This research was supported by grants from the Undergraduate Research Opportunities Program at UCI (A.R. and S.P.) and grants from the State of California Alzheimer's Disease Initiative and the National Institutes of Health grant no. HD38466, and Alzheimer's Disease Research Center grant no. AG16573 (J.B.)

1. 	Cryopreservation of Cortical Tissue Blocks
Material Preparation
<ol>
 <li>Prepare 1X Hank's Buffered Salt Solution (HBSS) by diluting at 10X stock solution in sterile water (1:9). Store HBSS at 4oC.</li>
 <li>Prepare 10% DMSO freezing medium by diluting DMSO in HBSS (1:9). The freezing medium should be made fresh before cryopreservation and stored at 4oC until chilled.</li>
 <li>A steel razor blade is used for the systematic chopping of tissue. Before use, the razor blade is sterilized by submersion in 70% ethanol for 2 hr. Immediately before tissue processing, rinse the razor blade with sterile water 3 times. Avoid leaving the razor in sterile water for an excessive amount of time, as it is prone to oxidation.</li>
 <li>A Nalgene freezing container is loaded with cryovials (2.5 mL) and then brought into a sterile biological cabinet. 1 mL of chilled freezing medium is added to each vial. The freezing container is then placed at 4oC for at least 2 hrs.</li>
</ol>
Cleaning, Chopping, and Freezing
<ol>
 <li>The brain tissue to be frozen is cleaned of meningeal membrane and blood vessels. Use sterile needle tips to carefully remove debris from the tissue while working on top of an ice pack. Cleaned tissue should be rinsed lightly with cold HBSS and transferred to a new Petri dish for chopping.</li>
 <li>Working on top of an ice pack, use a sterilized razor blade to rapidly chop the tissue into approximately 1 mm3 blocks. Working with small portions of cleaned tissue at a time results in better control over the chopping procedure, resulting in more uniform block sizes.</li>
 <li>Slowly add chopped tissue into 50 mL of HBSS in a 50 mL conical tube. Gently rinse the Petri dish with HBSS in order to collect any remaining tissue blocks. Allow the tissue blocks to descend to the bottom of the tube, creating a loosely packed tissue block pellet.</li>
 <li>While the tissue is settling, bring the previously chilled freezing container and cryovials into the biological safety cabinet. Uncap all cryovials to hasten the process of adding tissue.</li>
 <li>After all the tissue blocks have settled, gently aspirate the excess HBSS, leaving a very thin layer of media above the pellet. Centrifugation is discouraged as it will cause the tissue blocks to adhere to each other, significantly diminishing freezing efficiency.</li>
 <li>Slowly collect 200 &mu;L from the bottom of the loose tissue block pellet by using pipette tip with wide orifices (cut tip with sterile scissors). Transfer the material to one cryovial and move on to the next, again collecting tissue from the bottom of the pellet. It is essential that the entire tissue allocation procedure does not take more than 3-4 min per freezing container. This ensures that the tissue is exposed to DMSO for only a short period of time before freezing. After transferring the tissue to the vials, place the freezing container in a -80oC freezer for at least 4 hrs. Alternatively, the freezing container can be left overnight at -80 oC. Repeat this process for any remaining freezing containers.</li>
 <li>Transfer the cryovials from the freezing container(s) to a cryobox and place in a liquid nitrogen tank for long-term storage.</li>
</ol>
2. 	Thawing and Culturing of Frozen Cortical Tissue Blocks
Material Preparation
<ol>
 <li>One day prior to thawing tissue samples, poly-L-lysine coated Petri dishes/coverslips are prepared. In general, for cortical neurons we prepare coated dishes with 500 &mu;g/ mL or 1mg/ mL. Add enough poly-L-lysine solution (made in borate buffer) to fully cover the bottom of the dish. If glass coverslips are required, ensure that they are completely submerged under the poly-L-lysine solution. Incubate for at least 12 hr. Before use, thoroughly rinse the dishes with a liberal amount of sterile water 3 times, 5 min each.</li>
 <li>Warm HBSS is used to clean thawed tissue as well as to dissociate the tissue into cell suspensions. The volume of HBSS needed will vary depending on the amount of tissue to be thawed, but typically 1 cryovial will dilute in 50 mL of HBSS and will be dissociated in 10 mL of HBSS.</li>
 <li>Dulbecco's Modified Eagle Medium with 10% iron-supplemented bovine calf serum (EBS) and 1% antibiotic/antimycotic (AA) mixture (DMEM(++)) should be placed in a 37&deg;C water bath.</li>
 <li>Plating medium (NB(+++)) should be made fresh before use. This medium is prepared by adding 1% antibiotic/antimycotic plus B27 and N2 supplements.</li>
</ol>
Note: Media names appended with crosses (+) indicate the inclusion of additives to the base composition of the media. In this text, DMEM(++) denotes DMEM plus 10% EBS plus 1% AA, while NB(+++) denotes NB plus 1% AA plus B27 plus N2.
Thawing and Culturing
<ol>
 <li>Remove the cryovials from the liquid nitrogen tank and rapidly thaw at 37&deg;C in a water bath until only a small ice pellet is observed inside the vial content.</li>
 <li>Gently transfer the vial content to 50mL of warm HBSS in a conical tube. Use a wide orifice tip to gently dislodge any remaining tissue blocks stuck in the cryovial. Invert the tube 3-4 times and allow the tissue to slowly collect at the bottom. Tissue should settle quickly, but if the blocks are too small this may not occur and light centrifugation (~200 x g) is recommended.</li>
 <li>Aspirate excess HBSS.</li>
 <li>Add 10 mL of warm HBSS to the tissue pellet and 300 &mu;L of 0.25% trypsin and 50 &mu;L of DNAase. Incubate in a 37&deg;C water bath for 5 min, then place in an orbital shaker set to 80 rpm and 37&deg;C for an additional 5 min.</li>
 <li>After this period, bring the tissue blocks into a biological safety cabinet and use a 10 mL pipette to carefully agitate the tissue blocks until a cloudy cell suspension is formed. Deactivate the trypsin by adding 10 mL of warm DMEM(++) to the cell suspension.</li>
 <li>Centrifuge the dissociated cells at 1200 x g for 5 min. Aspirate and discard the supernatant, resuspend the cell pellet in 10 mL of warm DMEM(++) and quantify cell number using a hemocytometer.</li>
 <li>Plate cells on poly-L-lysine coated dishes. Typically, a 60-mm dish will be plated with 1x106 cells.</li>
 <li>Allow the cells to attach to the dish/coverslips for approximately 1 hr in the tissue culture incubator. It is recommended to monitor the progress of attachment in one plate every 10 min until effective attachment is observed. Then, gently replace the medium with fresh DMEM(++).</li>
 <li>After 24 hr, replace the DMEM(++) with warm NB(+++). Partial medium changes (50%) are carried out every 5 days with fresh NB(+++). Healthy cultures usually show signs of differentiation and growth of processes after 24 hrs. Under these conditions, and performing partial medium changes every 4-5 days, the cultures can be maintained for prolonged periods of time up to 4-6 weeks.</li>
 <li>For the generation of neuronal precursor cells (NPCs), a similar protocol is used with the exception that after centrifugation, DMEM without EBS is used. Cells are plated in T-25 flasks in DMEM/F12 (1:3) supplemented with B27 and EGF (20 ng/ mL) to allow formation of neuropheres. To differentiate NPCs, neurospheres are plated on laminin (10 ug/ mL) coated coverslips in DMEM/F12 (1:3) medium supplemented with N2.</li>
</ol>
3. Representative Results
Healthy cultures will show significant growth and differentiation after 5 days post-thawing and will usually stabilize in its growth by 10 days (Figure 1). Immuncocytochemical staining of these cultures reveals numerous astrocytes (Figure 2A) and neurons (Figure 2B) for both human and rat primary neuronal cultures. This protocol is also suitable for the generation of NPCs as free-floating neurospheres (Figure 3A), which under differentiating conditions, result in high quality mixed cultures (Figure 3B,C).
<img src="/files/ftp_upload/2384/2384fig1.jpg" alt="Figure 1" /> 
 Figure 1. Cell cultures of frozen human cortical tissue. Frozen human cortical tissue blocks are thawed and plated on poly-lysine coated coverslips and grown for 10 days. By day 5 cell expansion is apparent and by day 10 confluency is achieved. Scale bar: 100 &mu;m.
<img src="/files/ftp_upload/2384/2384fig2.jpg" alt="Figure 2" /> 
 Figure 2. Immunocytochemical staining of cell cultures. (A) Cultures were stained with the glial marker glial fibrillary acidic protein (GFAP, thick arrows). (B) Neurons were detected with the neuronal marker beta-tubulin III (TIII, thin arrows). (C) Both human and rat cultures show abundant glia and neurons after 10 days in culture. Primary antibodies: mouse anti-GFAP (1:1000) and rabbit anti-TIII (1:1000). Secondary antibodies: anti-mouse Alexa 594 (1:500) and anti-rabbit Alexa 488 (1:500). Nuclei staining: Hoestch blue (1:1000). Scale bar: 50 &mu;m.
<img src="/files/ftp_upload/2384/2384fig3.jpg" alt="Figure 3" /> 
 Figure 3. Generation of neurospheres. (A) Frozen tissue was processed as described above and allowed to propagate as neurospheres. (B) Neurospheres were plated on glass coverslips under differentiating conditions and grown for 10 days, resulting in cells migrating away from the neurosphere. (C) Both neurons and astrocytes are present in the expanding fringe of a differentiating neurosphere. Nuclear counterstaining, primary and secondary antibodies used as in Fig 2.

<ol>
	<li>Robbins, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cryopreservation+of+human+brain+tissue.">Cryopreservation of human brain tissue.</a> Exp Neurol. 107, 208-213 (1990).</li><li>Ware, C. B., Nelson, A. M., Blau, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Controlled-rate+freezing+of+human+ES+cells.">Controlled-rate freezing of human ES cells.</a> Biotechniques. 38, 879-880 (2005).</li><li>Paynter, S. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Principles+and+practical+issues+for+cryopreservation+of+nerve+cells.">Principles and practical issues for cryopreservation of nerve cells.</a> Brain Res Bull. 75, 1-14 (2008).</li><li>Thirumala, S., Gimble, J. M., Devireddy, R. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cryopreservation+of+stromal+vascular+fraction+of+adipose+tissue+in+a+serum-free+freezing+medium.">Cryopreservation of stromal vascular fraction of adipose tissue in a serum-free freezing medium.</a> J Tissue Eng Regen Med. 4, 224-232 (2010).</li></ol>

No conflicts of interest declared.

Cryopreservation offers an opportunity to bank precious brain tissue samples for future use. Here we describe a simple but effective protocol to generate both neuron-enriched cultures and neuronal precursor cells from frozen brain tissue blocks. This economical procedure avoids the costs of traditional cryopreservation techniques that utilize more expensive rate-controlled freezers. The protocol allows for the generation of viable neuronal cultures post-thawing by providing a rapid yet effective means to freeze tissue blocks. The entire freezing process can take as little as 20 minutes. In addition to primary neuronal cultures, using this method tissue blocks can also be thawed to generate NPCs grown as free floating neurospheres. In this regard, the lack of serum in our freezing medium ensures that cells are preserved in an undifferentiated state. Under differentiating conditions, neurospheres produced from frozen tissue show expansion and differentiation rates very comparable to neurospheres generated from fresh tissue.

<!DOCTYPE html PUBLIC "-//W3C//DTD XHTML 1.0 Transitional//EN" "http://www.w3.org/TR/xhtml1/DTD/xhtml1-transitional.dtd">
<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Dulbecco's Modified Eagle Medium (DMEM)</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>11995-073</td>
<td>High gluclose 1X</td>
</tr>
<tr>
<td>Bovine calf serum supplemented</td>
<td>&nbsp;</td>
<td>HYCLONE</td>
<td>SH30072.03</td>
<td>For culture medium </td>
</tr>
<tr>
<td>Neurobasal-A Medium (1X), liquid</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>1088-022</td>
<td>&nbsp;</td>
<tr>
<td>B27 Supplement</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>17504-044</td>
<td>Supplement for Neurobasal medium</td>
</tr>
<tr>
<td>N2 Supplement</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>17502-048</td>
<td>Supplement for Neurobasal medium</td>
</tr>
<tr>
<td>Trypsin (10X)</td>
<td>&nbsp;</td>
<td>Cellgro </td>
<td>25-054CI </td>
<td>Tissue dissociation</td>
</tr>
<tr>
<td>Deoxyribonuclease 1 from bovine pancreas (DNase)</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>D4527-30KU</td>
<td>Tissue dissociation</td>
</tr>
<tr>
<td>Hanks Balanced Salt Solution (HBSS) (10X)</td>
<td>&nbsp;</td>
<td>GIBCO</td>
<td>14065</td>
<td>Cleaning tissue, washes, and freezing medium</td>
</tr>
<tr>
<td>Poly-L-Lysine- 500mg</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>P2636</td>
<td>Substrate for adhesion of neuronal cells</td>
</tr>
<tr>
<td>antibiotic-antimycotic (100X), liquid</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>15240-062</td>
<td>To prevent contamination</td>
</tr>
<tr>
<td>Large orifice pipette Tips (1-200 ul)</td>
<td>&nbsp;</td>
<td>Fischer Scientific</td>
<td>02-681-141</td>
<td>To prevent shear stress to cells</td>
</tr>
<tr>
<td>Graduated pipette tips (101-1000 ul)</td>
<td>&nbsp;</td>
<td>USA Scientific</td>
<td>1111-2721</td>
<td>&nbsp;</td>
<tr>
<td>21 G1 precision guide needles</td>
<td>&nbsp;</td>
<td>Beckton Dickinson</td>
<td>305165</td>
<td>To clean tissue</td>
</tr>
<tr>
<td>10 ml pipette </td>
<td>&nbsp;</td>
<td>USA Scientific</td>
<td>1071-0810</td>
<td>Individually wrapped</td>
</tr>
<tr>
<td>50 ml tubes</td>
<td>&nbsp;</td>
<td>USA Scientific</td>
<td>926-9-04</td>
<td>&nbsp;</td>
<tr>
<td>Single edge razor blade</td>
<td>&nbsp;</td>
<td>Smith Brand</td>
<td>67-0238</td>
<td>To chop tissue</td>
</tr>
<tr>
<td>60 x 15 mm polystyrene petri dish </td>
<td>&nbsp;</td>
<td>USA Scientific</td>
<td>8609-0160</td>
<td>For general culture</td>
</tr>
<tr>
<td>100 x 15 mm polystyrene petri dish </td>
<td>&nbsp;</td>
<td>USA Scientific</td>
<td>8609-0010</td>
<td>For cleaning tissue</td>
</tr>
<tr>
<td>Cryogenic box </td>
<td>&nbsp;</td>
<td>NALGENE Labware</td>
<td>5026-1010</td>
<td>&nbsp;</td>
<tr>
<td>Freezing container</td>
<td>&nbsp;</td>
<td>NALGENE Labware</td>
<td>5100-0001</td>
<td>"Mr. Frosty"</td>
</tr>
<tr>
<td>2.0 ml cryogenic vials</td>
<td>&nbsp;</td>
<td>NALGENE Labware</td>
<td>5012-0020</td>
<td>&nbsp;</td>
<tr>
<td>DMSO</td>
<td>&nbsp;</td>
<td>Fischer Scientific</td>
<td>D128-500</td>
<td>For freezing medium</td>
</tr>
<tr>
<td>Ethanol 200 proof</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>E7023</td>
<td>For sterilizing razor blade</td>
</tr>		</tbody>
		</table>
		</div>

cryopreservation of cortical tissue blocks for the generation of highly enriched neuronal cultures

In this study, we outline a standardized protocol for the successful cryopreservation and thawing of cortical brain tissue blocks to generate highly enriched neuronal cultures. For this protocol the freezing medium used is 10% dimethyl sulfoxide (DMSO) diluted in Hank's Buffered Salt Solution (HBSS). Blocks of cortical tissue are transferred to cryovials containing the freezing medium and slowly frozen at -1&deg;C/min in a rate-controlled freezing container. Post-thaw processing and dissociation of frozen tissue blocks consistently produced neuronal-enriched cultures which exhibited rapid neuritic growth during the first 5 days in culture and significant expansion of the neuronal network within 10 days. Immunocytochemical staining with the astrocytic marker glial fibrillary acidic protein (GFAP) and the neuronal marker beta-tubulin class III, revealed high numbers of neurons and astrocytes in the cultures. Generation of neural precursor cell cultures after tissue block dissociation resulted in rapidly expanding neurospheres, which produced large numbers of neurons and astrocytes under differentiating conditions. This simple cryopreservation protocol allows for the rapid, efficient, and inexpensive preservation of cortical brain tissue blocks, which grants increased flexibility for later generation of neuronal, astrocyte, and neuronal precursor cell cultures.

Watch this Scientific Journal Video about Cryopreservation of Cortical Tissue Blocks for the Generation of Highly Enriched Neuronal Cultures at JoVE.com

Cryopreservation of Cortical Tissue Blocks for the Generation of Highly Enriched Neuronal Cultures

Here, we describe a method for efficient cryopreservation and thawing of cortical brain tissue blocks to generate highly enriched neuronal cultures. This simple protocol provides flexibility for later generation of neuronal, astrocyte, and neuronal precursor cell cultures.

In this study, we outline a standardized protocol for the successful cryopreservation and thawing of cortical brain tissue blocks to generate highly ...

cryopreservation-cortical-tissue-blocks-for-generation-highly

Research

JoVE Journal

Neuroscience

13.6K Views.  University of California, Irvine. Here, we describe a method for efficient cryopreservation and thawing of cortical brain tissue blocks to generate highly enriched neuronal cultures. This simple protocol provides flexibility for later generation of neuronal, astrocyte, and neuronal precursor cell cultures.

Video: Cryopreservation of Cortical Tissue Blocks for the Generation of Highly Enriched Neuronal Cultures

Organotypic Cerebellar Cultures: Apoptotic Challenges and Detection

Organotypic cultures of neuronal tissue were first introduced by Hogue in 1947 1,2 and have constituted a major breakthrough in the field of neuroscience. Since then, the technique was developed further and currently there are many different ways to prepare organotypic cultures. The method presented here was adapted from the one described by Stoppini et al. for the preparation of the slices and from Gogolla et al. for the staining procedure 3,4.
A unique feature of this technique is that it allows you to study different parts of the brain such as hippocampus or cerebellum in their original structure, providing a big advantage over dissociated cultures in which all the cellular organization and neuronal networks are disrupted. In the case of the cerebellum it is even more advantageous because it allows the study of Purkinje cells, extremely difficult to obtain as dissociated primary culture. This method can be used to study certain developmental features of the cerebellum in vitro, as well as for electrophysiological and pharmacological experiments in both wild type and mutant mice.
The method described here was designed to study the effect of apoptotic stimuli such as Fas ligand in the developing cerebellum, using TUNEL staining to measure apoptotic cell death. If TUNEL staining is combined with cell type specific markers, such as Calbindin for Purkinje cells, it is possible to evaluate cell death in a cell population specific manner. The Calbindin staining also serves the purpose of evaluating the quality of the cerebellar cultures.

This method describes the generation of organotypic cerebellar cultures and the effect of certain apoptotic stimuli on the viability of different cerebellar cell types.

Organotypic cultures of neuronal tissue were first introduced by Hogue in 1947 1,2 and have constituted a major breakthrough in the field of neuroscience. ...

Generation of Neural Stem Cells from Discarded Human Fetal Cortical Tissue

Neural stem cells (NSCs) reside along the ventricular zone neuroepithelium during the development of the cortical plate. These early progenitors ultimately give rise to intermediate progenitors and later, the various neuronal and glial cell subtypes that form the cerebral cortex. The capacity to generate and expand human NSCs (so called neurospheres) from discarded normal fetal tissue provides a means with which to directly study the functional aspects of normal human NSC development 1-5. This approach can also be directed toward the generation of NSCs from known neurological disorders, thereby affording the opportunity to identify disease processes that alter progenitor proliferation, migration and differentiation 6-9. We have focused on identifying pathological mechanisms in human Down syndrome NSCs that might contribute to the accelerated Alzheimer's disease phenotype 10,11. Neither in vivo nor in vitro mouse models can replicate the identical repertoire of genes located on human chromosome 21. 
Here we use a simple and reliable method to isolate Down syndrome NSCs from aborted human fetal cortices and grow them in culture. The methodology provides specific aspects of harvesting the tissue, dissection with limited anatomical landmarks, cell sorting, plating and passaging of human NSCs. We also provide some basic protocols for inducing differentiation of human NSCs into more selective cell subtypes.

A simple and reliable method on isolation and culture of neural stem cells from discarded human fetal cortical tissue is described. Cultures derived from known human neurological disorders can be used for characterization of pathological cellular and molecular processes, as well as provide a platform to assess pharmacological efficacy.

Neural stem cells (NSCs) reside along the ventricular zone neuroepithelium during the development of the cortical plate. These early progenitors ...

Multi-electrode Array Recordings of Neuronal Avalanches in Organotypic Cultures

The cortex is spontaneously active, even in the absence of any particular input or motor output. During development, this activity is important for the migration and differentiation of cortex cell types and the formation of neuronal connections1. In the mature animal, ongoing activity reflects the past and the present state of an animal into which sensory stimuli are seamlessly integrated to compute future actions. Thus, a clear understanding of the organization of ongoing i.e. spontaneous activity is a prerequisite to understand cortex function. 
Numerous recording techniques revealed that ongoing activity in cortex is comprised of many neurons whose individual activities transiently sum to larger events that can be detected in the local field potential (LFP) with extracellular microelectrodes, or in the electroencephalogram (EEG), the magnetoencephalogram (MEG), and the BOLD signal from functional magnetic resonance imaging (fMRI). The LFP is currently the method of choice when studying neuronal population activity with high temporal and spatial resolution at the mesoscopic scale (several thousands of neurons). At the extracellular microelectrode, locally synchronized activities of spatially neighbored neurons result in rapid deflections in the LFP up to several hundreds of microvolts. When using an array of microelectrodes, the organizations of such deflections can be conveniently monitored in space and time. 
Neuronal avalanches describe the scale-invariant spatiotemporal organization of ongoing neuronal activity in the brain2,3. They are specific to the superficial layers of cortex as established in vitro4,5, in vivo in the anesthetized rat 6, and in the awake monkey7. Importantly, both theoretical and empirical studies2,8-10 suggest that neuronal avalanches indicate an exquisitely balanced critical state dynamics of cortex that optimizes information transfer and information processing.
In order to study the mechanisms of neuronal avalanche development, maintenance, and regulation, in vitro preparations are highly beneficial, as they allow for stable recordings of avalanche activity under precisely controlled conditions. The current protocol describes how to study neuronal avalanches in vitro by taking advantage of superficial layer development in organotypic cortex cultures, i.e. slice cultures, grown on planar, integrated microelectrode arrays (MEA; see also 11-14).

A robust way to study neuronal avalanches, i.e. scale-invariant spatio-temporal activity bursts, indicative of critical state dynamics in cortex. Avalanches emerge spontaneously in developing superficial layers of cultured cortex which allows for long-term measurements of the activity with planar integrated multi-electrode arrays (MEA) under precisely controlled conditions.

The cortex is spontaneously active, even in the absence of any particular input or motor output.  During development, this activity is important for the ...

Derivation of Enriched Oligodendrocyte Cultures and Oligodendrocyte/Neuron Myelinating Co-cultures from Post-natal Murine Tissues

Identifying the molecular mechanisms underlying OL development is not only critical to furthering our knowledge of OL biology, but also has implications for understanding the pathogenesis of demyelinating diseases such as Multiple Sclerosis (MS). Cellular development is commonly studied with primary cell culture models. Primary cell culture facilitates the evaluation of a given cell type by providing a controlled environment, free of the extraneous variables that are present in vivo. While OL cultures derived from rats have provided a vast amount of insight into OL biology, similar efforts at establishing OL cultures from mice has been met with major obstacles. Developing methods to culture murine primary OLs is imperative in order to take advantage of the available transgenic mouse lines. 
Multiple methods for extraction of OPCs from rodent tissue have been described, ranging from neurosphere derivation, differential adhesion purification and immunopurification 1-3. While many methods offer success, most require extensive culture times and/or costly equipment/reagents. To circumvent this, purifying OPCs from murine tissue with an adaptation of the method originally described by McCarthy &amp; 
de Vellis 2 is preferred. This method involves physically separating OPCs from a mixed glial culture derived from neonatal rodent cortices. The result is a purified OPC population that can be differentiated into an OL-enriched culture. This approach is appealing due to its relatively short culture time and the unnecessary requirement for growth factors or immunopanning antibodies. 
While exploring the mechanisms of OL development in a purified culture is informative, it does not provide the most physiologically relevant environment for assessing myelin sheath formation. Co-culturing OLs with neurons would lend insight into the molecular underpinnings regulating OL-mediated myelination of axons. For many OL/neuron co-culture studies, dorsal root ganglion neurons (DRGNs) have proven to be the neuron type of choice. They are ideal for co-culture with OLs due to their ease of extraction, minimal amount of contaminating cells, and formation of dense neurite beds. While studies using rat/mouse myelinating xenocultures have been published 4-6, a method for the derivation of such OL/DRGN myelinating co-cultures from post-natal murine tissue has not been described. Here we present detailed methods on how to effectively produce such cultures, along with examples of expected results. These methods are useful for addressing questions relevant to OL development/myelinating function, and are useful tools in the field of neuroscience.

This article describes methods to derive enriched populations of murine oligodendrocyte precursor cells (OPCs) in primary culture, which differentiate to produce mature oligodendrocytes (OLs). In addition, this report describes techniques to produce murine myelinating co-cultures by seeding mouse OPCs onto a neurite bed of mouse dorsal root ganglion neurons (DRGNs).

Identifying the molecular mechanisms underlying OL development is not only critical to furthering our knowledge of OL biology, but also has implications ...

A Fully Automated and Highly Versatile System for Testing Multi-cognitive Functions and Recording Neuronal Activities in Rodents

We have developed a fully automated system for operant behavior testing and neuronal activity recording by which multiple cognitive brain functions can be investigated in a single task sequence. The unique feature of this system is a custom-made, acoustically transparent chamber that eliminates many of the issues associated with auditory cue control in most commercially available chambers. The ease with which operant devices can be added or replaced makes this system quite versatile, allowing for the implementation of a variety of auditory, visual, and olfactory behavioral tasks. Automation of the system allows fine temporal (10 ms) control and precise time-stamping of each event in a predesigned behavioral sequence. When combined with a multi-channel electrophysiology recording system, multiple cognitive brain functions, such as motivation, attention, decision-making, patience, and rewards, can be examined sequentially or independently.

In this report, we present a fully automated and highly versatile system capable of simultaneously testing multi-cognitive behaviors and recording neuronal activities for rodents.

We have developed a fully automated system for operant behavior testing and neuronal activity recording by which multiple cognitive brain functions can be ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

A Procedure for Implanting Organized Arrays of Microwires for Single-unit Recordings in Awake, Behaving Animals

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell level. The technique allows experimenters to examine temporally and regionally specific firing patterns in order to correlate recorded action potentials with ongoing behavior. Moreover, single-unit recordings can be combined with a plethora of other techniques in order to produce comprehensive explanations of neural function. In this article, we describe the anesthesia and preparation for microwire implantation. Subsequently, we enumerate the necessary equipment and surgical steps to accurately insert a microwire array into a target structure. Lastly, we briefly describe the equipment used to record from each individual electrode in the array. The fixed microwire arrays described are well-suited for chronic implantation and allow for longitudinal recordings of neural data in almost any behavioral preparation. We discuss tracing electrode tracks to triangulate microwire positions as well as ways to combine microwire implantation with immunohistochemical techniques in order to increase the anatomical specificity of recorded results.

Implanting organized arrays of microwires for use in single-unit electrophysiological recordings presents a number of technical challenges.&#160;Methods for performing this technique and the equipment necessary are described.&#160;Also, the beneficial use of organized microwire arrays to record from distinct neural subregions with high spatial selectivity is discussed.

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell ...

Preparation of Neuronal Co-cultures with Single Cell Precision

Microfluidic embodiments of the Campenot chamber have attracted great interest from the neuroscience community. These interconnected co-culture platforms can be used to investigate a variety of questions, spanning developmental and functional neurobiology to infection and disease propagation. However, conventional systems require significant cellular inputs (many thousands per compartment), inadequate for studying low abundance cells, such as primary dopaminergic substantia nigra, spiral ganglia, and Drosophilia melanogaster neurons, and impractical for high throughput experimentation. The dense cultures are also highly locally entangled, with few outgrowths (&#60;10%) interconnecting the two cultures. In this paper straightforward microfluidic and patterning protocols are described which address these challenges: (i) a microfluidic single neuron arraying method, and (ii) a water masking method for plasma patterning biomaterial coatings to register neurons and promote outgrowth between compartments. Minimalistic neuronal co-cultures were prepared with high-level (&#62;85%) intercompartment connectivity and can be used for high throughput neurobiology experiments with single cell precision.

Protocols for single neuron microfluidic arraying and water masking for the in-chip plasma patterning of biomaterial coatings are described. Highly interconnected co-cultures can be prepared using minimal cell inputs.

Microfluidic embodiments of the Campenot chamber have attracted great interest from the neuroscience community. These interconnected co-culture platforms ...

Prolonged Incubation of Acute Neuronal Tissue for Electrophysiology and Calcium-imaging

Acute neuronal tissue preparations, brain slices and retinal wholemount, can usually only be maintained for 6 - 8 h following dissection. This limits the experimental time, and increases the number of animals that are utilized per study. This limitation specifically impacts protocols such as calcium imaging that require prolonged pre-incubation with bath-applied dyes. Exponential bacterial growth within 3 - 4 h after slicing is tightly correlated with a decrease in tissue health. This study describes a method for&#160;limiting the proliferation of bacteria in acute preparations to maintain viable neuronal tissue for prolonged periods of time (&#62;24 h) without the need for antibiotics, sterile procedures, or tissue culture media containing growth factors. By cycling the extracellular fluid through UV irradiation and keeping the tissue in a custom holding chamber at 15 - 16 &#176;C, the tissue shows no difference in electrophysiological properties, or calcium signaling through intracellular calcium dyes at &#62;24 h postdissection. These methods will not only extend experimental time for those using acute neuronal tissue, but will reduce the number of animals required to complete experimental goals, and will set a gold standard for acute neuronal tissue incubation.

Once removed from the body, neuronal tissue is greatly affected by environmental conditions, leading to eventual degradation of the tissue after 6 - 8 h. Using a unique incubation method, which closely monitors and regulates the extracellular environment of the tissue, tissue viability can be significantly extended for &#62;24 h.

Acute neuronal tissue preparations, brain slices and retinal wholemount, can usually only be maintained for 6 - 8 h following dissection. This limits the ...

Time-dependent Increase in the Network Response to the Stimulation of Neuronal Cell Cultures on Micro-electrode Arrays

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network dynamics in neuronal cultures. In contrast with more traditional methods, such as patch-clamping, which can only record activity from a single cell, MEAs can record simultaneously from multiple sites in a network, without requiring the arduous task of placing each electrode individually. Moreover, numerous control and stimulation configurations can be easily applied within the same experimental setup, allowing for a broad range of dynamics to be explored. One of the key dynamics of interest in these in vitro studies has been the extent to which cultured networks display properties indicative of learning. Mouse neuronal cells cultured on MEAs display an increase in response following training induced by electrical stimulation. This protocol demonstrates how to culture neuronal cells on MEAs; successfully record from over 95% of the plated dishes; establish a protocol to train the networks to respond to patterns of stimulation; and sort, plot, and interpret the results from such experiments. The use of a proprietary system for stimulating and recording neuronal cultures is demonstrated. Software packages are also used to sort neuronal units. A custom-designed graphical user interface is used to visualize post-stimulus time histograms, inter-burst intervals, and burst duration, as well as to compare the cellular response to stimulation before and after a training protocol. Finally, representative results and future directions of this research effort are discussed.

Mouse neuronal cells cultured on multi-electrode arrays display an increase in response following electrical stimulation. This protocol demonstrates how to culture neurons, how to record activity, and how to establish a protocol to train the networks to respond to patterns of stimulation.

Micro-electrode arrays (MEAs) can be used to investigate drug toxicity, design paradigms for next-generation personalized medicine, and study network ...