transfect via electroporation and culture mouse hippocampal and cortical neurons. Short-term cultures may be used for studies of axon outgrowth and guidance, while long-term cultures can be used for studies of synaptogenesis and dendritic spine analysis.

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and ...

Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP. A significant loss of &Delta;&Psi;m renders cells depleted of energy with subsequent death. Reactive oxygen species (ROS) are important signaling molecules, but their accumulation in pathological conditions leads to oxidative stress. The two major sources of ROS in cells are environmental toxins and the process of oxidative phosphorylation. Mitochondrial dysfunction and oxidative stress have been implicated in the pathophysiology of many diseases; therefore, the ability to determine &Delta;&Psi;m and ROS can provide important clues about the physiological status of the cell and the function of the mitochondria. 
Several fluorescent probes (Rhodamine 123, TMRM, TMRE, JC-1) can be used to determine &Delta;&psi;m in a variety of cell types, and many fluorescence indicators (Dihydroethidium, Dihydrorhodamine 123, H2DCF-DA) can be used to determine ROS. Nearly all of the available fluorescence probes used to assess &Delta;&Psi;m or ROS are single-wavelength indicators, which increase or decrease their fluorescence intensity proportional to a stimulus that increases or decreases the levels of &Delta;&Psi;m or ROS. Thus, it is imperative to measure the fluorescence intensity of these probes at the baseline level and after the application of a specific stimulus. This allows one to determine the percentage of change in fluorescence intensity between the baseline level and a stimulus. This change in fluorescence intensity reflects the change in relative levels of &Delta;&Psi;m or ROS. In this video, we demonstrate how to apply the fluorescence indicator, TMRM, in rat cortical neurons to determine the percentage change in TMRM fluorescence intensity between the baseline level and after applying FCCP, a mitochondrial uncoupler. The lower levels of TMRM fluorescence resulting from FCCP treatment reflect the depolarization of mitochondrial membrane potential. We also show how to apply the fluorescence probe H2DCF-DA to assess the level of ROS in cortical neurons, first at baseline and then after application of H2O2. This protocol (with minor modifications) can be also used to determine changes in &#x2206;&#x03A8;m and ROS in different cell types and in neurons isolated from other brain regions.

We demonstrate application of the fluorescence indicator, TMRM, in cortical neurons to determine the relative changes in TMRM fluorescence intensity before and after application of a specific stimulus. We also show application of the fluorescence probe H2DCF-DA to assess the relative level of reactive oxygen species in cortical neurons.

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP.  A ...

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 ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Calcium Phosphate Transfection of Primary Hippocampal Neurons

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this method on hard-to-transfect cells like primary neurons. Here we describe our detailed protocol for calcium phosphate transfection of hippocampal neurons cocultured with astroglial cells.

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this ...

Primer for Immunohistochemistry on Cryosectioned Rat Brain Tissue: Example Staining for Microglia and Neurons

Immunohistochemistry is a widely used technique for detecting the presence, location, and relative abundance of antigens in situ. This introductory level protocol describes the reagents, equipment, and techniques required to complete immunohistochemical staining of rodent brain tissue, using markers for microglia and neuronal elements as an example. Specifically, this paper is a step-by-step protocol for fluorescent visualization of microglia and neurons via immunohistochemistry for Iba1 and Pan-neuronal, respectively. Fluorescence double-labeling is particularly useful for the localization of multiple proteins within the same sample, providing the opportunity to accurately observe interactions between cell types, receptors, ligands, and/or the extracellular matrix in relation to one another as well as protein co-localization within a single cell. Unlike other visualization techniques, fluorescence immunohistochemistry staining intensity may decrease in the weeks to months following staining, unless appropriate precautions are taken. Despite this limitation, in many applications fluorescence double-labeling is preferred over alternatives such as 3,3&#39;-diaminobenzidine tetrahydrochloride (DAB) or alkaline phosphatase (AP), as fluorescence is more time efficient and allows for more precise differentiation between two or more markers. The discussion includes troubleshooting tips and advice to promote success.

This introductory level protocol describes the reagents, equipment, and techniques required to complete immunohistochemical staining of rodent brains, using markers for microglia and neuronal elements as an example.

Immunohistochemistry is a widely used technique for detecting the presence, location, and relative abundance of antigens in situ. This introductory level ...

Rapid and Refined CD11b Magnetic Isolation of Primary Microglia with Enhanced Purity and Versatility

Microglia are the primary responders to central nervous system insults; however, much remains unknown about their role in regulating neuroinflammation. Microglia are mesodermal cells that function similarly to macrophages in surveying inflammatory stress. The classical (M1-type) and alternative (M2-type) activations of macrophages have also been extended to microglia in an effort to better understand the underlying interplay these phenotypes have in neuroinflammatory conditions such as Parkinson&#39;s, Alzheimer&#39;s, and Huntington&#39;s Diseases. In vitro experimentation utilizing primary microglia offers rapid and reliable results that may be extended to the in vivo environment. Although this is a clear advantage over in vivo experimentation, isolating microglia while achieving adequate yields of optimal purity has been a challenge. Common methods currently in use either suffer from low recovery, low purity, or both. Herein, we demonstrate a refinement of the column-free CD11b magnetic separation method that achieves a high cell recovery and enhanced purity in half the amount of time. We propose this optimized method as a highly useful model of primary microglial isolation for the purposes of studying neuroinflammation and neurodegeneration.

Here, we present a protocol to isolate microglia from postnatal mouse pups (day 1) for in vitro experimentation. This improvised method of isolation generates both high yield and purity, a significant advantage over alternate methods that allows broad range experimentation for the purposes of elucidating microglial biology.

Microglia are the primary responders to central nervous system insults; however, much remains unknown about their role in regulating neuroinflammation. ...

Modeling Neuronal Death and Degeneration in Mouse Primary Cerebellar Granule Neurons

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their post-natal development, abundance, and accessibility, CGNs are an ideal model to study neuronal processes, including neuronal development, neuronal migration, and physiological neuronal activity stimulation. In addition, CGN cultures provide an excellent model for studying different modes of cell death including excitotoxicity and apoptosis. Within a week in culture, CGNs express N-methyl-D-aspartate (NMDA) receptors, a specific ionotropic glutamate receptor with many critical functions in neuronal health and disease. The addition of low concentrations of NMDA in conjunction with membrane depolarization to rodent primary CGN cultures has been used to model physiological neuronal activity stimulation while the addition of high concentrations of NMDA can be employed to model excitotoxic neuronal injury. Here, a method of isolation and culturing of CGNs from 6 day old pups as well as genetic manipulation of CGNs by adenoviruses and lentiviruses are described. We also present optimized protocols on how to stimulate NMDA-induced excitotoxicity, low-potassium-induced apoptosis, oxidative stress and DNA damage following transduction of these neurons.

This protocol describes a simple method for isolating and culturing primary mouse cerebral granule neurons (CGNs) from 6-7 day old pups, efficient transduction of CGNs for loss and gain of function studies, and modelling NMDA-induced neuronal excitotoxicity, low-potassium-induced cell death, DNA-damage, and oxidative stress using the same culture model.

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their ...

Characterization and Isolation of Mouse Primary Microglia by Density Gradient Centrifugation

Microglia, the resident immune cells in the brain, are the first responders to inflammation or injury in the central nervous system. Recent research has revealed microglia to be dynamic, capable of assuming both pro-inflammatory and anti-inflammatory phenotypes. Both M1 (pro-inflammatory) and M2 (pro-reparative) phenotypes play an important role in neuroinflammatory conditions such as perinatal brain injury, and exhibit differing functions in response to certain environmental stimuli. The modulation of microglial activation has been noted to confer neuroprotection thus suggesting microglia may have therapeutic potential in brain injury. However, more research is required to better understand the role of microglia in disease, and this protocol facilitates that. The protocol described below combines a density gradient centrifugation process to reduce cellular debris, with magnetic separation, producing a highly pure sample of primary microglial cells that can be used for in vitro experimentation, without the need for 2-3 weeks culturing. Additionally, the characterization steps yield robust functional data about microglia, aiding studies to better our understanding of the polarization and priming of these cells, which has strong implications in the field of regenerative medicine.

A protocol for the isolation of primary microglia from murine brains is presented. This technique aids in furthering the current understanding of neurological conditions. Density gradient centrifugation and magnetic separation are combined to produce sufficient yield of a highly pure sample. Furthermore, we outline the steps for characterization of microglia.

Microglia, the resident immune cells in the brain, are the first responders to inflammation or injury in the central nervous system. Recent research has ...

Generating and Co-culturing Murine Primary Microglia and Cortical Neurons

Summary

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Chapters in this video

In utero Electroporation followed by Primary Neuronal Culture for Studying Gene Function in Subset of Cortical Neurons

Nucleofection and Primary Culture of Embryonic Mouse Hippocampal and Cortical Neurons

Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Bilaminar Co-culture of Primary Rat Cortical Neurons and Glia

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Calcium Phosphate Transfection of Primary Hippocampal Neurons

Primer for Immunohistochemistry on Cryosectioned Rat Brain Tissue: Example Staining for Microglia and Neurons

Rapid and Refined CD11b Magnetic Isolation of Primary Microglia with Enhanced Purity and Versatility

Modeling Neuronal Death and Degeneration in Mouse Primary Cerebellar Granule Neurons

Characterization and Isolation of Mouse Primary Microglia by Density Gradient Centrifugation