This work was supported by a research grant MOP 119271 to JMW by the Canadian Institute of Health Research. The authors would like to thank Ms. Yao Fang Tan for her outstanding technical assistance.

NOTE: All animal procedures conformed to the animal health and welfare guidelines of the Department of Comparative Medicine at the University of Toronto.
1. Preparation of Hippocampal Slices
<ol>
	<li>Sterilize the following instruments using the dry autoclave at 125 &#176;C: Scalpel handle (#3) (2), Standard pattern forceps, large (1), Small dissector scissor (angled to side) (1), Micro spoon (spoon and flat spatula ends) (1), Micro-spatulas (rounded and rounded tapered ends) (2), Fine paintbrush (1), Fire polished Pasteur pipette (2), Gauze squares, 2 x 2 inches (5).</li>
	<li>When sterile, put the instruments into a sterile container and keep covered until use. Immediately before dissection, immerse instruments in a 70% ethanol solution.</li>
	<li>Prepare a 6-well culture plate with culture insert before beginning the dissection procedure by adding 1 ml of culture medium/well and storing the plate in the incubator at 35 &#176;C and 5% CO2.</li>
</ol>
2. Arrange Dissection Tools in Sterile Laminar Flow hood
<ol>
	<li>Spray the laminar flow hood with 70% ethanol and remove sterilized dissection instruments from alcohol. Allow the instruments to dry while resting on a sterile Petri dish to avoid contact between the alcohol and the dissected brain tissue.</li>
	<li>Deposit 5-7 ml of sterilized ice-cold dissecting solution in 2 large sterile Petri dishes. One dish will chill and clean the head (dirty), the other one for cooling and rinsing the scooped out brain (clean). Place a sterilized filter paper in one of the Petri dish lids for dissecting out the brain.</li>
	<li>Place a small, sterilized filter paper in one of the small Petri dish lid for dissecting out the hippocampus. Deposit 3-5 ml of sterilized ice-cold dissecting solution in 2 small, sterile Petri dishes. One dish will hold the scooped out hippocampus, one will hold sections during separation of hippocampal sections under dissection microscope.</li>
	<li>Prepare the tissue chopper by taping a piece of sterilized filter paper to the cutting stage and mounting a sterile razor blade. Wet the filter paper with sterilized dissecting solution.</li>
	<li>Spray a clean bio-bag with alcohol and place it in laminar flow clean bench.</li>
</ol>
3. Hippocampal Dissection
<ol>
	<li>Spray the P7 Sprague Dawley rat pup with 70% ethanol outside of the laminar flow clean bench and quickly decapitate the animal using large sterile surgical scissors inside the laminar flow bench. Let the head drop into ice-cold dissecting solution in one of the Petri dishes.</li>
	<li>In the Petri dish, rinse off the blood and quickly transfer the head to sterilized filter paper, ventral side down.</li>
	<li>Using the scalpel, cut along the dorsal surface in the sagittal plane to expose the underlying skull. Cut through the skin, but not the underlying bone, which is soft and easily penetrable in rats of this age. Set aside this &#8220;dirty&#8221; scalpel and do not use on brain tissue.</li>
	<li>Using the small dissector scissors (angled to side) and forceps, cut open the skull along sagittal suture of the skull to bregma, the anatomical point on the skull where the coronal suture is intersected perpendicularly by the sagittal suture. Use forceps to pull skull flaps up and away from the midline of the skull.</li>
	<li>Place the micro spoon on the underside of the brain, beneath the brain stem, to gently lift the brain out of the skull. Lift the brain to expose the optic nerves and olfactory bulb on basal surface of brain. Cut these structures with small scissors to fully detach the brain from skull. Remove and transfer the intact brain to the other large Petri dish containing ice-cold dissecting solution.</li>
	<li>Using the micro spoon, transfer the brain to a small Petri-dish lid containing sterile filter paper. With a sterile Pasteur pipette, rinse the brain with a few drops of dissecting solution to keep tissue moist.</li>
	<li>Using a &#8220;clean&#8221; scalpel blade cut the two hemispheres apart. Transfer the left hemisphere back to large Petri dish with micro spoon and place hemisphere pia side down in ice-cold dissecting solution for subsequent use.</li>
	<li>View the medial face of the right hemisphere and identify the edge fornix, a prominent band of white matter along the medial edge of the hippocampus. Using a sterile scalpel, make a sagittal cut through the fornix, but take care because only 0.5 cm of the scalpel tip will be sufficient to cut the fornix.</li>
	<li>Using 2 micro-spatulas, remove the first hippocampus from right hemisphere by placing the right-hand-spatula on the brain stem and lifting the overlying cortex with the left-hand spatula. Gently lift the cortex to reveal the lateral ventricle and medial surface of the hippocampus. A white curved line, the fimbria, should now be visible.</li>
	<li>Align the curvature of spatula with the curvature of the fimbria and gently press the spatula under the fimbria. Slide the spatula left and ride along rostral-caudal axis and then lift spatula in dorsal direction to remove hippocampus.</li>
	<li>Transfer the hippocampus to a 2nd small Petri dish with ice-cold dissecting solution. Repeat the same procedure on the left hemisphere to remove the left hippocampus.</li>
	<li>Using a micro spatula, carefully transfer the hippocampi to the tissue chopper stage. Arrange them adjacent and parallel to each other and perpendicular to the axis of the chopper blade. Use a paintbrush to position the tissue and add a few drops of the dissecting solution on top of the hippocampi.</li>
	<li>Cut the tissue in 400 &#956;m slices without pausing to remove individual slices (usually they will not adhere to the blade). After the whole hippocampus has been cut, use the paintbrush to gently transfer the sections to a 2nd small Petri dish with dissection solution.</li>
	<li>Under a dissecting microscope, carefully separate the floating slices using amicro-spatula and paintbrush.</li>
	<li>Remove the pre-prepared culture plate with culture insert from the incubator and place in a laminar flow hood.</li>
	<li>Using a fire-polished Pasteur pipette, draw 4-5 slices into the pipette and transfer slices to the apical surface of culture insert membrane. Next, adjust the positioning with a paintbrush and leave space between individual sections and the border of the culture insert.</li>
	<li>Using a sterile Pasteur pipette, remove the excess dissecting solution from apical surface of membrane. 
	NOTE: While removing solution avoid drawing tissue sections into the pipette. Alternatively, use a regular pipette (P200 or P1000) with sterilized pipette tips to slowly remove solution.</li>
	<li>Place the culture plate with serum-containing culture medium and the hippocampal slices back into the incubator at 35 &#176;C and 5% CO2. 
	NOTE: If experiment calls for generating cultures from multiple animals, thoroughly clean the laminar flow clean bench between dissections. Return instruments to alcohol solution and replace all Petri-dishes, filter papers and sterile razor blades prior to second dissection.</li>
</ol>
4. Feeding and Maintaining Organotypic Slices
<ol>
	<li>Feed cultures in a sterile laminar flow clean bench.</li>
	<li>Perform the first feeding of the cultured sections 2 days post-dissection. Aspirate old culture medium using sterilized glass pipette.</li>
	<li>Use sterile 5 ml serological pipette to add 1 ml of fresh, sterile, serum-containing medium to the wells.</li>
	<li>Gently replace the culture insert and take care to remove any air bubbles that may have formed underneath membrane surface.</li>
	<li>After the first feeding, change medium every other day.</li>
</ol>
5. Incubating Tissue Slices with Thymidine Analogues to Label Newborn Neurons
<ol>
	<li>In order to study the maturation and integration of dentate granule cells in the hippocampus, incubate the organotypic slices with a thymidine analogue, such as Bromodeoxyuridine (BrdU) or Chlorodeoxyuridine (CldU). Here, we use CldU because of its higher solubility in saline.</li>
	<li>After 3DIV, add 1 &#956;l of 10 mg/ml CldU stock solution in saline to 1 ml of culture medium for a final concentration of 10 &#956;g/ml. If BrdU is used, correct the concentration for differences in molecular weight. Add medium with CldU to culture wells and incubate tissue with CldU-containing medium for 2 hr at 35 &#176;C.</li>
	<li>Following 2 hr of incubation at 35 &#176;C, remove the CldU-containing medium and replace with regular feeding medium to resume normal feeding schedule (outlined above).</li>
</ol>
6. Tissue Fixation and Storage
<ol>
	<li>Establish a timeline for applying treatments and fixing tissue samples before starting culture experiments (Figure 2B).</li>
	<li>At a predetermined day post-dissection, prepare the following in a laboratory fume hood: a 10-50 ml beaker containing 4% paraformaldehyde (PFA) in phosphate buffered saline (PBS); an empty beaker for discarded culture medium; small forceps; and a 1,000 ml pipette with disposable tips.</li>
	<li>Remove the culture plate(s) from incubation chamber and transfer to a fume hood. Tilt the individual well plate inserts with forceps. Next, use a pipette to remove culture medium and transfer to a disposal beaker. Tilt the culture plate at an angle to help ensure that all culture medium is removed.</li>
	<li>Finish removing medium for one culture plate at a time. When medium has been removed, add 1 ml of 4% PFA to each culture well and seal well plate with parafilm. Repeat for as many culture plates as necessary and transfer plates to refrigerator at 4 &#176;C for 24 hr.</li>
	<li>After 24 hr, prepare the following in a fume hood: 10-50 ml beaker containing 0.1% Sodium Azide in PBS; an empty beaker for discarded PFA (follow safety precautions when discarding toxic substances); small forceps; and 1,000 ml pipette with disposable tips.</li>
	<li>Follow the procedure outlined in 6.4 for adding PFA, but add 1 ml of PBS with sodium azide instead. Once completely transferred, seal well plates for future use by wrapping edges in parafilm and storing in refrigerator at 4 &#176;C.</li>
</ol>
7. Sectioning Tissue for Immunohistochemistry
<ol>
	<li>Perform tissue sectioning using a vibratome. The following series of steps help maximize the yield of usable tissue sections from organotypic cultures. 
	NOTE: Immediately following hippocampal dissection and plating of slices, the tissue has a thickness of approximately 400 &#956;m. However, after 2-3 weeks in the incubation chamber, tissue slices will begin to flatten, resulting in a final section thickness of 150-300 &#956;m.</li>
	<li>Prepare the following items to section cultured tissue: a #11 scalpel blade and handle, a glass Petri dish containing ice cold PBS, a micro-dissection forceps, and a clean vibratome cutting stage to mount tissue.</li>
	<li>Use a scalpel to carefully cut along the perimeter of the circular insert membrane so that it can be detached from the plastic insert housing. Leave ample space between the cultured slice and the scalpel.</li>
	<li>Transfer the detached insert membrane to a Petri dish containing PBS. After rinsing in PBS, use forceps to transfer the membrane to the vibratome mounting stage.</li>
	<li>Next, use the scalpel to eliminate excess membrane surrounding cultured slices and cut away excess material to create clean edges. This step will ensure the membrane is flat and can easily adhere to the cutting surface.</li>
	<li>Place 1-2 drops of adhesive on the vibratome cutting stage and spread in even layer using a 22 G needle. Spread adhesive into a rectangular shape with the long edge parallel to the cutting blade of the vibratome. Perform this step quickly, to prevent the adhesive from drying.</li>
	<li>Use forceps to transfer the trimmed membrane containing hippocampal slices to the cutting stage and gently position the membrane on glue and ensure that there are no air bubbles.</li>
	<li>As the superglue dries, transfer the vibratome stage with the glued membrane back to the PBS containing Petri dish. Prepare the vibratome blade and a 48 well plate containing sodium azide to store tissue sections.</li>
	<li>Use the vibratome to generate 30 &#956;m sections of the organotypic slice tissue and transfer to a 48 well plate containing sodium azide for storage and subsequent immunohistochemical staining.</li>
	<li>Refer to pre-existing protocols for immunohistochemical staining26,29.</li>
</ol>

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The authors declare that they have no competing financial interests.

Following CldU (or BrdU) administration, the timeline of application of pharmacological agents can be chosen to target newborn DGCs during particular developmental windows. For example, a hypothetical agent can be applied during the second week post-CldU injection, which is proposed to coincide with the age of immature neurons that are at a developmental stage where GABA is depolarizing. Future studies using this protocol could adapt the pharmacological agent and the window of exposure to &#8220;tailor&#8221; the approac...

Adult neurogenesis in the mammalian hippocampus represents a remarkable example of the brain&#8217;s innate capacity for adaptability and plasticity. Dentate granule cells (DGCs) are generated from a renewable pool of neural progenitor cells in the hippocampal dentate gyrus, which is one of the two presently well-characterized neurogenic regions in the mammalian brain, and is thought to be particularly important for learning and memory. The hippocampus is part of the limbic system and has a deep location within the mammalian brain, which makes it a difficult target for precise pharmacological manipulation. Additionally, aberrant neurogenesis has been implicated in conditions, such as epilepsy, schizophrenia, and Alzheimer&#8217;s disease, which has prompted interest in understanding the influence of various pharmacological agents during the maturation and survival of newborn neurons. The distinction between postnatal and adult neurogenesis is blurred and previous studies have shown that many features of in vivo neuronal development in the early postnatal period and adulthood are similar25. Here we emphasize postnatal neurogenesis and suggest possible applications to adult neurogenesis.
Organotypic slice cultures provide an efficient in vitro method for studying various physiological properties of the mammalian hippocampus. The value of slice cultures prepared from rodent brains can be summarized in three main qualities: 1) the protocol is straightforward and requires readily available materials; 2) slice cultures allow for pharmacological studies that eliminate complex variables such as the deep anatomic location of the hippocampus and circumvents the blood brain barrier1; and 3) the well characterized structure of the hippocampus and tri-synaptic circuit is preserved2. Previous investigators have used the organotypic hippocampal culture to study synaptic development and physiology3,4, gliogenesis5-7, ischemic brain damage8,9, neuroprotection and neurorepair10-12 as well as epilepsy13-15.The slice cultures could also provide a useful model system allowing for the monitoring of cell development in conjunction with labeling of cells with green fluorescent protein (GFP) or other vital markers.
Slice cultures have also been previously employed to study postnatal hippocampal neurogenesis16-19, but one important factor in the majority of these studies is the well-characterized degeneration that results from explanting tissue from adult animals after approximately 2 weeks in vitro20,21. For this reason, slice cultures are typically prepared from early post-natal (P5-P10) mice or rat pups, which utilizes the improved viability of early postnatal brain tissue for culturing22. While previous studies have shown that the early postnatal and adult hippocampus differ with regards to synaptic physiology and the expression of specific neuronal subtypes23,24, there is substantial conservation of the choreographed developmental program that newborn dentate granule cells proceed through during maturation25. Additionally, recent studies have suggested that the physiological characteristics of newborn DGCs in culture are very similar to immature neurons in the acute hippocampal slice preparation16.

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			<td rowspan="2" style="width:131px;">Company</td>
			<td rowspan="2" style="width:135px;">Catalog Number</td>
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			<td height="20" style="height:20px;width:265px;">5-chloro-2&#39;-deoxyuridine (CldU)</td>
			<td style="width:131px;">MP Biomedicals</td>
			<td align="right" style="width:135px;">105478</td>
			<td style="width:239px;">Hazardous, Carcinogenic</td>
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			<td height="80" style="height:80px;width:265px;">Cell culture inserts, 30mm diameter, 0.4&#181;m pore size</td>
			<td style="width:131px;">Thermo scientific&#160;</td>
			<td align="right" style="width:135px;">140660</td>
			<td style="width:239px;">Nuclon delta coating on these inserts provides better tissue adhesion and improves slice quality.</td>
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			<td style="width:131px;">Gibco</td>
			<td style="width:135px;">16050-122</td>
			<td style="width:239px;">Make 50 mL aliquots and store at -20&#176;C</td>
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			<td style="width:131px;">Kimberly-Clarke</td>
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			<td style="width:131px;">Fisher Brand</td>
			<td style="width:135px;">FB0875712 and FB0875713A</td>
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			<td height="40" style="height:40px;width:265px;">Scalpel blades #11</td>
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			<td style="width:131px;">Sorfa Medical Plastic Co.</td>
			<td style="width:135px;">P8050</td>
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			<td style="width:135px;">11000-12</td>
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organotypic slice cultures for studies of postnatal neurogenesis

Here we describe a technique for studying hippocampal postnatal neurogenesis in the rodent brain using the organotypic slice culture technique. This method maintains the characteristic topographical morphology of the hippocampus while allowing direct application of pharmacological agents to the developing hippocampal dentate gyrus. Additionally, slice cultures can be maintained for up to 4 weeks and thus, allow one to study the maturation process of newborn granule neurons. Slice cultures allow for efficient pharmacological manipulation of hippocampal slices while excluding complex variables such as uncertainties related to the deep anatomic location of the hippocampus as well as the blood brain barrier. For these reasons, we sought to optimize organotypic slice cultures specifically for postnatal neurogenesis research.

Determining if organotypic cultures would be suitable for adult neurogenesis research required that they satisfy two main criteria: 1) that slices maintain characteristic morphological features of hippocampal slices after 10-21 days in vitro (DIV), and 2) that newborn DGCs can be quantified using standard immunohistochemical techniques commonly employed in adult neurogenesis research. Regarding the first criterion, Figure 1A and 1B highlight the preserved hippocampal morphology....

Watch this Scientific Journal Video about Organotypic Slice Cultures for Studies of Postnatal Neurogenesis at JoVE.com

Organotypic Slice Cultures for Studies of Postnatal Neurogenesis

Here we describe a technique for studying hippocampal postnatal neurogenesis using the organotypic slice culture technique. This method allows for in vitro manipulation of adult neurogenesis and allows for the direct application of pharmacological agents to the cultured hippocampus.

Here we describe a technique for studying hippocampal postnatal neurogenesis in the rodent brain using the organotypic slice culture technique. This ...

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12.2K Views.  University of Toronto. Here we describe a technique for studying hippocampal postnatal neurogenesis using the organotypic slice culture technique. This method allows for in vitro manipulation of adult neurogenesis and allows for the direct application of pharmacological agents to the cultured hippocampus.