The authors would like to thank Irene Choi for illustrating embryo heads in Fig 1 and the other members of the laboratory for help and discussion. This work is supported by NIH grant DE012728 (T.W.)

1. Prepare Dispase II
<ol>
	<li>Prepare fresh Hepes-buffered Saline (HBS) (50 mM Hepes/KOH pH 7.4, 150 mM NaCl). Add 2 ml 0.5 M Hepes/KOH pH 7.4 and 1.2 ml 2.5 M NaCl into 16.8 ml Ultrapure H2O.</li>
	<li>Make 10 mg/ml Dispase II. Weigh 0.2 g Dispase II (Roche, Cat# 4942078001) and dissolve in 20 ml HBS. Prepare 1.0 ml aliquots in individual Eppendorf tubes and store at -20 &deg;C for long-term storage.</li>
	<li>Dilute 10 mg/ml Dispase II 1:10 in PBS before use. Put the diluted Dispase II on ice.</li>
</ol>
2. Dissect CD-1 E10.5 Embryos from Pregnant Female
All animal experiments were performed in accordance with protocols approved by the University of Colorado Denver (UCD) Animal Care and Usage Committee. CD-1 (ICR) mice were obtained from Harlan Laboratories (Indianapolis, IN).
<ol>
	<li>Clean the dissection area and tools with 70% ethanol.</li>
	<li>Euthanize pregnant CD-1 female using approved protocols.</li>
	<li>Spray the abdomen with 70% ethanol then open the abdomen. Use one set of forceps to pull the uterus and another to tear away the mesometrium from the uterus. Remove the uterus from the body. Repeat for uterine horn on other side of the abdomen.</li>
	<li>Briefly rinse the uterus with ice cold PBS in a 10-cm Petri dish to wash out blood.</li>
	<li>Transfer the uterus into a new Petri dish with ice cold PBS. Separate into one-embryo fragments by cutting between implantation sites.</li>
	<li>Transfer one embryo into ice-cold PBS under a stereomicroscope. Use fine forceps to tear off the muscular wall of uterus starting at one of the cut sites to expose the embryo with the yolk sac. Peel off the yolk sac and amnion. Transfer dissected embryo to a 6-cm Petri dish with PBS on ice. Proceed until all embryos have been collected in this way.</li>
</ol>
3. Dispase II Treatment
<ol>
	<li>Wash all the embryos with PBS in a 6-cm Petri dish. Replace PBS with 10 ml diluted Dispase II in PBS. Cover the Petri dish and incubate at 37 &deg;C for 25 min - a bacterial incubator or warm room should suffice - we have not used a CO2 tissue culture incubator for this step.</li>
	<li>Take the Petri dish out and observe the embryo under a stereomicroscope. The ectoderm should have become loose, typified by a clear gap between this tissue layer and the underlying mesenchyme. If not, incubate for an additional 5 min at 37 &deg;C. Then put the Petri dish on ice.</li>
</ol>
4. Dissect out the Intact Facial Prominences
<ol>
	<li>Use a disposable glass pipette (VWR, Cat# 14672-380) to transfer one Dispase II treated embryo into ice-cold PBS in a 6-cm Petri dish under a stereomicroscope.</li>
	<li>Use one pair of forceps to hold the embryo. Use another pair of fine forceps carefully to cut the boundaries of the facial prominences attached to head (Figure 1 shows the boundaries of the facial prominences). Start from one side of the MnP, then the MxP, and last the FNP. Do the same to the other side to dissect the facial prominences out intact (Figure 2A).</li>
</ol>
5. Separate Facial Ectoderm from Mesenchyme
<ol>
	<li>Observe the facial prominences under a stereomicroscope with 3x working distance. Use a long glass Pasteur pipette to gently pipette the facial prominences up and down 5 times. After this, the ectoderm is easy to peel off.</li>
	<li>Use one pair of forceps to hold the facial prominences. Use the other pair of forceps to peel off the ectoderm very slowly. Figure 2B shows the separated sheets of facial ectoderm.</li>
	<li>Use a new long glass Pasteur pipette to transfer the facial ectoderm into a 1.5 ml Eppendorf tube with PBS on ice. Transfer the mesenchyme to another 1.5 ml Eppendorf tube with PBS on ice.</li>
	<li>Use new ice cold PBS for each dissection. Collect samples.</li>
	<li>Wash the samples with PBS. Centrifuge at 4 &deg;C, 500 x g for 3 min.</li>
	<li>Aspirate the PBS using long glass Pasteur pipettes. If the samples are to be used for ChIP assay, proceed to step 6.1. For RNA extraction, proceed to step 6.2. For Protein extraction, proceed to step 6.3.</li>
</ol>
6. Process Samples for Further Experimentation
<ol>
	<li>Samples for ChIP assay
	<ol>
		<li>Add 1 ml PBS with 1% formaldehyde to crosslink. Homogenize the samples.</li>
		<li>Incubate at room temperature for 10 min with rotation.</li>
		<li>Add 110 &mu;l of 1.25 M Glycine to each tube to quench unreacted formaldehyde.</li>
		<li>Mix and incubate at room temperature for 5 min.</li>
		<li>Spin at 500 x g at 4 &deg;C for 3 min. Remove supernatant.</li>
		<li>Add 1 ml cold PBS to wash the tissues.</li>
		<li>Spin at 500 x g at 4 &deg;C for 3 min.</li>
		<li>Remove PBS and repeat PBS wash once.</li>
		<li>Remove supernatant, Snap freeze the tissues in liquid nitrogen. Store the samples at -80 &deg;C for subsequent pooling and processing.</li>
	</ol>
	</li>
	<li>Samples for RNA extraction
	<ol>
		<li>Place the facial ectoderm and mesenchyme in tubes containing RNAlater (Ambion).</li>
		<li>Incubate the samples in RNAlater Solution overnight at 4 &deg;C to allow thorough penetration of the tissue.</li>
		<li>Store at -20 &deg;C for subsequent pooling and processing.</li>
	</ol>
	</li>
	<li>Samples for protein extraction</li>
</ol>
Add lysis buffer to the sample for subsequent processing. Or snap freeze the tissues in liquid nitrogen. Store the samples at -80 &deg;C for subsequent pooling and processing.
Notes
<ol>
	<li>Time constraints. Plan on 4-6 hr from the start of dissection through to the beginning of section 6.</li>
	<li>When proficient, it takes ~10-15 min per embryo to perform sections 4 and 5 (prominence dissection and ectoderm separation).</li>
	<li>If the samples are for RNA isolation, every solution should be treated with Diethylpyrocarbonate first. Also clean the forceps and the working area with RNase Away.</li>
	<li>Keep the samples on ice at all times except for the Dispase II treatment and dissections. Change to new ice cold PBS for each dissection to make sure the PBS used for dissection is cold.</li>
	<li>Sharp forceps are critical to obtain precise dissection. Clean the forceps completely to prevent any contamination.</li>
	<li>For collecting samples from different prominences, dissect each prominence separately after Dispase II treatment. Then separate the ectoderm and mesenchyme.</li>
	<li>After treatment with Dispase II, the facial ectoderm is easy to peel off. Pay special attention to contamination from other ectoderm. Discard all potentially contaminated tissue.</li>
	<li>Once you start, don't stop until the samples reach the appropriate point in Section 6. Finish separation of facial ectoderm and mesenchyme for all the embryos as quickly as you can.</li>
	<li>The enzyme activities of Dispase II might be different between batches and brands. It is necessary to adjust the treatment time for different brands and batches of Dispase II.</li>
</ol>

<ol>
	<li>Wilkie, A. O., Morriss-Kay, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetics+of+craniofacial+development+and+malformation.">Genetics of craniofacial development and malformation.</a> Nat. Rev. Genet. 2, 458-468 (2001).</li><li>Gritli-Linde, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+2+-+The+Etiopathogenesis+of+cleft+lip+and+cleft+palate:+usefulness+and+caveats+of+mouse+models.">Chapter 2 - The Etiopathogenesis of cleft lip and cleft palate: usefulness and caveats of mouse models.</a> Current Topics in Developmental Biology. 84, 37-138 (2008).</li><li>Schutte, B. C., Murray, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+many+faces+and+factors+of+orofacial+clefts.">The many faces and factors of orofacial clefts.</a> Human Molecular Genetics. 8, 1-7 (1999).</li><li>Chai, Y., Maxson, R. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Recent+advances+in+craniofacial+morphogenesis.">Recent advances in craniofacial morphogenesis.</a> Dev. Dyn. 235, 2353-2375 (2006).</li><li>Jiang, R., Bush, J. O., Lidral, A. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+the+upper+lip:+Morphogenetic+and+molecular+mechanisms.">Development of the upper lip: Morphogenetic and molecular mechanisms.</a> Dev. Dyn. 235, 1152-1166 (2006).</li><li>Reid, B. S., Yang, H., Melvin, V. S., Taketo, M. M., Williams, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ectodermal+WNT/&#946;-catenin+signaling+shapes+the+mouse+face.">Ectodermal WNT/&#946;-catenin signaling shapes the mouse face.</a> Developmental Biology. 349, 261-269 (2011).</li><li>Feng, W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatial+and+temporal+analysis+of+gene+expression+during+growth+and+fusion+of+the+mouse+facial+prominences.">Spatial and temporal analysis of gene expression during growth and fusion of the mouse facial prominences.</a> PLoS ONE. 4, e8066 (2009).</li><li>Chia, R., Achilli, F., Festing, M. F., Fisher, E. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+origins+and+uses+of+mouse+outbred+stocks.">The origins and uses of mouse outbred stocks.</a> Nat. Genet. 37 (11), 1181-1186 (2005).</li><li>Nagai, T., Aruga, J., Takada, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+expression+of+the+mouse+Zic1,+Zic2,+and+Zic3+gene+suggests+an+essential+role+for+Zic+genes+in+body+pattern+formation.">The expression of the mouse Zic1, Zic2, and Zic3 gene suggests an essential role for Zic genes in body pattern formation.</a> Developmental Biology. 182, 299-313 (1997).</li><li>Britsch, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+transcription+factor+Sox10+is+a+key+regulator+of+peripheral+glial+development.">The transcription factor Sox10 is a key regulator of peripheral glial development.</a> Genes Dev. 15 (1), 66-78 (2001).</li></ol>

The authors have no competing interests or other conflicts associated with the contents of this article.

This protocol provides a straightforward method to separate embryonic mouse facial ectoderm and mesenchyme based on an initial Dispase II treatment step. In previous studies, we have performed dissection of the facial prominences prior to Dispase II treatment, but we have consistently found that the mesenchyme becomes "sticky" and more difficult to manipulate. Our new protocol avoids this problem. Combination Dispase II treatment with gentle physical force by pipetting up and down makes separation of ectoderm and mesench...

<table>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <td>Dispase II (neutral protease, grade II)</td>
 <td>Roche</td>
 <td>4942078001</td>
 <td>Make 10 mg/ml Dispase II stock solution in HBS. Aliquot and store at -20 &deg;C . </td>
 </tr>
 <td>RNAlater</td>
 <td>Ambion</td>
 <td>AM7021</td>
 <td>&nbsp;</td>
 </tr>
</table>

separation of mouse embryonic facial ectoderm and mesenchyme

Orofacial clefts are the most frequent craniofacial defects, which affect 1.5 in 1,000 newborns worldwide1,2. Orofacial clefting is caused by abnormal facial development3. In human and mouse, initial growth and patterning of the face relies on several small buds of tissue, the facial prominences4,5. The face is derived from six main prominences: paired frontal nasal processes (FNP), maxillary prominences (MxP) and mandibular prominences (MdP). These prominences consist of swellings of mesenchyme that are encased in an overlying epithelium. Studies in multiple species have shown that signaling crosstalk between facial ectoderm and mesenchyme is critical for shaping the face6. Yet, mechanistic details concerning the genes involved in these signaling relays are lacking. One way to gain a comprehensive understanding of gene expression, transcription factor binding, and chromatin marks associated with the developing facial ectoderm and mesenchyme is to isolate and characterize the separated tissue compartments.
Here we present a method for separating facial ectoderm and mesenchyme at embryonic day (E) 10.5, a critical developmental stage in mouse facial formation that precedes fusion of the prominences. Our method is adapted from the approach we have previously used for dissecting facial prominences7. In this earlier study we had employed inbred C57BL/6 mice as this strain has become a standard for genetics, genomics and facial morphology8. Here, though, due to the more limited quantities of tissue available, we have utilized the outbred CD-1 strain that is cheaper to purchase, more robust for husbandry, and tending to produce more embryos (12-18) per litter than any inbred mouse strain8. Following embryo isolation, neutral protease Dispase II was used to treat the whole embryo. Then, the facial prominences were dissected out, and the facial ectoderm was separated from the mesenchyme. This method keeps both the facial ectoderm and mesenchyme intact. The samples obtained using this methodology can be used for techniques including protein detection, chromatin immunoprecipitation (ChIP) assay, microarray studies, and RNA-seq.

After Dispase II treatment, the ectoderm of the embryo tends to be loose. The facial prominences should be intact after dissection (Figure 2A). The isolated facial ectoderm is clear and free of mesenchyme tissue (Figure 2B). To determine the efficacy of the protocol and to detect cross contamination we assayed for forebrain expressed Zic39, ectodermal specific cdh15 and mesenchymal specific sox1010 using reverse transcriptase PCR. Our studies confirmed th...

Watch this Scientific Journal Video about Separation of Mouse Embryonic Facial Ectoderm and Mesenchyme at JoVE.com

Separation of Mouse Embryonic Facial Ectoderm and Mesenchyme

A protocol for separation of embryo facial ectoderm and mesenchyme is described. We use Dispase II to treat whole embryos first, dissect whole facial prominences out, and then separate the facial ectoderm and mesenchyme.

 Orofacial clefts are the most frequent  craniofacial defects, which affect 1.5 in 1,000 newborns worldwide1,2. Orofacial clefting is caused by abnormal ...

separation-of-mouse-embryonic-facial-ectoderm-and-mesenchyme

Research

JoVE Journal

Biology

10.9K vistas.  University of Colorado Denver Anschutz Medical Campus. Un protocolo para la separación de ectodermo y el mesénquima facial embrión se describe. Utilizamos Dispasa II para el tratamiento de embriones enteros en primer lugar, analizar las prominencias faciales completas, y luego separar el ectodermo facial y mesénquima.

Video: Separation of Mouse Embryonic Facial Ectoderm and Mesenchyme

Electrophoretic Separation of Proteins

Electrophoresis is used to separate complex mixtures of proteins (e.g., from cells, subcellular fractions, column fractions, or immunoprecipitates), to investigate subunit compositions, and to verify homogeneity of protein samples. It can also serve to purify proteins for use in further applications. In polyacrylamide gel electrophoresis, proteins migrate in response to an electrical field through pores in a polyacrylamide gel matrix; pore size decreases with increasing acrylamide concentration. The combination of pore size and protein charge, size, and shape determines the migration rate of the protein. In this unit, the standard Laemmli method is described for discontinuous gel electrophoresis under denaturing conditions, i.e., in the presence of sodium dodecyl sulfate (SDS). 

In this video, we demonstrate a method for electrophoretic separation of proteins using poly-acrylimide gel electrophoresis (PAGE).

Electrophoresis is used to separate complex mixtures of proteins (e.g., from cells, subcellular fractions, column fractions, or immunoprecipitates), to ...

Isolation and Derivation of Mouse Embryonic Germinal Cells

The ability of embryonic germinal cells (EG) to differentiate into primordial germinal cells (PGCs) and later into gametes during early developmental stages is a perfect model to address our hypothesis about cancer and infertility. This protocol shows how to isolate primordial germinal cells from developing gonads in 10.5-11.5 days post coitum (dpc) mouse embryos.  Developing gonadal ridges from mouse embryos (C57BL6J) were dissociated by mechanical disruption with collagenase, then plated in a mouse embryo fibroblast feeder layer (MEF-CF1) that was previously mitotically inactivated with mitomycin C in the presence of knockout media and supplemented with Leukemia Inhibitor Factor (LIF), basic Fibroblast Growth Factor (bFGF), and Stem Cell Factor (SCF).  Using these optimized methods for PCG identification, isolation, and establishment of culture conditions permits long term cultures of EG cells for more than 40 days.  The embryonic germinal cell lines showed embryonic phenotype and expression of common used markers of the pluripotent state.  Isolation and derivation of germinal cells in culture provide a tool to understand their development in vitro and offer the opportunity to monitor cumulative damage at genetic and epigenetic levels after exposure to oxidative stress.

The ability of embryonic germinal cells to differentiate into primordial germinal cells during early development stages is a perfect model to address our hypothesis about cancer and infertility. This protocol shows how to isolate primordial germinal cells from developing gonads in 10.5-11.5 days post coitum mouse embryos.

The ability of embryonic germinal cells (EG) to differentiate into primordial germinal cells (PGCs) and later into gametes during early developmental ...

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

Adenovirus-mediated Genetic Removal of Signaling Molecules in Cultured Primary Mouse Embryonic Fibroblasts

The ability to genetically remove specific components of various cell signalling cascades has been an integral tool in modern signal transduction analysis. One particular method to achieve this conditional deletion is via the use of the Cre-loxP system. This method involves flanking the gene of interest with loxP sites, which are specific recognition sequences for the Cre recombinase protein. Exposure of the so-called floxed (flanked by loxP site) DNA to this enzyme results in a Cre-mediated recombination event at the loxP sites, and subsequent excision of the intervening gene3. Several different methods exist to administer Cre recombinase to the site of interest. In this video, we demonstrate the use of an adenovirus containing the Cre recombinase gene to infect primary mouse embryonic fibroblasts (MEFs) obtained from embryos containing a floxed Rac1 allele1. Our rationale for selecting Rac1 MEFs for our experiments is that clear morphological changes can be seen upon deletion of Rac1, due to alterations in the actin cytoskeleton2,5. 72 hours following viral transduction and Cre expression, cells were stained using the actin dye phalloidin and imaged using confocal laser scanning microscopy. It was observed that MEFs which had been exposed to the adeno-Cre virus appeared contracted and elongated in morphology compared to uninfected cells, consistent with previous reports2,5. The adenovirus method of Cre recombinase delivery is advantageous as the adeno-Cre virus is easily available, and gene deletion via Cre in nearly 100% of the cells can be achieved with optimized adenoviral infection.

In this video we use an adenovirus carrying the Cre recombinase gene to infect primary mouse embryonic fibroblasts carrying a floxed Rac1 allele.

The ability to genetically remove specific components of various cell signalling cascades has been an integral tool in modern signal transduction ...

Isolation and Culture of Cells from the Nephrogenic Zone of the Embryonic Mouse Kidney

Embryonic development of the kidney has been extensively studied both as a model for epithelial-mesenchymal interaction in organogenesis and to gain understanding of the origins of congenital kidney disease. More recently, the possibility of steering na&iuml;ve embryonic stem cells toward nephrogenic fates has been explored in the emerging field of regenerative medicine. Genetic studies in the mouse have identified several pathways required for kidney development, and a global catalog of gene transcription in the organ has recently been generated <a href="http://www.gudmap.org/">http://www.gudmap.org/</a>, providing numerous candidate regulators of essential developmental functions. Organogenesis of the rodent kidney can be studied in organ culture, and many reports have used this approach to analyze outcomes of either applying candidate proteins or knocking down the expression of candidate genes using siRNA or morpholinos. However, the applicability of organ culture to the study of signaling that regulates stem/progenitor cell differentiation versus renewal in the developing kidney is limited as cultured organs contain a compact extracellular matrix limiting diffusion of macromolecules and virus particles. To study the cell signaling events that influence the stem/progenitor cell niche in the kidney we have developed a primary cell system that establishes the nephrogenic zone or progenitor cell niche of the developing kidney ex vivo in isolation from the epithelial inducer of differentiation. Using limited enzymatic digestion, nephrogenic zone cells can be selectively liberated from developing kidneys at E17.5. Following filtration, these cells can be cultured as an irregular monolayer using optimized conditions. Marker gene analysis demonstrates that these cultures contain a distribution of cell types characteristic of the nephrogenic zone in vivo, and that they maintain appropriate marker gene expression during the culture period. These cells are highly accessible to small molecule and recombinant protein treatment, and importantly also to viral transduction, which greatly facilitates the study of candidate stem/progenitor cell regulator effects. Basic cell biological parameters such as proliferation and cell death as well as changes in expression of molecular markers characteristic of nephron stem/progenitor cells in vivo can be successfully used as experimental outcomes. Ongoing work in our laboratory using this novel primary cell technique aims to uncover basic mechanisms governing the regulation of self-renewal versus differentiation in nephron stem/progenitor cells.

In this report we describe a method for the isolation and culture of the progenitor cell niche from the embryonic mouse kidney that can be used to study signaling pathways regulating stem/progenitor cells of the developing kidney. These cultured cells are highly accessible to small molecule and recombinant protein treatment, and importantly also to viral transduction, which allows efficient manipulation of candidate pathways.

Embryonic development of the kidney has been extensively studied both as a model for epithelial-mesenchymal interaction in organogenesis and  to gain ...

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Cell polarity is a fundamental property of eukaryotic cells that is dynamically regulated by both intrinsic and extrinsic factors during embryonic development 1, 2. One of the signaling pathways involved in this regulation is the Wnt pathway, which is used many times during embryogenesis and critical for human disease3, 4, 5. Multiple molecular components of this pathway coordinately regulate signaling in a spatially-restricted manner, but the underlying mechanisms are not fully understood. Xenopus embryonic epithelial cells is an excellent system to study subcellular localization of various signaling proteins. Fluorescent fusion proteins are expressed in Xenopus embryos by RNA microinjection, ectodermal explants are prepared and protein localization is evaluated by epifluorescence. In this experimental protocol we describe how subcellular localization of Diversin, a cytoplasmic protein that has been implicated in signaling and cell polarity determination6, 7 is visualized in Xenopus ectodermal cells to study Wnt signal transduction8. Coexpression of a Wnt ligand or a Frizzled receptor alters the distribution of Diversin fused with red fluorescent protein, RFP, and recruits it to the cell membrane in a polarized fashion 8, 9. This ex vivo protocol should be a useful addition to in vitro studies of cultured mammalian cells, in which spatial control of signaling differs from that of the intact tissue and is much more difficult to analyze.

Polarized Translocation of Fluorescent Proteins in Xenopus Ectoderm in Response to Wnt Signaling

Xenopus embryonic ectoderm has become an attractive model for studies of cell polarity. An assay is described, in which subcellular distribution of fluorescent proteins is assessed in ectoderm cells. This protocol will help address questions related to spatial control of signaling.

Cell polarity is a fundamental property of eukaryotic cells that is dynamically regulated by both intrinsic and extrinsic factors during embryonic ...

Electroporation of Craniofacial Mesenchyme

Electroporation is an efficient method of delivering DNA and other charged macromolecules into tissues at precise time points and in precise locations. For example, electroporation has been used with great success to study neural and retinal development in Xenopus, chicken and mouse 1-10. However, it is important to note that in all of these studies, investigators were not targeting soft tissues. Because we are interested in craniofacial development, we adapted a method to target facial mesenchyme.
When we searched the literature, we found, to our surprise, very few reports of successful gene transfer into cartilaginous tissue. The majority of these studies were gene therapy studies, such as siRNA or protein delivery into chondrogenic cell lines, or, animal models of arthritis 11-13. In other systems, such as chicken or mouse, electroporation of facial mesenchyme has been challenging (personal communications, Dept of Craniofacial Development, KCL). We hypothesized that electroporation into procartilaginous and cartilaginous tissues in Xenopus might work better. In our studies, we show that gene transfer into the facial cartilages occurs efficiently at early stages (28), when the facial primordium is still comprised of soft tissue prior to cartilage differentiation.
Xenopus is a very accessible vertebrate system for analysis of craniofacial development. Craniofacial structures are more readily visible in Xenopus than in any other vertebrate model, primarily because Xenopus embryos are fertilized externally, allowing analyses of the earliest stages, and facilitating live imaging at single cell resolution, as well as reuse of the mothers 14. Among vertebrate models developing externally, Xenopus is more useful for craniofacial analysis than zebrafish, as Xenopus larvae are larger and easier to dissect, and the developing facial region is more accessible to imaging than the equivalent region in fish. In addition, Xenopus is evolutionarily closer to humans than zebrafish (&tilde;100 million years closer) 15. Finally, at these stages, Xenopus tadpoles are transparent, and concurrent expression of fluorescent proteins or molecules will allow easy visualization of the developing cartilages. We anticipate that this approach will allow us to rapidly and efficiently test candidate molecules in an in vivo model system.

Craniofacial cartilages develop in close contact with other tissues and are difficult to manipulate in live animals. We are using electroporation to deliver molecular tools during growth of the craniofacial skeleton while bypassing early embryonic effects. This approach will allow us to efficiently test candidate molecules in vivo.

Electroporation is an efficient method of delivering DNA and other charged macromolecules into tissues at precise time points and in precise locations. ...

Efficient Differentiation of Mouse Embryonic Stem Cells into Motor Neurons

Direct differentiation of embryonic stem (ES) cells into functional motor neurons represents a promising resource to study disease mechanisms, to screen new drug compounds, and to develop new therapies for motor neuron diseases such as spinal muscular atrophy (SMA) and amyotrophic lateral sclerosis (ALS). Many current protocols use a combination of retinoic acid (RA) and sonic hedgehog (Shh) to differentiate mouse embryonic stem (mES) cells into motor neurons1-4. However, the differentiation efficiency of mES cells into motor neurons has only met with moderate success. We have developed a two-step differentiation protocol5 that significantly improves the differentiation efficiency compared with currently established protocols. The first step is to enhance the neuralization process by adding Noggin and fibroblast growth factors (FGFs). Noggin is a bone morphogenetic protein (BMP) antagonist and is implicated in neural induction according to the default model of neurogenesis and results in the formation of anterior neural patterning6. FGF signaling acts synergistically with Noggin in inducing neural tissue formation by promoting a posterior neural identity7-9. In this step, mES cells were primed with Noggin, bFGF, and FGF-8 for two days to promote differentiation towards neural lineages. The second step is to induce motor neuron specification. Noggin/FGFs exposed mES cells were incubated with RA and a Shh agonist, Smoothened agonist (SAG), for another 5 days to facilitate motor neuron generation. To monitor the differentiation of mESs into motor neurons, we used an ES cell line derived from a transgenic mouse expressing eGFP under the control of the motor neuron specific promoter Hb91. Using this robust protocol, we achieved 51&plusmn;0.8% of differentiation efficiency (n = 3; p &lt; 0.01, Student's t-test)5. Results from immunofluorescent staining showed that GFP+ cells express the motor neuron specific markers, Islet-1 and choline acetyltransferase (ChAT). Our two-step differentiation protocol provides an efficient way to differentiate mES cells into spinal motor neurons.

We developed a new protocol to improve efficiency of in vitro differentiation of mouse embryonic stem cells into motor neurons. The differentiated ES cells acquired motor neurons features as evidenced by expression of neuronal and motor neuron markers using immunohistochemical techniques.

Direct differentiation of embryonic stem (ES) cells into functional  motor neurons represents a promising resource to study disease mechanisms, to  screen ...

Preparation of Mouse Embryonic Fibroblast Cells Suitable for Culturing Human Embryonic and Induced Pluripotent Stem Cells

In general, human embryonic stem cells (hESCs) and human induced pluripotent stem cells (hiPSCs)1 can be cultured under variable conditions. However, it is not easy to establish an effective system for culturing these cells. Since the culture conditions can influence gene expression that confers pluripotency in hESCs and hiPSCs, the optimization and standardization of the culture method is crucial.
The establishment of hESC lines was first described by using MEFs as feeder cells and fetal bovine serum (FBS)-containing culture medium2. Next, FBS was replaced with knockout serum replacement (KSR) and FGF2, which enhances proliferation of hESCs3. Finally, feeder-free culture systems enable culturing cells on Matrigel-coated plates in KSR-containing conditioned medium (medium conditioned by MEFs)4. Subsequently, hESCs culture conditions have moved towards feeder-free culture in chemically defined conditions5-7. Moreover, to avoid the potential contamination by pathogens and animal proteins culture methods using xeno-free components have been established8.
To obtain improved conditions mouse feeder cells have been replaced with human cell lines (e.g. fetal muscle and skin cells9, adult skin cells10, foreskin fibroblasts11-12, amniotic mesenchymal cells13). However, the efficiency of maintaining undifferentiated hESCs using human foreskin fibroblast-derived feeder layers is not as high as that from mouse feeder cells due to the lower level of secretion of Activin A14. Obviously, there is an evident difference in growth factor production by mouse and human feeder cells.
Analyses of the transcriptomes of mouse and human feeder cells revealed significant differences between supportive and non-supportive cells. Exogenous FGF2 is crucial for maintaining self-renewal of hESCs and hiPSCs, and has been identified as a key factor regulating the expression of Tgf&beta;1, Activin A and Gremlin (a BMP antagonist) in feeder cells. Activin A has been shown to induce the expression of OCT4, SOX2, and NANOG in hESCs15-16.
For long-term culture, hESCs and hiPSCs can be grown on mitotically inactivated MEFs or under feeder-free conditions in MEF-CM (MEF-Conditioned Medium) on Matrigel-coated plates to maintain their undifferentiated state. Success of both culture conditions fully depends on the quality of the feeder cells, since they directly affect the growth of hESCs.
Here, we present an optimized method for the isolation and culture of mouse embryonic fibroblasts (MEFs), preparation of conditioned medium (CM) and enzyme-linked immunosorbent assay (ELISA) to assess the levels of Activin A within the media.

The quality of mouse embryonic fibroblasts (MEFs) is dictated by the right strain of mouse such as CF-1. Pluripotency-supportive MEFs and conditioned media (CM) obtained from these should contain optimal concentrations of Activin A, Gremlin and Tgf&beta;1 needed for the Activin/Nodal and FGF pathways to co-operatively maintain self-renewal and pluripotency.

In general, human embryonic stem cells (hESCs) and human induced pluripotent stem cells (hiPSCs)1 can be cultured under variable conditions. However, it ...

A System for ex vivo Culturing of Embryonic Pancreas

The pancreas controls vital functions of our body, including the production of digestive enzymes and regulation of blood sugar levels1. Although in the past decade many studies have contributed to a solid foundation for understanding pancreatic organogenesis, important gaps persist in our knowledge of early pancreas formation2. A complete understanding of these early events will provide insight into the development of this organ, but also into incurable diseases that target the pancreas, such as diabetes or pancreatic cancer. Finally, this information will generate a blueprint for developing cell-replacement therapies in the context of diabetes.
 During embryogenesis, the pancreas originates from distinct embryonic outgrowths of the dorsal and ventral foregut endoderm at embryonic day (E) 9.5 in the mouse embryo3,4. Both outgrowths evaginate into the surrounding mesenchyme as solid epithelial buds, which undergo proliferation, branching and differentiation to generate a fully mature organ2,5,6. Recent evidences have suggested that growth and differentiation of pancreatic cell lineages, including the insulin-producing &beta;-cells, depends on proper tissue-architecture, epithelial remodeling and cell positioning within the branching pancreatic epithelium7,8. However, how branching morphogenesis occurs and is coordinated with proliferation and differentiation in the pancreas is largely unknown. This is in part due to the fact that current knowledge about these developmental processes has relied almost exclusively on analysis of fixed specimens, while morphogenetic events are highly dynamic.
 Here, we report a method for dissecting and culturing mouse embryonic pancreatic buds ex vivo on glass bottom dishes, which allow direct visualization of the developing pancreas (Figure 1). This culture system is ideally devised for confocal laser scanning microscopy and, in particular, live-cell imaging. Pancreatic explants can be prepared not only from wild-type mouse embryos, but also from genetically engineered mouse strains (e.g. transgenic or knockout), allowing real-time studies of mutant phenotypes. Moreover, this ex vivo culture system is valuable to study the effects of chemical compounds on pancreatic development, enabling to obtain quantitative data about proliferation and growth, elongation, branching, tubulogenesis and differentiation. In conclusion, the development of an ex vivo pancreatic explant culture method combined with high-resolution imaging provides a strong platform for observing morphogenetic and differentiation events as they occur within the developing mouse embryo.

A System for ex vivo Culturing of Embryonic Pancreas

Here, we describe a method for isolation, culture and manipulation of mouse embryonic pancreas. This represents an excellent ex vivo system for studying various aspects of pancreatic development, including morphogenesis, differentiation and growth. Pancreatic bud explants can be cultured for several days and used in a range of different applications, including whole-mount immunofluorescence and live imaging.

The pancreas controls vital functions of our body,  including the production of digestive enzymes and regulation of blood sugar  levels1. Although in the ...