This work was supported by the Direcci&#243;n General de Asuntos del Personal Acad&#233;mico (DGAPA)-Universidad Nacional Aut&#243;noma de M&#233;xico [grant numbers IN211117 and IN213314] and Consejo Nacional de Ciencia y Tecnolog&#237;a (CONACyT) [grant number 1887 CONACyT-Fronteras de la Ciencia] awarded to JC-M. JC M-L received a postdoctoral fellowship from the Consejo Nacional de Ciencia y Tecnolog&#237;a (CONACyT-Fronteras de la Ciencia-1887). The authors appreciate the help of Lic. Lucia&#160;Brito from Instituto de Investigaciones Biom&#233;dicas, UNAM in the preparation references of this manuscript.

This research was reviewed and approved by the Institutional Review Board for the Care and Use of Laboratory Animals of the Instituto de Investigaciones Biom&#233;dicas, Universidad Nacional Aut&#243;noma de M&#233;xico (UNAM, Mexico City, Mexico).
1. Egg incubation and embryo staging
NOTE: Fertilized hen eggs can be obtained from local farms. Fertilized White Leghorn chicken eggs are most commonly used. Store the freshly fertilized chicken eggs at 15 &#176;C for up to 1 week prior to incubation.
<ol>
	<li>Incubate the fertilized chicken eggs vertically with the pointy side down in a humidified incubator at 38 &#176;C and 70% relative humidity until they reach the 28 HH stage (approximately 5.5 days according to Hamilton and Hamburger, 1951)22. Rotate the eggs during the incubation to prevent the embryo from adhering to the shell membrane. 
	NOTE: The choice of the developmental stage is informed by the experimental aims. In this case, the 28 HH stage is optimal for interdigit bead implantation to induce an ectopic digit or promote cell death.</li>
	<li>Remove the eggs from the incubator, swab them with 70% ethanol, and allow them to air dry. Disinfect the working area, microscopes, and instruments with 70% ethanol.</li>
	<li>Candle the egg to identify blood vessels and locate the embryo. Discard eggs that do not have an embryo.</li>
	<li>Using the end of non-toothed forceps, open a window in around five eggs by tapping the blunt end of an egg, and remove a 1-cm2 section of the shell with the forceps.</li>
	<li>Transfer the egg into a carton or plastic holder and place it under the stereomicroscope. Remove the air membrane by puncturing and pulling it out with fine surgical forceps. Remove any small piece of eggshell that could contact the embryo. 
	NOTE: The air membrane is the white, opaque membrane observed immediately after windowing the egg.</li>
	<li>Observe under the microscope while opening the amniotic sac slightly, tearing it using the fine surgical forceps. Be careful not to damage the vasculature of the chorioallantoic membrane. 
	&#8203;NOTE: The amnion is the transparent membrane closely surrounding the embryo that encapsulates it in amniotic fluid.</li>
	<li>Stage the embryos in ovo to determine whether they are in the desired stage. Embryos in earlier stages can be returned to the incubator after sealing the eggshell window with tape.</li>
</ol>
2. Bead preparation
NOTE: Depending on the experimental aim and treatment in question, alternative bead types (e.g., Affi-Gel, AG1-X2, heparin) may be more suitable. Affi-Gel beads are optimal for proteins (e.g., TGF-&#223;1), while heparin beads are ideal for growth factors (e.g., FGFs, WNT) and AG1-X2 beads for chemicals solubilized in organic solvents (e.g., DMSO).
<ol>
	<li>Affi-Gel and heparin bead preparation
	<ol>
		<li>Cut a square of parafilm to fit a 45-mm Petri dish. Place the parafilm across the bottom of the petri dish to cover it and fix it to the bottom of the Petri dish by pushing each vertex using the end of non-toothed forceps. Set aside.</li>
		<li>Using a pipette or spatula, transfer the beads into a microcentrifuge tube and wash them twice in 1x PBS by settling and pipetting.</li>
		<li>Transfer ~40-50 beads with a micropipette to the center of the parafilmed Petri dish from step 1.</li>
		<li>Using the microscope, select ~30 Affi-Gel or heparin beads ~100 &#181;m in diameter for use. Use a microscope eyepiece reticle to size the beads or use the third interdigit of an HH 28 embryo as a reference; the bead must be smaller than the interdigit.</li>
		<li>Carefully remove as much of the excess PBS surrounding the beads as possible and soak them in 2-5 &#181;L of the treatment solution. Assure that the solution completely covers beads.</li>
		<li>In parallel, prepare control beads by soaking in a solution that contains the same amount of vehicle as used in the experimental treatment solution. 
		NOTE: Concentrations need to be calculated for each treatment according to the experiment. Use the appropriate personal protective equipment when handling potentially harmful reagents.</li>
		<li>Incubate the beads in the solution for 30 min at room temperature. To prevent the beads from drying out during the incubation, pipette a few drops of 1x PBS or water around the beads to humidify the local atmosphere and cover the dish with parafilm to slow evaporation.</li>
		<li>Place the Petri dish on ice and implant the beads within the same day.</li>
	</ol>
	</li>
	<li>AG 1-X2 bead preparation
	<ol>
		<li>Cut a square of parafilm to fit a 45-mm Petri dish. Cover the bottom of the Petri dish with parafilm and affix by pushing each vertex using the end of non-toothed forceps. Set aside.</li>
		<li>Use a spatula to transfer the AG 1-X2 beads into a microcentrifuge tube. Add 30-50 &#181;L of the treatment solution at the desired final concentration.</li>
		<li>In parallel, incubate the control beads in a solution with the vehicle alone, prepared without the experimental chemical or protein of interest.</li>
		<li>Incubate the beads for 20 min while slowly shaking at room temperature. Wrap the microcentrifuge tubes with foil to protect from light during all incubation, given that many of these molecules are light-sensitive</li>
		<li>Remove as much of the solution as possible using a pipette and stain with 2% phenol red dissolved in water at room temperature for 2 min with mild agitation in a vortex. 
		NOTE: Dyeing the transparent AG 1-X2 beads facilitates their implantation in the embryo.</li>
		<li>Remove the 2% phenol red and wash the beads twice in 1x PBS to remove any excess dye.</li>
		<li>With a micropipette tip, transfer ~40-50 AG 1-X2 beads to the center of the parafilm-covered Petri dish prepared in step 1 and soak these in 5 &#181;L of 1x PBS. Under the microscope, select around 30 beads that are ~100 &#181;m diameter in size. Discard the unused beads.</li>
		<li>Beads are ready to implant. Ensure the beads are submerged in the PBS throughout the implantation by pipetting a few drops of 1x PBS or water around the beads to humidify the local atmosphere and/or cover the dish with parafilm to slow evaporation.</li>
	</ol>
	</li>
</ol>
3. Embryo manipulation and interdigit implantation
<ol>
	<li>Before manipulating the embryos, arrange two stereomicroscopes next to each other on a benchtop. One is for embryo manipulation and bead implantation, and the other is for maintaining the treated beads ready to implant into the embryo.</li>
	<li>Using non-toothed forceps, create a window in the remaining eggs as described in the egg incubation and embryo staging section.</li>
	<li>Under the microscope, open the amniotic sac by tearing the amniotic membrane with fine surgical forceps near the right hindlimb only by the amount needed to accomplish the procedure (i.e., as little as possible).</li>
	<li>Using fine forceps, hold the embryo by the amniotic membrane to expose the right hindlimb. Using a fine tungsten needle, make a hole centered in the distal-most of the third interdigit of the hindlimb.</li>
	<li>Without releasing the embryo and the exposed hindlimb, take one treated bead from the other microscope using one of the tips of the forceps. The forceps must be in the open position for one bead to adhere to the forceps tip.</li>
	<li>Transfer the bead into the chick embryo near the hindlimb and position it on top of the interdigit hole.</li>
	<li>Close the forceps to apply pressure to the bead until it enters the hole.</li>
	<li>Release the embryo and seal the eggshell window with tape.</li>
	<li>Return the eggs to the incubator until they have reached the required stage. Repeat the procedure until the required number of manipulated embryos is reached. It is imperative to ensure that the beads do not dry out at any point. 
	NOTE: To observe a complete ectopic finger, incubate for ~72 h after bead implantation. In contrast, early differentiation genes (e.g., Sox9) are expressed ~30 min after the implantation of a TGF-&#223;1-soaked bead.</li>
</ol>

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The authors have nothing to disclose.

The main advantage of the experimental tool detailed in this protocol is being able to control the time and location of the exposure to beads soaked in a given experimental molecule. Combining the correct positioning with precise developmental timing provides enormous possibilities to study cell differentiation processes. Performing these experiments in undifferentiated tissue enables investigating the first crucial events in cellular lineage. For example, placing a TGF&#223;-soaked bead in the interdigital tissue of emb...

Breakthroughs in the molecular mechanisms that control gene expression during embryonic development have allowed us to understand how cell fate is determined. Commitment to different cell lineages occurs once cells begin the molecular expression of transcription factors1. This expression pattern is highly coordinated in space and time and thereby directs the shaping, positioning, and patterning of cells, tissues, and organs1,2,3,4,5. Embryonic induction is the process by which cells are committed to specific lineages by establishing hierarchies that restrict cells&#39; potentiality, which even include the generation of the basic body plan as occurs with the Spemann organizer6,7. The blastopore dorsal lip induces a second embryonic axis in a host embryo8,9. Today, with the aid of grafting and other classical experiments combined with molecular approaches, it is known that different transcription factors and growth factors function to direct embryonic induction in the Spemann organizer10. Thus, experimental manipulation is an important tool to understand cell differentiation, morphogenesis, and patterning processes during embryogenesis.
Interestingly, in embryonic systems where tissue transplantation is difficult or when the inducers are already well known, carriers are used to deliver molecules (e.g., proteins, chemicals, toxins, etc.) to regulate cell differentiation, morphogenesis, and even patterning. One such carrier system involves implanting beads soaked in a specific molecule in any experimental model organism at any developmental time point to determine the effect of the said reagent or direct the differentiation of the said model. For example, by implanting retinoic acid (RA)-soaked beads into the chicken wing limb bud, Cheryl Tickle et al. (1985) demonstrated that RA induces the expression of sonic hedgehog in the zone of polarizing activity (ZPA)11,12. The same experimental strategy was used to discover that RA controls the asymmetry of somites and cell death in the limb bud during digit development and in other embryonic limb regions13,14,15. Other factors, mainly proteins (e.g., fibroblast growth factors [FGF], transforming growth factor-beta [TGF-&#223;]) have been used to induce limbs in early embryos&#39; flanks and new digits in the interdigital region, respectively16,17,18,19,20,21. These experiments evidence the power and utility of this technique for determining the stage of commitment or competence of tissues or groups of cells exposed to the molecules.
In this protocol, the chick limb at the stage of digit formation served as the experimental model to present step-by-step how to prepare and implant the soaked beads. However, this experimental tool is not limited to this application but can be exploited in any experimental animal model and any timepoint in vitro and in vivo to study induction, differentiation, cell death, and patterning.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Affi-Gel Blue Gel beads</td>
			<td>Bio-Rad</td>
			<td>153-7302</td>
			<td></td>
		</tr>
		<tr>
			<td>AG1-X2 beads</td>
			<td>Bio-Rad</td>
			<td>1400123</td>
			<td></td>
		</tr>
		<tr>
			<td>Egg incubator</td>
			<td>Incumatic de Mexico</td>
			<td>Incumatic 1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine surgical forceps</td>
			<td>Fine Science Tools</td>
			<td>9115-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparine Sepharose beads</td>
			<td>Abcam</td>
			<td>ab193268</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dish</td>
			<td>Nest</td>
			<td>705001</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereomicroscope</td>
			<td>Zeiss</td>
			<td>Stemi DV4</td>
			<td></td>
		</tr>
		<tr>
			<td>Tape</td>
			<td>NA</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Tungsten needle</td>
			<td>GoodFellow</td>
			<td>E74-15096/01</td>
			<td></td>
		</tr>
	</tbody>
</table>

analysis of cell differentiation, morphogenesis, and patterning during chicken embryogenesis using the soaked-bead assay

A multitude of genetic programs is activated during embryonic development that orchestrates cell differentiation to generate an astounding diversity of somatic cells, tissues, and organs. The precise activation of these genetic programs is regulated by morphogens, diffusible molecules that direct cell fate at different thresholds. Understanding how genetic activation coordinates morphogenesis requires the study of local interactions triggered by morphogens during development. The use of beads soaked in proteins or drugs implanted into distinct regions of the embryo enables studying the role of specific molecules in the establishment of digits and other developmental processes. This experimental technique provides information on the control of cell induction, cell fate, and pattern formation. Thus, this soaked bead assay is an extremely powerful and valuable experimental tool applicable to other embryonic models.

Using soaked beads to evaluate cell behavior in the embryonic chick limb 
To assure the efficacy of this assay, the bead must be placed consistently and precisely in the correct location; in this case, the distal-most of the third interdigit beneath the apical ectodermal ridge AER (Figure 1A). This positioning permits the molecule in question to spread equally throughout the interdigital tissue. Moreover, the zone beneath the AER contains undifferentiated cells that are...

Watch this Scientific Journal Video about Analysis of Cell Differentiation, Morphogenesis, and Patterning During Chicken Embryogenesis Using the Soaked-Bead Assay at JoVE.com

Analysis of Cell Differentiation, Morphogenesis, and Patterning During Chicken Embryogenesis Using the Soaked-Bead Assay

The soaked bead assay involves targeted delivery of test reagent at any developmental time point to study the regulation of cell differentiation and morphogenesis. A detailed protocol, applicable to any experimental animal model, for preparing three different types of soaked beads and implanting these in the interdigit of a chicken embryo is presented.

A multitude of genetic programs is activated during embryonic development that orchestrates cell differentiation to generate an astounding diversity of ...

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3.0K Views.  Universidad Nacional Autónoma de México. The soaked bead assay involves targeted delivery of test reagent at any developmental time point to study the regulation of cell differentiation and morphogenesis. A detailed protocol, applicable to any experimental animal model, for preparing three different types of soaked beads and implanting these in the interdigit of a chicken embryo is presented.

Video: Analysis of Cell Differentiation, Morphogenesis, and Patterning During Chicken Embryogenesis Using the Soaked-Bead Assay

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Differentiation of the SH-SY5Y Human Neuroblastoma Cell Line

Having appropriate in vivo and in vitro systems that provide translational models for human disease is an integral aspect of research in neurobiology and the neurosciences. Traditional in vitro experimental models used in neurobiology include primary neuronal cultures from rats and mice, neuroblastoma cell lines including rat B35 and mouse Neuro-2A cells, rat PC12 cells, and short-term slice cultures. While many researchers rely on these models, they lack a human component and observed experimental effects could be exclusive to the respective species and may not occur identically in humans. Additionally, although these cells are neurons, they may have unstable karyotypes, making their use problematic for studies of gene expression and reproducible studies of cell signaling. It is therefore important to develop more consistent models of human neurological disease. 
The following procedure describes an easy-to-follow, reproducible method to obtain homogenous and viable human neuronal cultures, by differentiating the chromosomally stable human neuroblastoma cell line, SH-SY5Y. This method integrates several previously described methods1-4 and is based on sequential removal of serum from media. The timeline includes gradual serum-starvation, with introduction of extracellular matrix proteins and neurotrophic factors. This allows neurons to differentiate, while epithelial cells are selected against, resulting in a homogeneous neuronal culture. Representative results demonstrate the successful differentiation of SH-SY5Y neuroblastoma cells from an initial epithelial-like cell phenotype into a more expansive and branched neuronal phenotype. This protocol offers a reliable way to generate homogeneous populations of neuronal cultures that can be used for subsequent biochemical and molecular analyses, which provides researchers with a more accurate translational model of human infection and disease.

It is critical in neurobiology and neurovirology to have a reliable, replicable in vitro system that serves as a translational model for what occurs in vivo in human neurons. This protocol describes how to culture and differentiate SH-SY5Y human neuroblastoma cells into viable neurons for use in in vitro applications.

Having appropriate in vivo and in vitro systems that provide translational models for human disease is an integral aspect of research in neurobiology and ...

Studying Wnt Signaling During Patterning of Conducting Airways

Wnt signaling pathways play critical roles during development of the respiratory tract. Defining precise mechanisms of differentiation and morphogenesis controlled by Wnt signaling is required to understand how tissues are patterned during normal development. This knowledge is also critical to determine the etiology of birth defects such as lung hypoplasia and tracheobronchomalacia. Analysis of earliest stages of development of respiratory tract imposes challenges, as the limited amount of tissue prevents the performance of standard protocols better suited for postnatal studies. In this paper, we discuss methodologies to study cell differentiation and proliferation in the respiratory tract. We describe techniques such as whole mount staining, processing of the tissue for confocal microscopy and immunofluorescence in paraffin sections applied to developing tracheal lung. We also discuss methodologies for the study of tracheal mesenchyme differentiation, in particular cartilage formation. Approaches and techniques discussed in the current paper circumvent the limitation of material while working with embryonic tissue, allowing for a better understanding of the patterning process of developing conducting airways.

The use of reporter mice coupled to whole mount and section staining, microscopy and in vivo assays facilitates the analysis of mechanisms underlying the normal patterning of the respiratory tract. Here we describe how these techniques contributed to the analysis of Wnt signaling during tracheal development.

Wnt signaling pathways play critical roles during development of the respiratory tract. Defining precise mechanisms of differentiation and morphogenesis ...

Differentiation of Atrial Cardiomyocytes from Pluripotent Stem Cells Using the BMP Antagonist Grem2

Protocols for generating populations of cardiomyocytes from pluripotent stem cells have been developed, but these generally yield cells of mixed phenotypes. Researchers interested in pursuing studies involving specific myocyte subtypes require a more directed differentiation approach. By treating mouse embryonic stem (ES) cells with Grem2, a secreted BMP antagonist that is necessary for atrial chamber formation in vivo, a large number of cardiac cells with an atrial phenotype can be generated. Use of the engineered Myh6-DSRed-Nuc pluripotent stem cell line allows for identification, selection, and purification of cardiomyocytes. In this protocol embryoid bodies are generated from Myh6-DSRed-Nuc cells using the hanging drop method and kept in suspension until differentiation day 4 (d4). At d4 cells are treated with Grem2 and plated onto gelatin coated plates. Between d8-d10 large contracting areas are observed in the cultures and continue to expand and mature through d14. Molecular, histological and electrophysiogical analyses indicate cells in Grem2-treated cells acquire atrial-like characteristics providing an in vitro model to study the biology of atrial cardiomyocytes and their response to various pharmacological agents.

Generating cardiomyocytes from pluripotent stem cells in vitro allows access to large amounts of cardiac tissue in vitro for basic science and clinical applications. This protocol uses the atrializing factor Grem2 to both increase the numbers of cardiomyocytes obtained and to generate cardiomyocytes with an atrial phenotype.

Protocols for generating populations of cardiomyocytes from pluripotent stem cells have been developed, but these generally yield cells of mixed ...

Prediction and Validation of Gene Regulatory Elements Activated During Retinoic Acid Induced Embryonic Stem Cell Differentiation

Embryonic development is a multistep process involving activation and repression of many genes. Enhancer elements in the genome are known to contribute to tissue and cell-type specific regulation of gene expression during the cellular differentiation. Thus, their identification and further investigation is important in order to understand how cell fate is determined. Integration of gene expression data (e.g., microarray or RNA-seq) and results of chromatin immunoprecipitation (ChIP)-based genome-wide studies (ChIP-seq) allows large-scale identification of these regulatory regions. However, functional validation of cell-type specific enhancers requires further in vitro and in vivo experimental procedures. Here we describe how active enhancers can be identified and validated experimentally. This protocol provides a step-by-step workflow that includes: 1) identification of regulatory regions by ChIP-seq data analysis, 2) cloning and experimental validation of putative regulatory potential of the identified genomic sequences in a reporter assay, and 3) determination of enhancer activity in vivo by measuring enhancer RNA transcript level. The presented protocol is detailed enough to help anyone to set up this workflow in the lab. Importantly, the protocol can be easily adapted to and used in any cellular model system.

In this work we provide an experimental workflow of how active enhancers can be identified and experimentally validated.

Embryonic development is a multistep process involving activation and repression of many genes. Enhancer elements in the genome are known to contribute to ...

Light-mediated Reversible Modulation of the Mitogen-activated Protein Kinase Pathway during Cell Differentiation and Xenopus Embryonic Development

Kinase activity is crucial for a plethora of cellular functions, including cell proliferation, differentiation, migration, and apoptosis. During early embryonic development, kinase activity is highly dynamic and widespread across the embryo. Pharmacological and genetic approaches are commonly used to probe kinase activities. Unfortunately, it is challenging to achieve superior spatial and temporal resolution using these strategies. Furthermore, it is not feasible to control the kinase activity in a reversible fashion in live cells and multicellular organisms. Such a limitation remains a bottleneck for achieving a quantitative understanding of kinase activity during development and differentiation. This work presents an optogenetic strategy that takes advantage of a bicistronic system containing photoactivatable proteins Arabidopsis thaliana cryptochrome 2 (CRY2) and the N-terminal domain of cryptochrome-interacting basic-helix-loop-helix (CIBN). Reversible activation of the mitogen-activated protein kinase (MAPK) signaling pathway is achieved through light-mediated protein translocation in live cells. This approach can be applied to mammalian cell cultures and live vertebrate embryos. This bicistronic system can be generalized to control the activity of other kinases with similar activation mechanisms and can be applied to other model systems.

Light-mediated Reversible Modulation of the Mitogen-activated Protein Kinase Pathway during Cell Differentiation and Xenopus Embryonic Development

This protocol describes an optogenetic strategy to modulate mitogen-activated protein kinase (MAPK) activity during cell differentiation and Xenopus embryonic development. This method allows for the reversible activation of the MAPK signaling pathway in mammalian cell culture and in multicellular live organisms, like Xenopus embryos, with high spatial and temporal resolution.

Kinase activity is crucial for a plethora of cellular functions, including cell proliferation, differentiation, migration, and apoptosis. During early ...

Using In Vivo and Tissue and Cell Explant Approaches to Study the Morphogenesis and Pathogenesis of the Embryonic and Perinatal Aorta

The aorta is the largest artery in the body. The aortic wall is composed of an inner layer of endothelial cells, a middle layer of alternating elastic lamellae and smooth muscle cells (SMCs), and an outer layer of fibroblasts and extracellular matrix. In contrast to the widespread study of pathological models (e.g., atherosclerosis) in the adult aorta, much less is known about the embryonic and perinatal aorta. Here, we focus on SMCs and provide protocols for the analysis of the morphogenesis and pathogenesis of embryonic and perinatal aortic SMCs in normal development and disease. Specifically, the four protocols included are: i) in vivo embryonic fate mapping and clonal analysis; ii) explant embryonic aorta culture; iii) SMC isolation from the perinatal aorta; and iv) subcutaneous osmotic mini-pump placement in pregnant (or non-pregnant) mice. Thus, these approaches facilitate the investigation of the origin(s), fate, and clonal architecture of SMCs in the aorta in vivo. They allow for modulating embryonic aorta morphogenesis in utero by continuous exposure to pharmacological agents. In addition, isolated aortic tissue explants or aortic SMCs can be used to gain insights into the role of specific gene targets during fundamental processes such as muscularization, proliferation, and migration. These hypothesis-generating experiments on isolated SMCs and the explanted aorta can then be assessed in the in vivo context through pharmacological and genetic approaches.

Using In Vivo and Tissue and Cell Explant Approaches to Study the Morphogenesis and Pathogenesis of the Embryonic and Perinatal Aorta

Protocols for studying the embryonic and perinatal murine aorta using in vivo clonal analysis and fate mapping, aortic explants, and isolated smooth muscle cells are detailed here. These diverse approaches facilitate the investigation of the morphogenesis of the embryonic and perinatal aorta in normal development and the pathogenesis in disease.

The aorta is the largest artery in the body. The aortic wall is composed of an inner layer of endothelial cells, a middle layer of alternating elastic ...

Visualization of the Superior Ocular Sulcus during Danio rerio Embryogenesis

Congenital ocular coloboma is a genetic disorder that is typically observed as a cleft in the inferior aspect of the eye resulting from incomplete choroid fissure closure. Recently, the identification of individuals with coloboma in the superior aspect of the iris, retina, and lens led to the discovery of a novel structure, referred to as the superior fissure or superior ocular sulcus (SOS), that is transiently present on the dorsal aspect of the optic cup during vertebrate eye development. Although this structure is conserved across mice, chick, fish, and newt, our current understanding of the SOS is limited. In order to elucidate factors that contribute to its formation and closure, it is imperative to be able to observe it and identify abnormalities, such as delay in the closure of the SOS. Here, we set out to create a standardized series of protocols that can be used to efficiently visualize the SOS by combining widely available microscopy techniques with common molecular biology techniques such as immunofluorescent staining and mRNA overexpression. While this set of protocols focuses on the ability to observe SOS closure delay, it is adaptable to the experimenter&#39;s needs and can be easily modified. Overall, we hope to create an approachable method through which our understanding of the SOS can be advanced to expand the current knowledge of vertebrate eye development.

Visualization of the Superior Ocular Sulcus during Danio rerio Embryogenesis

Here, we present a standardized series of protocols to observe the superior ocular sulcus, a recently-identified, evolutionarily-conserved structure in the vertebrate eye. Using zebrafish larvae, we demonstrate techniques necessary to identify factors that contribute to the formation and closure of the superior ocular sulcus.

Congenital ocular coloboma is a genetic disorder that is typically observed as a cleft in the inferior aspect of the eye resulting from incomplete choroid ...

Analysis of Cardiac Chamber Development During Mouse Embryogenesis Using Whole Mount Epifluorescence

The goal of this protocol is to describe a method for the dissection of mouse embryos and visualization of embryonic mouse ventricular chambers during heart development using ventricular specific fluorescent reporter knock-in mice (MLC-2v-tdTomato mice). Heart development involves a linear heart tube formation, the heart tube looping, and four chamber septation. These complex processes are highly conserved in all vertebrates. The mouse embryonic heart has been widely used for heart developmental studies. However, due to their extremely small size, dissecting mouse embryonic hearts is technically challenging. In addition, visualization of cardiac chamber formation often needs in situ hybridization, beta-galactosidase staining using LacZ reporter mice, or immunostaining of sectioned embryonic hearts. Here, we describe how to dissect mouse embryonic hearts and directly visualize ventricular chamber formation of MLC-2v-tdTomato mice using whole mount epifluorescent microscopy. With this method, it is possible to directly examine heart tube formation and looping, and four chamber formation without further experimental manipulation of mouse embryos. Although the MLC-2v-tdTomato reporter knock-in mouse line is used in this protocol as an example, this protocol can be applied to other heart-specific fluorescent reporter transgenic mouse lines.

We present the protocols to examine mouse heart development using whole mount epifluorescent microscopy on mouse embryos dissected from ventricular specific MLC-2v-tdTomato reporter knock-in mice. This method allows us to directly visualize each stage of the ventricular formation during mouse heart development without labor-intensive histochemical methods.

The goal of this protocol is to describe a method for the dissection of mouse embryos and visualization of embryonic mouse ventricular chambers during ...

Isotropic Light-Sheet Microscopy and Automated Cell Lineage Analyses to Catalogue Caenorhabditis elegans Embryogenesis with Subcellular Resolution

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous system can be observed, with single cell resolution, in vivo. Here, we present an integrated protocol for the examination of neurodevelopment in C. elegans embryos. Our protocol combines imaging, lineaging and neuroanatomical tracing of single cells in developing embryos. We achieve long-term, four-dimensional (4D) imaging of living C. elegans&#160;embryos with nearly isotropic spatial resolution through the use of Dual-view Inverted Selective Plane Illumination Microscopy (diSPIM). Nuclei and neuronal structures in the nematode embryos are imaged and isotropically fused to yield images with resolution of ~330 nm in all three dimensions. These minute-by-minute high-resolution 4D data sets are then analyzed to correlate definitive cell-lineage identities with gene expression and morphological dynamics at single-cell and subcellular levels of detail. Our protocol is structured to enable modular implementation of each of the described steps and enhance studies on embryogenesis, gene expression, or neurodevelopment.

Here, we present a combinatorial approach using high-resolution microscopy, computational tools, and single-cell labeling in living C. elegans embryos to understand single cell dynamics during neurodevelopment.

Caenorhabditis elegans (C. elegans) stands out as the only organism in which the challenge of understanding the cellular origins of an entire nervous ...