This protocol is based on techniques learned in the laboratory of Francis-Andr&eacute; Wollman at the Institut de Biologie Physico-Chimique, Paris, France. The authors owe a debt of gratitude to Jacqueline Girard-Bascou, who originally introduced D.B.S. to Chlamydomonas genetic techniques. Chlamydomonas research in the authors&rsquo; laboratory is supported by National Science Foundation award MCB-0646350.

Part 1: Preparation of strains for gametogenesis
Note: All manipulations should be carried out at room temperature. Higher or lower temperatures will affect the speed of zygote germination and may adversely affect viability. Important consideration: It is essential that the strains to be mated are growing robustly and are free of contaminating fungi or bacteria. If the strains have not been transferred regularly, passage on rich medium should precede generation of gametes.
<ol>
<li>Strains to be mated are transferred onto TAP plates (100&times;15 mm), 1-2 strains per plate. The cells should be grown in a concentrated strip about 1 cm in width.</li>
<li>Incubation on TAP plates should continue until a thick layer of robustly-growing cells is present, with intermediate transfers if necessary. Strains that have not been passaged recently may require one passage and a total of 1 week, whereas wild-type strains in regular use may only take a few days and no passages. Mutants that are light-sensitive may require additional time.</li>
<li>Transfer all the cells from the TAP plates to N10 plates, which are deficient for nitrogen and will begin to induce gametogenesis. Transfer the cells as a thick slab, much as they were present on the TAP plate. This will facilitate subsequent collection of the cells. After 3 days on N10, the cells are prepared to undergo gametogenesis. Cell division will be slow on N10, however cells should remain dark green. Any contaminated strains should be discarded as viable zygotes will not be obtained.</li>
</ol>
Part 2: Mating
<ol>
<li>Each strain is transferred into a 50 ml erlenmeyer flask and resuspended in approximately 2.5 ml sterile distilled water. This can be done either using a wire loop or a sterile stick; what is important is that the cells are well suspended and not in clumps. The exact volume of water should be adjusted so that the cell concentration, as gauged by eye, is about the same for each strain.</li>
<li>Cells are agitated under strong light (1.260 &mu;E/m2/sec) for 2 hours, during which time the cells will become motile gametes. Motility should be checked under a light microscope. If the gametes are swimming poorly, mating is likely to be inefficient. However, even poorly motile strains will mate a reduced frequency.</li>
<li>Next, equal volumes of mt+ and mt- gametes are combined into a single 50 ml flask. Gametes are left under strong light (1.260 &mu;E/m2/sec) without agitation.</li>
<li>Mating can be checked immediately under the light microscope. Pairs of gametes form rapidly, and will appear as rapidly moving cells joined at the flagella. If mating appears to be extremely rapid, samples should be taken within 30 min -1 hr for generation of zygotes. If not, mating can be checked each hour, although this is optional. Since times giving the optimal density of zygotes will vary widely between crosses, it is best to take aliquots of the mating mixture at different times after combining gametes, e.g. after 1 hr, 2 hrs, 3 hrs and 4 hrs. This is done by transferring 300 &micro;L samples to TAP-3% Difco agar 60&times;15 mm plates; 3-4 spots per plate. Be careful not to agitate the mating mixture as this may disrupt mating pairs. Difco agar is used because it has fewer contaminants than lower-priced agars, and it will be easier to see zygotes under the microscope. The unused mating mixture can be left in the flask overnight, and the quality of the mating reaction will be obvious the next day. When mating is efficient, the zygotes will adhere to the wall or bottom of the flask and the medium will appear clear. Even agitation of the flask will fail to dislodge the zygotes. If nothing sticks to the glass when the mixture is agitated with reasonable vigor, then mating was inefficient or did not take place.</li>
<li>The TAP-Difco plates containing mating mixture are dried in a sterile hood, with the lids ajar, until any surface liquid has been absorbed. The plates are typically left in the light (700 nE/m2/sec) for 18 hours, then wrapped in foil, since zygospore maturation will only take place in the dark. The strains crossed, mating time and date are written on both the plates and the foil. Plates are stored 5 to 7 days and can be kept longer, although zygote germination efficiency declines progressively.</li>
</ol>
Part 3: Tetrad Dissection
<ol>
<li>Vegetative cells should be scraped off dried spots of mating mixture using a dull blade, such as a scalpel. The blade can be used to expose an area of zygotes, which will stick tightly to the 3% agar. This step is usually done in the afternoon, so that the zygotes have germinated by the next morning, but have not undergone a second, vegetative division which would yield 8 progeny. Chlamydomonas biologists quickly learn to recognize zygotes under the microscope, since they are larger than vegetative cells, and feature a thick zygospore wall which gives a black outline. Zygospores often appear yellow but may also be somewhat green. The most important character is that they stick to agar, while vegetative cells are readily moved aside. However, if the plates are overly dry this will not be the case. Also, if the mating mixture was too dense, zygotes may form a mat which will make transfer of individual zygotes difficult or impossible. Appropriate densities are established in step 2.1, and will be quickly learned with experience.</li>
<li>Glass needles are prepared for gathering zygotes and separating tetrad progeny. These are prepared by pulling 3 mm glass rods in a small flame to generate a long, thin thread, which can be broken off. The tips of these tapered ends are smoothed or bent into hooks, by brief heating in a low flame (e.g. alcohol or a burning match). Prepare 3 or more glass needles, since they break fairly easily, especially for inexperienced users.</li>
<li>Zygotes are gathered on the 3% TAP-Difco plate using a glass needle into a small pile. A square of agar (up to 0.5 cm2) containing the zygotes is formed and excised using a dull scalpel. Then, the agar block is placed face down on a 15 mm 1.5% TAP-Difco plate, and slid along a horizontal line about 1.5 cm from the top of the plate, to distribute the zygotes. The top horizontal line, and a grid with approximately 1.5 cm horizontal and 0.5 cm vertical spacing are etched into the back of the plates. The grid will be used the following day as a guide for separating the tetrad progeny (the grid can also be etched the day of dissection). Finally, individual zygotes are move to the intersections of the top horizontal lines with each vertical line, with about 20-30 zygotes per plate. Although only 12 or so will be dissected, not every zygote germinates, and some will divide twice before bursting, so extra zygotes are needed.</li>
<li>After distributing the zygotes, and vegetative cells that have been accidentally transferred from the 3% agar plate are killed by exposing the inverted plate for 30 seconds over a glass dish containing a thin (1-2 mm) layer of chloroform. The distance between the plate and chloroform should be about 5 cm. If too much chloroform is used, or if the distance is too small or great, one risks either killing the zygotes, or not killing the vegetative cells. The chloroform treatment must be done immediately after the zygospores have been distributed on the germination plate, to ensure selectivity.</li>
<li>Germination takes 16-20 hours for most strains, but will vary with genotype and with the age of the zygotes. For germination, the plates are placed under low light (239 nE/m2/sec), or in medium light dimmed by covering the plate with a tissue.</li>
<li>Germinated zygotes can be identified under the microscope as swollen or sometimes burst cells with daughter cells inside them. If the zygospore has not ruptured, gently touching with a glass needle will break the wall if germination has occurred. Daughter cells are gently dragged to the grid points below the zygote. It is easiest to do this by capturing the cell in a small amount of liquid medium released from the agar by the pressure of the glass needle, and dragging this &ldquo;puddle&rdquo; behind the needle. One form of &ldquo;insurance&rdquo; that one has dissected a zygote and not a vegetative cell that has twice divided, is that apart from the four daughter cells, the &ldquo;skin&rdquo; of the zygospore wall will remain.</li>
<li>The dissected progeny are grown under appropriate conditions until visible clonies can be picked for subsequent analysis. Plates should be monitored daily for contamination.</li>
</ol>
Note: Generation of vegetative diploids is not covered here, but essentially involves plating the mating mixture on medium which will not support the growth of either parental strain, but will support the growth of the diploid (e.g. two different auxotropic markers). Plates are kept in the light to avoid the formation of zygotes.

<ol>
	<li>Harris, E. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Chlamydomonas+Sourcebook:+Introduction+to+Chlamydomonas+and+Its+Laboratory+Use.">The Chlamydomonas Sourcebook: Introduction to Chlamydomonas and Its Laboratory Use.</a> Conf Proc IEEE Eng Med Biol Soc. , 119-273 (2005).</li></ol>

The authors have nothing to disclose.

Genetic analysis is simple in Chlamydomonas, but it requires particular techniques to generate gametes and separate tetrad progeny. With healthy strains, the techniques are readily acquired. With some mutant strains, it becomes more of an art, and readers should refer Volume 1 of the recently-published Chlamydomonas Sourcebook for historical notes and practical hints. Additionally, it should be noted that there are many variations on the basic theme described above, ranging from the method for inducing ...

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		<th>Type</th>
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		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>CC-125</td>
<td>Mating type plus strain</td>
<td>Chlamydomonas Stock Center</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>CC-124</td>
<td>Mating type minus strain</td>
<td>Chlamydomonas Stock Center</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
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<td>Tris-Acetate-Phosphate (TAP) </td>
<td>Growth medium</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>N10</td>
<td>Growth medium</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
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<td>Difco agar</td>
<td>Reagent</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
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<td>Fisherbrand Stainless-Steel Blades</td>
<td>Tools</td>
<td>Fisher Scientific</td>
<td>08-916-5B</td>
<td>&nbsp;</td>
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<td>BD Surgical Blade Handles</td>
<td>Tools</td>
<td>Fisher Scientific</td>
<td>08-914-5</td>
<td>&nbsp;</td>
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<td>Fisherbrand Metal Scalpels</td>
<td>Tools</td>
<td>Fisher Scientific</td>
<td>08-920A</td>
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mating and tetrad separation of chlamydomonas reinhardtii for genetic analysis

The unicellular green alga Chlamydomonas reinhardtii (Chlamydomonas) has become a popular organism for research in diverse areas of cell biology and genetics because of its simple life cycle, ease of growth and manipulation for genetic analysis, genomic resources, and transformability of the nucleus and both organelles. Mating strains is a common practice when genetic approaches are used in Chlamydomonass, to create vegetative diploids for analysis of dominance, or following tetrad dissection to ascertain nuclear vs. organellar inheritance, to test allelism, to analyze epistasy, or to generate populations for the purpose of map-based cloning. Additionally, genetic crosses are routinely used to combine organellar genotypes with particular nuclear genotypes. Here we demonstrate standard methods for gametogenesis, mating, zygote germination and tetrad separation. This protocol consists of an easy-to-follow series of steps that will make genetic approaches amenable to scientists who are less familiar with Chlamydomonas. Key parameters and trouble spots are explained. Finally, resources for further information and alternative methods are provided.

Watch this Scientific Journal Video about Mating and Tetrad Separation of Chlamydomonas reinhardtii for Genetic Analysis at JoVE.com

Mating and Tetrad Separation of Chlamydomonas reinhardtii for Genetic Analysis

Mating and tetrad separation are required for genetic analysis in Chlamydomonas reinhardtii. Here we demonstrate standard methods for gametogenesis, mating, zygote germination and tetrad dissection. This protocol consists of an easy-to-follow series of steps that will make genetic approaches amenable to scientists who are less familiar with Chlamydomonas.

Mating and Tetrad Separation of Chlamydomonas reinhardtii for Genetic Analysis

The unicellular green alga Chlamydomonas reinhardtii (Chlamydomonas) has become a popular organism for research in diverse areas of cell biology and ...

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19.9K vistas.  Cornell University. Apareamiento y la separación tétrada se requieren para el análisis genético en Chlamydomonas reinhardtii. Aquí nos demuestran los métodos estándar para la gametogénesis, el apareamiento, la germinación de cigoto y la disección tétrada. Este protocolo consiste en una serie fácil de seguir los pasos que harán que los enfoques genéticos susceptibles de científicos que están menos familiarizados con Chlamydomonas.

Video: Mating and Tetrad Separation of Chlamydomonas reinhardtii for Genetic Analysis

Light/dark Transition Test for Mice

Although all of the mouse genome sequences have been determined, we do not yet know the functions of most of these genes. Gene-targeting techniques, however, can be used to delete or manipulate a specific gene in mice. The influence of a given gene on a specific behavior can then be determined by conducting behavioral analyses of the mutant mice. As a test for behavioral phenotyping of mutant mice, the light/dark transition test is one of the most widely used tests to measure anxiety-like behavior in mice. The test is based on the natural aversion of mice to brightly illuminated areas and on their spontaneous exploratory behavior in novel environments. The test is sensitive to anxiolytic drug treatment. The apparatus consists of a dark chamber and a brightly illuminated chamber. Mice are allowed to move freely between the two chambers. The number of entries into the bright chamber and the duration of time spent there are indices of bright-space anxiety in mice. To obtain phenotyping results of a strain of mutant mice that can be readily reproduced and compared with those of other mutants, the behavioral test methods should be as identical as possible between laboratories. The procedural differences that exist between laboratories, however, make it difficult to replicate or compare the results among laboratories. Here, we present our protocol for the light/dark transition test as a movie so that the details of the protocol can be demonstrated. In our laboratory, we have assessed more than 60 strains of mutant mice using the protocol shown in the movie. Those data will be disclosed as a part of a public database that we are now constructing.

Visualization of the protocol will facilitate understanding of the details of the entire experimental procedure, allowing for standardization of the protocols used across laboratories and comparisons of the behavioral phenotypes of various strains of mutant mice assessed using this test.

The light/dark transition test is one of the most widely used tests to measure anxiety-like behavior in mice. Here, we present a movie that shows detailed procedures on how we conduct the test.

Although all of the mouse genome sequences have been determined, we do not yet know the functions of most of these genes. Gene-targeting techniques, ...

Method for Culture of Early Chick Embryos ex vivo (New Culture)

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  the formation and patterning of brain, neural tube, somite and heart primordia. Applications of chick embryo culture include electroporation of DNA or RNA constructs in order to analyze gene function, grafts of growth factor coated beads such as FGFs and BMPs , as well as whole mount in situ hybridization and immunohistochemistry. This video demonstrates the different steps in chick embryo culture; First, the embryo is explanted in saline. Then, the embryo is centered on a glass ring. The membranes surrounding the embryo are lifted along the walls of the ring. The ring is then placed in a culture dish containing a pool of albumine. The culture dish is sealed and placed in a humid chamber, where the embryo is cultured for up to 24 hrs. Finally, the embryo is removed from the ring, fixed and processed for further applications. A troubleshooting guide is also presented.

Method for Culture of Early Chick Embryos ex vivo New Culture

This video demonstrates New culture, a method by which chick embryos are cultured outside the egg for up to 24 hr. This method enables one to study early development (primitive streak to 14 som.), a period corresponding to E7-9 in mouse. Applications of this technique include electroporation, in situ hybridization and immunohistochemistry.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

Double Whole Mount in situ Hybridization of Early Chick Embryos

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  gene expression in brain, neural tube, somite and heart primordia formation. This video demonstrates the different steps in 2-color whole mount in situ hybridization; First, the embryo is dissected from the egg and fixed in paraformaldehyde. Second, the embryo is processed for prehybridization. The embryo is then hybridized with two different probes, one coupled to DIG, and one coupled to FITC.  Following overnight hybridization, the embryo is incubated with DIG coupled antibody. Color reaction for DIG substrate is performed, and the region of interest appears blue. The embryo is then incubated with FITC coupled antibody. The embryo is processed for color reaction with FITC, and the region of interest appears red. Finally, the embryo is fixed and processed for phtograph and sectioning. A troubleshooting guide is also presented.

This video demonstrates 2-color whole mount in situ hybridization, a method by which the spatial and temporal expression pattern of 2 different genes can be visualized in young chick embryos. This method was originally introduced by David Wilkinson, Domingos Henrique, Phil Ingham and David Ish -Horowicz.

Method for Whole Mount Antibody Staining in Chick

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying antibody expression in developing brain, neural tube and somite. This video demonstrates the different steps in whole-mount antibody staining using HRP conjugated secondary antibodies; First, the embryo is dissected from the egg and fixed in paraformaldehyde. Second, endogenous peroxidase is inactivated; The embryo is then exposed to primary antibody. After several washes, the embryo is incubated with secondary antibody conjugated to HRP. Peroxidase activity is revealed using reaction with diaminobenzidine substrate. Finally, the embryo is fixed and processed for photography and sectioning. The advantage of this method over the use of fluorescent antibodies is that embryos can be processed for wax sectioning, thus enabling the study of antigen sites in cross section. This method was originally introduced by Jane Dodd and Tom Jessell 1.

This video demonstrates whole mount immunohistochemistry, a method by which the spatial and temporal expression pattern of an antigen can be visualized in young chick embryos. This method was originally introduced by Jane Dodd and Tom Jessell.

Assay for Neural Induction in the Chick Embryo

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying the formation and initial patterning of the nervous system. This video demonstrates how to graft organizer tissue into a host, a method by which Hensen s node (the organizer in the chick embryo) is grafted to a host competent ectoderm. The organizer graft instructs overlying na ve tissue to adopt a neural fate via neural inducing signals. This mechanism is referred to as neural induction, and constitutes the initial step in the formation of brain and spinal cord in amniotes. This method is essentially used for the characterization of putative neural inducing molecules in chick. This video demonstrates the different steps in the assay for neural induction; First, the donnor embryo is explanted and pinned on a dish. Then, the host embryo is prepared for New culture. The graft is excised and transplanted to the host area pellucida margin. The host is cultured for 18-22 hrs. The assembly is fixed and processed for further applications (e.g. in situ hybridization). This method was originally devised by Waddington 1,2 and Gallera 3,4.

Neural induction is the first step in the formation of the brain. It is a mechanism by which Hensen's node (organizer), instructs adjacent tissue to adopt a neural fate, i.e. to give rise to the nervous system. This video demonstrates an assay for neural induction in chick embryo.

A Neuronal and Astrocyte Co-Culture Assay for High Content Analysis of Neurotoxicity

High Content Analysis (HCA) assays combine cells and detection reagents with automated imaging and powerful image analysis algorithms, allowing measurement of multiple cellular phenotypes within a single assay. In this study, we utilized HCA to develop a novel assay for neurotoxicity. Neurotoxicity assessment represents an important part of drug safety evaluation, as well as being a significant focus of environmental protection efforts. Additionally, neurotoxicity is also a well-accepted in vitro marker of the development of neurodegenerative diseases such as Alzheimer's and Parkinson's diseases. Recently, the application of HCA to neuronal screening has been reported. By labeling neuronal cells with &beta;III-tubulin, HCA assays can provide high-throughput, non-subjective, quantitative measurements of parameters such as neuronal number, neurite count and neurite length, all of which can indicate neurotoxic effects. However, the role of astrocytes remains unexplored in these models. Astrocytes have an integral role in the maintenance of central nervous system (CNS) homeostasis, and are associated with both neuroprotection and neurodegradation when they are activated in response to toxic substances or disease states. GFAP is an intermediate filament protein expressed predominantly in the astrocytes of the CNS. Astrocytic activation (gliosis) leads to the upregulation of GFAP, commonly accompanied by astrocyte proliferation and hypertrophy. This process of reactive gliosis has been proposed as an early marker of damage to the nervous system. The traditional method for GFAP quantitation is by immunoassay. This approach is limited by an inability to provide information on cellular localization, morphology and cell number. We determined that HCA could be used to overcome these limitations and to simultaneously measure multiple features associated with gliosis - changes in GFAP expression, astrocyte hypertrophy, and astrocyte proliferation - within a single assay. In co-culture studies, astrocytes have been shown to protect neurons against several types of toxic insult and to critically influence neuronal survival. Recent studies have suggested that the use of astrocytes in an in vitro neurotoxicity test system may prove more relevant to human CNS structure and function than neuronal cells alone. Accordingly, we have developed an HCA assay for co-culture of neurons and astrocytes, comprised of protocols and validated, target-specific detection reagents for profiling &beta;III-tubulin and glial fibrillary acidic protein (GFAP). This assay enables simultaneous analysis of neurotoxicity, neurite outgrowth, gliosis, neuronal and astrocytic morphology and neuronal and astrocytic development in a wide variety of cellular models, representing a novel, non-subjective, high-throughput assay for neurotoxicity assessment. The assay holds great potential for enhanced detection of neurotoxicity and improved productivity in neuroscience research and drug discovery.

This article describes a novel protocol and reagent set designed for sensitive measurement of neurotoxic effects of compounds and treatments on co-cultures of neurons and astrocytes using high content analysis. Results demonstrate that high content analysis represents an exciting novel technology for neurotoxicity assessment.

High Content Analysis (HCA) assays combine cells and detection reagents with automated imaging and powerful image analysis algorithms, allowing ...

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 ...

Rapid Genetic Analysis of Epithelial-Mesenchymal Signaling During Hair Regeneration

Hair follicle morphogenesis, a complex process requiring interaction between epithelia-derived keratinocytes and the underlying mesenchyme, is an attractive model system to study organ development and tissue-specific signaling. Although hair follicle development is genetically tractable, fast and reproducible analysis of factors essential for this process remains a challenge. Here we describe a procedure to generate targeted overexpression or shRNA-mediated knockdown of factors using lentivirus in a tissue-specific manner. Using a modified version of a hair regeneration model 5, 6, 11, we can achieve robust gain- or loss-of-function analysis in primary mouse keratinocytes or dermal cells to facilitate study of epithelial-mesenchymal signaling pathways that lead to hair follicle morphogenesis. We describe how to isolate fresh primary mouse keratinocytes and dermal cells, which contain dermal papilla cells and their precursors, deliver lentivirus containing either shRNA or cDNA to one of the cell populations, and combine the cells to generate fully formed hair follicles on the backs of nude mice. This approach allows analysis of tissue-specific factors required to generate hair follicles within three weeks and provides a fast and convenient companion to existing genetic models.

Tissue-specific analysis of a hair follicle regeneration model using lentivirus to mediate gain- or loss-of-function.

Hair follicle morphogenesis, a complex process requiring interaction between epithelia-derived keratinocytes and the underlying mesenchyme, is an ...

Alternative Cultures for Human Pluripotent Stem Cell Production, Maintenance, and Genetic Analysis

Human pluripotent stem cells (hPSCs) hold great promise for regenerative medicine and biopharmaceutical applications. Currently, optimal culture and efficient expansion of large amounts of clinical-grade hPSCs are critical issues in hPSC-based therapies. Conventionally, hPSCs are propagated as colonies on both feeder and feeder-free culture systems. However, these methods have several major limitations, including low cell yields and generation of heterogeneously differentiated cells. To improve current hPSC culture methods, we have recently developed a new method, which is based on non-colony type monolayer (NCM) culture of dissociated single cells. Here, we present detailed NCM protocols based on the Rho-associated kinase (ROCK) inhibitor Y-27632. We also provide new information regarding NCM culture with different small molecules such as Y-39983 (ROCK I inhibitor), phenylbenzodioxane (ROCK II inhibitor), and thiazovivin (a novel ROCK inhibitor). We further extend our basic protocol to cultivate hPSCs on defined extracellular proteins such as the laminin isoform 521 (LN-521) without the use of ROCK inhibitors. Moreover, based on NCM, we have demonstrated efficient transfection or transduction of plasmid DNAs, lentiviral particles, and oligonucleotide-based microRNAs into hPSCs in order to genetically modify these cells for molecular analyses and drug discovery. The NCM-based methods overcome the major shortcomings of colony-type culture, and thus may be suitable for producing large amounts of homogeneous hPSCs for future clinical therapies, stem cell research, and drug discovery.

Here, we present human pluripotent stem cell (hPSC) culture protocols, based on non-colony type monolayer (NCM) growth of dissociated single cells. This new method, utilizing Rho-associated kinase inhibitors or the laminin isoform 521 (LN-521), is suitable for producing large amounts of homogeneous hPSCs, genetic manipulation, and drug discovery.

Human pluripotent stem cells (hPSCs) hold great promise for regenerative medicine and biopharmaceutical applications. Currently, optimal culture and ...

DNA Electroporation, Isolation and Imaging of Myofibers

Mature muscle has a unique structure that is amenable to live cell imaging. Herein, we describe the experimental protocol for expressing fluorescently labeled proteins in the flexor digitorum brevis (FDB) muscle. Conditions have been optimized to provide a large number of high quality myofibers expressing the electroporated plasmid while minimizing muscle damage. The method employs fluorescent tags on various proteins. Combining this expression method with high resolution confocal microscopy permits live cell imaging, including imaging after laser-induced damage. Fluorescent dyes combined with imaging of fluorescently-tagged proteins provides information regarding the basic structure of muscle and its response to stimuli.

This protocol utilizes electroporation to introduce and express fluorescently labeled proteins in mouse muscle fibers. Following recovery after electroporation, fibers are isolated. Individual fibers are then imaged using high resolution confocal microscopy to visualize muscle structure.

Mature muscle has a unique structure that is amenable to live cell imaging. Herein, we describe the experimental protocol for expressing fluorescently ...