JLN was supported by MH 68347.

Prior to hippocampal isolation
Before beginning the hippocampal isolation, make sure that all the tools are sterile. Spray down the culture hood with 70% ethanol, and place the tools inside the hood. You will need 6- and 10-cm Petri dishes, sterile poly-L coated glass coverslips, pipettors and tips, disposable pipettes, and an electric pipettor. From this point on, remember to use correct sterile technique.
<ol>
<li>Turn on the water bath and make sure that it is heated up to 37&deg;C. </li>
<li>The following solutions that are stored at 4&deg;C will be needed: 
<ul>
<li>Modified Eagle&rsquo;s Medium (MEM)</li>
<li>Neurobasal </li>
<li>Hank&rsquo;s buffered saline solution (HBSS)</li>
<li>Borate buffer solution</li>
<li>Sodium pyruvate solution</li>
<li>Sterile-filtered 20% glucose solution in distilled water </li>
<li>Make sure to spray all the bottles down with 70% ethanol before placing them in the hood. </li>
</ul>
</li>
<li>Take these frozen solutions out of the freezer and place them in a heated water bath: 
<ul>
<li>A 5 ml aliquot of horse serum</li>
<li>A 1 ml aliquot of B-27 supplement</li>
<li>A 1 ml aliquot of 100X antibiotic (penicillin plus streptomycin)</li>
<li>And a 0.5 ml aliquot of L-glutamine</li>
</ul>
</li>
<li>After the solutions have thawed, take them out of the water bath, spray them down with 70% ethanol, and place them in the hood. Now the plating and Neurobasal medium can be prepared.</li>
<li>After the solutions are prepared, cap the containers tightly and place them in the 37&deg;C bath to warm up while performing the hippocampal isolation.</li>
<li>Also, take an aliquot of the protease trypsin (2.5%) out of the freezer and place it in the water bath. Trypsin will digest the dissected hippocampus, which will be isolated in the next step.</li>
<li>Take a 15ml conical tube, fill it with HBSS, and label it with the treatment group. It is here that isolated hippocampi will be collected in the next step. </li>
</ol>
Hippocampal Isolation
<ol>
<li>To begin the hippocampal isolation, make sure newborn pups are clean and have had their milk bands, placentas, and umbilical cords removed by their mother.</li>
<li>Place the pups in a Petri dish, spray with 70% ethanol to clean, and place them in the hood. Animals are euthanized immediately before culturing.</li>
<li>For further sterilization take one pup from the dish, dip the pup into 70% ethanol, and then into two washes of sterile HBSS.</li>
<li>Remove the head from the body with scissors. Using the same scissors, cut through the skin and the skull.</li>
<li>Using a pair of fine tweezers, peel the skull away from the brain and place the brain into a small Petri dish that contains a small amount of sterile HBSS.</li>
<li>Peel back the cerebral hemispheres. The hippocampus is a small, seahorse-shaped structure in the medial temporal lobe.</li>
<li>Remove the hippocampus and place into 3 ml of HBSS in a 15 ml tube. Repeat these steps with each pup, and place each isolated hippocampus into the 15 ml tube. Now, it is time to dissociate the tissue to single cells. </li>
</ol>
Hippocampal cell dissociation
<ol>
<li>After all hippocampi have been isolated, fill the 15 ml tube to 4.5 ml with HBSS.</li>
<li>Remove the trypsin from the water bath, spray with ethanol, and place in the hood. Add 0.5 ml of trypsin to the tube, and incubate for 15 minutes at 37&deg;C.</li>
<li>In the hood, remove the HBSS/trypsin solution from the tube with a sterile pipette, being careful not to disturb the hippocampi that have settled to the bottom of the tube. Add 5 ml of HBSS to the tube and swirl gently. Incubate at 37&deg;C for 5 minutes.</li>
<li>Repeat this step twice, removing the old HBSS and replacing with fresh solution.</li>
<li>Take an aliquot of DNAse I out of the freezer. Add 0.5 ml of DNase to the hippocampi in 4.5 ml HBSS. The DNAse is added to promote enzyme inactivation.</li>
<li>Triturate the solution (or pipette up and down) until it is homogenous. Be careful not to introduce bubbles into the homogenate.</li>
<li>Determine cell viability with the trypan blue exclusion method.</li>
<li>Take a 10 cm Petri dish containing 6 to 8 coverslips, and add 10 ml of plating medium. Pipette the desired number of cells to the dish containing the coverslips. Swirl gently to disperse the cells, and make sure the coverslips do not overlap.</li>
<li>Allow the cells to attach for 2-4 hours in a humidified 37&deg;C incubator with 5% CO2. After confirming that the cells are viable and have attached, transfer the coverslips to individual dishes containing Neurobasal medium. Place these dishes back into the incubator.</li>
<li>Once a week, replace one-third of the medium with fresh Neurobasal medium and experimental treatments, as needed. Now, the functional properties of these cells in culture are ready to be studied.</li>
</ol>
Fura-2 Calcium Imaging of Hippocampal Neurons
<ol>
<li>After cultured hippocampal neurons have been in vitro for 48 hours (but not earlier), calcium imaging experiments can begin. At this point, these neurons should have begun to extend processes. The optimal age range for calcium imaging is from day in vitro 3 to 7.</li>
<li>Load cultures with Fura-2 AM for 30 minutes at 37&deg;C, after which they can be imaged under a 20X objective using a Xenon arc lamp to excite the calcium indicator dye and a 512 bit ccd camera, which samples every 150ms.</li>
</ol>

<ol>
	<li>Banker, G., Goslin, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Culturing Nerve Cells, 2nd Edition. , (1998).</li></ol>

This protocol was developed as a modification of the seminal work of Gary Banker and Kim Goslin 1. This book is an essential resource for anyone interested in culturing cells -- not only the neuron-enriched cultures described in the current protocol.
The three most critical factors in cell culturing are: sterility, speed, the choice of medium used.
Sterility - a laminar flow/sterile hood, aseptic conditions, and a sterile incubator are ...

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		<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>Antibiotic antimitotic</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>15240062</td>
<td>&nbsp;</td>
<tr>
<td>B-27 Supplement</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>17504044</td>
<td>Serum free</td>
</tr>
<tr>
<td>Boric Acid</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>B6768</td>
<td>&nbsp;</td>
<tr>
<td>Dnase I</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>DN25</td>
<td>&nbsp;</td>
<tr>
<td>Fura-2, AM cell permeant</td>
<td>&nbsp;</td>
<td>Molecular Probes</td>
<td>F1221</td>
<td>&nbsp;</td>
<tr>
<td>Glucose</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>G7528</td>
<td>&nbsp;</td>
<tr>
<td>HBSS (10X)</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>14185052</td>
<td>&nbsp;</td>
<tr>
<td>HEPES</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>15630080</td>
<td>&nbsp;</td>
<tr>
<td>L-Glutamine</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>G7513</td>
<td>&nbsp;</td>
<tr>
<td>MEM</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>51200038</td>
<td>&nbsp;</td>
<tr>
<td>Neurobasal</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>12348017</td>
<td>Without phenol red</td>
</tr>
<tr>
<td>Poly-L-Lysine Hydrobromide</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>P6282</td>
<td>&nbsp;</td>
<tr>
<td>Pyruvic Acid</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>P2256</td>
<td>&nbsp;</td>
<tr>
<td>Sodium Pyruvate</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>P2256</td>
<td>&nbsp;</td>
<tr>
<td>Sodium Tetraborate</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Trypsin, 2.5% (10X)</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>15090046</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

primary culture of hippocampal neurons from p0 newborn rats

The physiological properties of hippocampal neurons are commonly investigated, especially because of the involvement of the hippocampus in learning and memory. Primary hippocampal cell culturing allows neuroscientists to examine the activity and properties of neurons at the individual cell and single synapse level. In this video, we will demonstrate how to isolate and grow primary hippocampal cells from newborn rats. The hippocampus may be isolated from each newborn animal in as short as 2 to 3 minutes, and the cultures can be maintained for up to two weeks. We will also briefly demonstrate how to use these hippocampal neurons for ratiometric calcium imaging. While this protocol describes the process for the hippocampus, with little to no modification, it can be applied to other regions of the brain.

Watch this Scientific Journal Video about Primary Culture of Hippocampal Neurons from P0 Newborn Rats at JoVE.com

Primary Culture of Hippocampal Neurons from P0 Newborn Rats

The dissection and growth of cells from an individual brain area facilitates investigation of cellular and physiological parameters. We describe a method for primary cell culturing that produces neuron-enriched cultures in a serum-free environment.

The physiological properties of hippocampal neurons are commonly investigated, especially because of the involvement of the hippocampus in learning and ...

primary-culture-of-hippocampal-neurons-from-p0-newborn-rats

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76.4K Views.  Michigan State University (MSU). The dissection and growth of cells from an individual brain area facilitates investigation of cellular and physiological parameters. We describe a method for primary cell culturing that produces neuron-enriched cultures in a serum-free environment.

Video: Primary Culture of Hippocampal Neurons from P0 Newborn Rats

Gene-gun Transfection of Hippocampal Neurons

Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  Vital fluorescent probes, particularly green fluorescent protein (GFP) have revolutionized cell biology stemming from the ability to label specific intracellular compartments and cellular structures. For example, the live imaging of GFP (and its spectral variants) chimeras have allowed for a dynamic 
analysis of the cytoskeleton, organelle transport, and membrane dynamics in a multitude of organisms and cell types [1-3].  Although live imaging has become prevalent, this approach still poses many technical challenges, particularly in primary cultured neurons.  One challenge is the expression of GFP-tagged proteins in post-mitotic neurons; the other is the ability to capture fluorescent images while minimizing phototoxicity, photobleaching, 
and maintaining general cell health.  Here we provide a protocol that describes a lipid-based transfection method that yields a relatively low transfection rate (~0.5%), however is ideal for the imaging of fully polarized neurons.  A low transfection rate is essential so that single axons and dendrites can be characterized as to their orientation to the cell body to confirm directionality of transport, i.e., anterograde v. retrograde.  Our approach to imaging 
GFP expressing neurons relies on a standard wide-field fluorescent microscope outfitted with a CCD camera, image capture software, and a heated imaging chamber. We have imaged a wide variety of organelles or structures, for example, dense-core vesicles, mitochondria, growth cones, and actin without any special optics or excitation requirements other than a fluorescent light source. Additionally, spectrally-distinct, fluorescently 
labeled proteins, e.g., GFP and dsRed-tagged proteins, can be visualized near simultaneously to characterize co-transport or other coordinated cellular events.  The imaging approach described here is flexible for a variety of imaging applications and can be adopted by a laboratory for relatively little cost provided a microscope is available.

Live cell imaging is of particular utility when studying the dynamics of organelle trafficking. Here we describe a protocol for live imaging of dense-core vesicles in cultured neurons using wide-field fluorescence microscopy. This protocol is flexible and can be adapted to image other organelles such as mitochondria, endosomes, and peroxisomes.

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  ...

Primary Culture of Adult Rat Heart Myocytes

Cultured primary adult rodent heart cells are an important model system for cardiovascular research. Nevertheless, establishment of robust, viable cultured adult myocytes can be a technically challenging, rate-limiting step for many researchers. Here we described a protocol to obtain a high yield of adult rat heart myocytes that remain viable in culture for several days. The heart is isolated and perfused with collagenase and protease under low Ca2+ conditions to recover single myocytes. Ca2+-tolerant cells are obtained by stepwise increases in extracellular Ca2+ concentration in three subsequent wash steps. Cells are filtered, resuspended in culture medium, and plated on laminin coated slips. Cultured myocytes obtained using this protocol are viable for up to four days and are suitable for most experiments including electrophysiology, biochemistry, imaging and molecular biology.

In this paper, we described a typical way to isolate and culture adult rat heart myocytes. Collagenase and protease are used to digest and isolate single myocytes. Myocytes cultured follow this protocol meet most experiment requirements.

Cultured primary adult rodent heart cells are an important model system for cardiovascular research.  Nevertheless, establishment of robust, viable ...

Dissection of Hippocampal Dentate Gyrus from Adult Mouse

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

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

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

Isolation and Primary Culture of Rat Hepatic Cells

Primary hepatocyte culture is a valuable tool that has been extensively used in basic research of liver function, disease, pathophysiology, pharmacology and other related subjects. The method based on two-step collagenase perfusion for isolation of intact hepatocytes was first introduced by Berry and Friend in 1969 1 and, since then, has undergone many modifications. The most commonly used technique was described by Seglenin 1976 2. Essentially, hepatocytes are dissociated from anesthetized adult rats by a non-recirculating collagenase perfusion through the portal vein. The isolated cells are then filtered through a 100 &mu;m pore size mesh nylon filter, and cultured onto plates. After 4-hour culture, the medium is replaced with serum-containing or serum-free medium, e.g. HepatoZYME-SFM, for additional time to culture. These procedures require surgical and sterile culture steps that can be better demonstrated by video than by text. Here, we document the detailed steps for these procedures by both video and written protocol, which allow consistently in the generation of viable hepatocytes in large numbers.

Primary hepatocytes provide a valuable tool to evaluate biochemical, molecular, and metabolic functions in a physiologically relevant experimental system. We describe a reliable protocol for rat in situ liver perfusion, which consistently generates viable hepatocytes up to 1.0 &times; 108 cells per preparation with cell viability between 88 ~ 96%.

Primary hepatocyte culture is a valuable tool that has been extensively used in basic research of liver function, disease, pathophysiology, pharmacology ...

Isolation and Primary Culture of Mouse Aortic Endothelial Cells

The vascular endothelium is essential to normal vascular homeostasis. Its dysfunction participates in various cardiovascular disorders. The mouse is an important model for cardiovascular disease research. This study demonstrates a simple method to isolate and culture endothelial cells from the mouse aorta without any special equipment. To isolate endothelial cells, the thoracic aorta is quickly removed from the mouse body, and the attached adipose tissue and connective tissue are removed from the aorta. The aorta is cut into 1 mm rings. Each aortic ring is opened and seeded onto a growth factor reduced matrix with the endothelium facing down. The segments are cultured in endothelial cell growth medium for about 4 days. The endothelial sprouting starts as early as day 2. The segments are then removed and the cells are cultured continually until they reach confluence. The endothelial cells are harvested using neutral proteinase and cultured in endothelial cell growth medium for another two passages before being used for experiments. Immunofluorescence staining indicated that after the second passage the majority of cells were double positive for Dil-ac-LDL uptake, Lectin binding, and CD31 staining, the typical characteristics of endothelial cells. It is suggested that cells at the second to third passages are suitable for in vitro and in vivo experiments to study the endothelial biology. Our protocol provides an effective means of identifying specific cellular and molecular mechanisms in endothelial cell physiopathology.

The vascular endothelial cells play a significant role in many important cardiovascular disorders. This article describes a simple method to isolate and expand endothelial cells from the mouse aorta without using any special equipment. Our protocol provides an effective means of identifying mechanisms in endothelial cell physiopathology.

The vascular endothelium is essential to normal vascular homeostasis. Its dysfunction participates in various cardiovascular disorders. The mouse is an ...

Isolation and Culture of Primary Mouse Keratinocytes from Neonatal and Adult Mouse Skin

The keratinocyte (KC) is the predominant cell type in the epidermis, the outermost layer of the skin. Epidermal KCs play a critical role in providing skin defense by forming an intact skin barrier against environmental insults, such as UVB irradiation or pathogens, and also by initiating an inflammatory response upon those insults. Here we describe methods to isolate KCs from neonatal mouse skin and from adult mouse tail skin. We also describe culturing conditions using defined growth supplements (dGS) in comparison to chelexed fetal bovine serum (cFBS). Functionally, we show that both neonatal and adult KCs are highly responsive to high calcium-induced terminal differentiation, tight junction formation and stratification. Additionally, cultured adult KCs are susceptible to UVB-triggered cell death and can release large amounts of TNF upon UVB irradiation. Together, the methods described here will be useful to researchers for the setup of in vitro models to study epidermal biology in the neonatal mouse and/or the adult mouse.

Epidermal keratinocytes form a functional skin barrier and are positioned at the front line of host defense against external environmental insults. Here we describe methods for isolation and primary culture of epidermal keratinocytes from neonatal and adult mouse skin, and induction of terminal differentiation and UVB-triggered inflammatory response from keratinocytes.

The keratinocyte (KC) is the predominant cell type in the epidermis, the outermost layer of the skin. Epidermal KCs play a critical role in providing skin ...

Isolation and Culture of Primary Aortic Endothelial Cells from Miniature Pigs

Xenotransplantation is a promising way to resolve the shortage of human organs for patients with end-stage organ failure, and the pig is considered as a suitable organ source. Immune rejection and coagulation are two major hurdles for the success of xenotransplantation. Vascular endothelial cell (EC) injury and dysfunction are important for the development of the inflammation and coagulation responses in xenotransplantation. Thus, isolation of porcine aortic endothelial cells (pAECs) is necessary for investigating the immune rejection and coagulation responses. Here, we have developed a simple enzymatic approach for the isolation, characterization, and expansion of highly purified pAECs from miniature pigs. First, the miniature pig was anaesthetized with ketamine, and a length of aorta was excised. Second, the endothelial surface of aorta was exposed to 0.005% collagenase IV digestive solution for 15 min. Third, the endothelial surface of the aorta was scraped in only one direction with a cell scraper (&#60;10 times), and was not compressed during the process of scraping. Finally, the isolated pAECs of Day 3, and after passage 1 and passage 2, were identified by flow cytometry with an anti-CD31 antibody. The percentages of CD31-positive cells were 97.4% &#177; 1.2%, 94.4% &#177; 1.1%, and 92.4% &#177; 1.7% (mean &#177; SD), respectively. The concentration of Collagenase IV, the digestive time, the direction, and frequency and intensity of scraping are critical for decreasing fibroblast contamination and obtaining high-purity and a large number of ECs. In conclusion, our enzymatic method is a highly-effctive method for isolating ECs from the miniature pig aorta, and the cells can be expanded in vitro to investigate the immune and coagulation responses in xenotransplantation.

An effective enzymatic method for isolation of primary porcine aortic endothelial cells (pAECs) from miniature pigs is described. The isolated primary pAECs can be used to investigate the immune and coagulation response in xenotransplantation.

Xenotransplantation is a promising way to resolve the shortage of human organs for patients with end-stage organ failure, and the pig is considered as a ...

Isolation and Differentiation of Primary White and Brown Preadipocytes from Newborn Mice

The understanding of the mechanisms underlying adipocyte differentiation and function has greatly benefited from the use of immortalized white preadipocyte cell lines. These cultured cell lines, however, have limitations. They do not fully capture the diverse functional spectrum of the heterogenous adipocyte populations that are now known to exist within white adipose depots. To provide a more physiologically relevant model to study the complexity of white adipose tissue, a protocol has been developed and optimized to enable simultaneous isolation of primary white and brown adipocyte progenitors from newborn mice, their rapid expansion in culture, and their differentiation in vitro into mature, fully functional adipocytes. The primary advantage of isolating primary cells from newborn, rather than adult mice, is that the adipose depots are actively developing and are, therefore, a rich source of proliferating preadipocytes. Primary preadipocytes isolated using this protocol differentiate rapidly upon reaching confluence and become fully mature in 4-5 days, a temporal window that accurately reflects the appearance of developed fat pads in newborn mice. Primary cultures prepared using this strategy can be expanded and studied with high reproducibility, making them suitable for genetic and phenotypic screens and enabling the study of the cell-autonomous adipocyte phenotypes of genetic mouse models. This protocol offers a simple, rapid, and inexpensive approach to study the complexity of adipose tissue in vitro.

This report describes a protocol for the simultaneous isolation of primary brown and white preadipocytes from newborn mice. Isolated cells can be grown in culture and induced to differentiate into fully mature white and brown adipocytes. The method enables genetic, molecular, and functional characterization of primary fat cells in culture.

The understanding of the mechanisms underlying adipocyte differentiation and function has greatly benefited from the use of immortalized white ...

Isolation and Culture of Primary Oral Keratinocytes from the Adult Mouse Palate

For years, most studies involving keratinocytes have been conducted using human and mouse skin epidermal keratinocytes. Recently, oral keratinocytes have attracted attention because of their unique function and characteristics. They maintain the homeostasis of the oral epithelium and serve as resources for applications in regenerative therapies. However, in vitro studies that use oral primary keratinocytes from adult mice have been limited due to the lack of an efficient and well-established culture protocol. Here, oral primary keratinocytes were isolated from the palate tissues of adult mice and cultured in a commercial low-calcium medium supplemented with a chelexed-serum. Under these conditions, keratinocytes were maintained in a proliferative or stem cell-like state, and their differentiation was inhibited even after increased passages. Marker expression analysis showed that the cultured oral keratinocytes expressed the basal cell markers p63, K14, and &#945;6-integrin and were negative for the differentiation marker K13 and the fibroblast marker PDGFR&#945;. This method produced viable and culturable cells suitable for downstream applications in the study of oral epithelial stem cell functions in vitro.

The present protocol describes the isolation and culture of oral keratinocytes derived from the adult mouse palate. An evaluation method using immunostaining is also reported.

For years, most studies involving keratinocytes have been conducted using human and mouse skin epidermal keratinocytes. Recently, oral keratinocytes have ...