This work is supported by the Eunice Kennedy Shriver National Institute of Child Health and Human Development, National Institutes of Health (K25HD097288, R21HD112663).

The protocol has been approved by the Institutional Animal Care and Use Committee of Wayne State University.
1. Experimental preparation
<ol>
	<li>Use a 70% ethanol solution to clean and sterilize the scissors and tweezers. Also, prepare disposable pipettes and a syringe.</li>
	<li>Prepare a wash medium by adding 3.595 g of NaCl to 495 mL of deionized water. Then, add 5 ml of Penicillin-Streptomycin (5 U/mL) to the medium. Fill a 100 mm Petri dish with the wash medium and warm it to 37 &#176;C.</li>
	<li>Prepare culture dishes according to Figure 1, which illustrates the overall configuration.
	<ol>
		<li>Prepare a piece of filter paper, cut it into a rectangular shape of approximately 17 mm &#215; 20 mm, and remove the four corners. Create a hollow center in the filter paper, approximately 7 mm &#215; 10 mm in size for securing the embryo (Figure 1, left). 
		NOTE: The collection of embryos is described in step 2.</li>
		<li>Attach a commercial available ring (&#934; = 1 inch, see Table of Materials) with a flexible film (i.e., paraffin film) layer, ensuring that the flexible film also has a hollow center. This ring with the film will be used to hold the filter paper with the embryo.</li>
		<li>Finally, place the prepared ring into a 35 mm Petri dish (Figure 1, right). 
		&#8203;NOTE: The filter paper is for securing and holding the extracted embryo by placing the filter paper on the membrane and leaving the embryo in the center hollow area (which will be detailed in step 2). The four corners were cut to fit the size of the outside ring (not necessary if it is already fitted). A 35 mm glass-bottom culture dish is used for better laser beam propagation. The dish has an inner well (&#934; = 23 mm), which can be filled with albumen for ex ovo culture. The diameter of the ring must be larger than the diameter of the inner well so that the well can be covered by the flexible film on the ring (Figure 1, inset). The size of the filter paper is not restricted and can be modified according to the size of the chosen ring and the culture dish. Alternatively, the bottom of the dish can be pre-coated with an agarose layer (thickness: ~1 mm). By removing agarose from the center of the layer using a blade, a well with a similar shape to the hollow center of the flexible film can be created for loading albumen. One critical point is that the four sides of the hollow filter paper need to have enough width (&#62; 5 mm) to ensure the attachment of the embryo.</li>
	</ol>
	</li>
</ol>
2. Extraction and ex ovo culture of the chicken embryo
NOTE: This step is modified from previously published reports40,41.
<ol>
	<li>After pre-culture, retrieve the egg from the incubator and place it on its long axis on the egg carton tray. Clean the eggshell with 70% ethanol, ensuring to cover the entire surface, and then allow it to rest for 15 min.</li>
	<li>Hold the egg on its short axis and crack it at the bottom. Open the egg over a clean 100 mm Petri dish to extract its contents, as illustrated in Figure 2. Ensure not to rotate the egg while holding and cracking it.</li>
	<li>Using a pipette, transfer and collect approximately 10 mL of the thin albumen (i.e., the liquid albumen42) into a 15 mL tube for ex ovo culture use. Fill the center well of the culture dish with the collected thin albumen (about 0.9 mL), as shown in Figure 3. The filling process should be slow, avoiding the formation of any bubbles inside the well. Ensure that the culture dish with albumen is warmed to 37 &#176;C. 
	NOTE: This dish will be used for culturing the embryo ex ovo. New culture dishes filled with wash medium will be used during Brillouin imaging (see step 3).</li>
	<li>Using tissue paper, gently remove the thick albumen (i.e., viscous albumen) attached to the embryo by carefully separating the albumen from the vitelline membrane, as demonstrated in Figure 4. Avoid direct contact with the vitelline membrane.</li>
	<li>After removing all the thick albumen, carefully affix the filter paper to the vitelline membrane. Ensure the body axis of the embryo aligns with the long axis of the center rectangle on the filter paper. Use scissors to cut the membrane surrounding the filter paper.</li>
	<li>Using tweezers, gently pull the isolated filter paper away from the yolk in an oblique direction. Flip the filter paper upside down to position the embryo with its dorsal side down (for inverted microscope configuration). Carefully immerse the entire filter paper from one side of the long axis into the 100 mm Petri dish with the wash medium in an oblique fashion.</li>
	<li>Wash away any remaining yolk by gently spraying the wash medium parallel to the filter paper using a clean pipette. Avoid directly spraying the wash medium onto the membrane.</li>
	<li>After clearing all the yolk, carefully remove the filter paper from the wash medium and use tissue paper to absorb any excess medium from the edges. Then, place the filter paper with the embryo onto the culture dish, ensuring that the dorsal side of the embryo is facing downward, as illustrated in Figure 5. To maintain humidity, place a moistened tissue paper in the dish.</li>
	<li>Transfer the culture dish to the on-stage incubator for ex ovo culture.</li>
</ol>
3. Brillouin measurement of the embryo
<ol>
	<li>Prepare another set of culture dishes and fill them with wash medium. Ensure the wash medium is warmed to 37 &#176;C.</li>
	<li>When the embryo reaches the desired developmental stage, transfer the filter paper with the embryo to the culture dish filled with wash medium. Place the culture dish into the on-stage incubator of the Brillouin microscope (see Table of Materials).
	<ol>
		<li>Before conducting the Brillouin measurement, save a bright-field image of the entire embryo for reference. The detailed configuration of the Brillouin microscope has been previously described30 and is summarized in the Results section (Figure 6).</li>
	</ol>
	</li>
	<li>Measure the Brillouin signal of water and methanol, which will be used in step 4 for the calibration process.</li>
	<li>Adjust the incident laser power and electron-multiplying charge-coupled device (EMCCD, see Table of Materials) camera exposure time to achieve at least 10,000 counts of Brillouin signal. Using the bright-field image as a guide, set up the scan range and step size. Acquire a Brillouin image of the region of interest by scanning the embryo using a 2D translational stage. 
	NOTE: The horizontal scanning range can be determined based on the bright-field image, and the vertical (i.e., depth) scanning range can be determined based on the signal strength obtained from a quick and coarse scan. To prevent any photodamage to the embryo, limit the incident power to 25 mW, and set the exposure time of the EMCCD camera to 50 ms. Choose a step size of 2 &#181;m in the horizontal direction and 1 &#181;m in the vertical direction to balance imaging quality and acquisition time.</li>
	<li>After completing the scan, carefully remove the filter paper with the embryo and use tissue paper to absorb any excess wash medium. Then, place the filter paper back into the culture dish filled with thin albumen for continuous culturing in the on-stage incubator.</li>
	<li>Repeat steps 3.2-3.5 at regular time intervals (e.g., 1.5 h) to capture time-lapse Brillouin images as the embryo develops.</li>
	<li>Reconstruct the 2D Brillouin image following step 4.</li>
</ol>
4. Brillouin image reconstruction
<ol>
	<li>Calibrate the Brillouin spectrometer using Brillouin signals of water and methanol to calculate the free spectral range (FSR) and the pixel-to-frequency conversion ratio (PR) of the spectrometer30. The calibrated values of FSR and PR will be used to calculate the Brillouin shift of the embryo at each pixel. 
	NOTE: Figure 7A displays a raw Brillouin spectrum captured by the EMCCD camera. By vertically summing the spectrum and then doing a Lorentzian fitting, the peak distance &#916;d of the two dots can be obtained (Figure 7B). By getting the peak distance of water &#916;dwater and methanol &#916;dmethanol, the FSR and PR can be computed based on the equations30:&#160;PR = 2&#183;(&#969;methanol-&#969;water)/(&#916;dwater- &#916;dmethanol), and FSR = 2&#183;&#969;water&#160;+ PR&#183;&#916;dwater, with the known Brillouin shift of water &#969;water = 6.01 GHz and methanol &#969;methanol = 4.49 GHz at 660 nm.</li>
	<li>Obtain the peak distance of the Brillouin signal at each pixel of the sample. Calculate the Brillouin shift based on the calibrated FSR and PR: &#969;sample = 0.5 (FSR - PR x &#916;dsample)30, where &#969;sample is the Brillouin shift of the sample, and &#916;dsample is the peak distance of the Brillouin signal.</li>
	<li>Reconstruct the 2D Brillouin image based on the Brillouin shifts of all pixels. 
	NOTE: To conduct local analysis, one can select a region of interest (e.g., the neural plate) from the acquired Brillouin image and quantify its mechanical properties. A common approach is to calculate the average Brillouin shift of the selected region.</li>
</ol>

Longitudinal Monitoring of Neural Plate Mechanics in Chick Embryo Development

The authors declare that they have no conflict of interest.

The early development of the embryo can be easily affected by external disturbances. Therefore, utmost caution is required during the sample extraction and transfer. One potential issue is the detachment of the embryo from the filter paper, which can lead to the shrinking of the vitelline membrane and result in a tilted artifact of the neural plate in Brillouin imaging. Furthermore, this shrinking may halt the development of the embryo. Attention should be paid to several critical steps to prevent detachment. First, in s...

Neural tube defects (NTDs) are severe birth defects of the central nervous system caused by failures in neural tube closure (NTC) during embryonic development1. The etiology of NTDs is complex. Studies have shown that NTC involves a sequence of morphogenetic processes, including convergent extension, bending of the neural plate (e.g., apical constriction), elevating the neural fold, and finally adhesion of the neural fold. These processes are regulated by multiple molecular and genetic mechanisms2,3, and any malfunction in these processes may result in NTDs4,5,6. As mounting evidence suggests that mechanical cues also play crucial roles during NTC3,7,8,9,10,11, and relationships have been found between genes and mechanical cues12,13,14, it becomes imperative to investigate the tissue biomechanics during neurulation.
Several techniques have been developed for measuring the mechanical properties of embryonic tissues, including laser ablation (LA)15, tissue dissection and relaxation (TDR)16,17, micropipette aspiration (MA)18, Atomic Force Microscopy (AFM)-based nanoindentation19, microindenters (MI)&#160;and microplates (MP)20, micro rheology (MR) with optical/magnetic tweezers21,22,23, and droplet-based sensors24. Existing methods can measure mechanical properties at spatial resolutions ranging from subcellular to tissue scales. However, most of these methods are invasive because they require contact with the sample (e.g., MA, AFM, MI, and MP), external material injection (e.g., MR and droplet-based sensors), or tissue dissection (e.g., LA and TDR). As a result, it is challenging for existing methods to monitor the mechanical evolution of neural plate tissue in situ25. Recently, reverberant optical coherence elastography has shown promise for non-contact mechanical mapping with high spatial resolution26.
Confocal Brillouin microscopy is an emerging optical modality that enables non-contact quantification of tissue biomechanics with subcellular resolution27,28,29,30. Brillouin microscopy is based on the principle of spontaneous Brillouin light scattering, which is the interaction between the incident laser light and the acoustic wave induced by thermal fluctuations within the material. Consequently, the scattered light experiences a frequency shift, known as the Brillouin shift &#969;R, following the equation31:
<img alt="Equation 1" src="/files/ftp_upload/66117/66117eq01.jpg" style="margin: auto;" /> (1)
Here, <img alt="Equation 2" src="/files/ftp_upload/66117/66117eq02.jpg" style="margin: auto;" /> is the refractive index of the material,&#160;&#955; is the wavelength of the incident light, M&#39; is the longitudinal modulus, &#961; is the mass density, and &#952; is the angle between the incident light and the scattered light. For the same type of biological materials, the ratio of refractive index and density <img alt="Equation 3" src="/files/ftp_upload/66117/66117eq03.jpg" style="margin: auto;" /> is approximately constant28,32,33,34,35,36. Thus, the Brillouin shift can be directly used to estimate relative mechanical changes in physiological processes. The feasibility of Brillouin microscopy has been validated in various biological samples29,37,38. Recently, time-lapse mechanical imaging of a live chick embryo was demonstrated by combining a Brillouin microscope with an on-stage incubation system39. This protocol provides detailed descriptions of sample preparation, experiment implementation, and data post-processing and analysis. We hope this effort will facilitate the widespread adoption of non-contact Brillouin technology for studying biomechanical regulation in embryo development and birth defects.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>100 mm Petri dish&#160;</td>
			<td>Fisherbrand</td>
			<td>FB0875713</td>
			<td></td>
		</tr>
		<tr>
			<td>2D motorized stage&#160;</td>
			<td>Prior Scientific</td>
			<td>H117E2</td>
			<td></td>
		</tr>
		<tr>
			<td>35 mm Petri dish</td>
			<td>World Precision Instruments</td>
			<td>FD35-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Brillouin Microscope with on-stage incubator</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>This is a custom-built Brillouin Microscope system based on Ref. 30</td>
		</tr>
		<tr>
			<td>Chicken eggs</td>
			<td>University of Connecticut</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>CMOS camera</td>
			<td>Thorlabs</td>
			<td>CS2100M-USB</td>
			<td></td>
		</tr>
		<tr>
			<td>EMCCD camera</td>
			<td>Andor</td>
			<td>iXon</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Decon Laboratories, Inc.</td>
			<td>#2701</td>
			<td></td>
		</tr>
		<tr>
			<td>Filter paper</td>
			<td>Whatman</td>
			<td>1004-070</td>
			<td></td>
		</tr>
		<tr>
			<td>Incubator for in ovo culture</td>
			<td>GQF Manufacturing Company Inc.&#160;</td>
			<td>GQF 1502&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>Ring</td>
			<td>Thorlabs</td>
			<td>SM1RR</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope body</td>
			<td>Olympus</td>
			<td>IX73</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma-Aldrich</td>
			<td>S9888</td>
			<td></td>
		</tr>
		<tr>
			<td>On-stage incubator</td>
			<td>Oko labs</td>
			<td>OKO-H301-PRIOR-H117</td>
			<td></td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>Bemis</td>
			<td>PM-996</td>
			<td></td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin</td>
			<td>Gibco</td>
			<td>15070-063</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipettes</td>
			<td>Fisherbrand</td>
			<td>13-711-6M</td>
			<td></td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>Artman instruments</td>
			<td>N/A</td>
			<td>3pc Micro Scissors 5</td>
		</tr>
		<tr>
			<td>Syringe</td>
			<td>BD</td>
			<td>305482</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue paper</td>
			<td>Kimwipes</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Tube</td>
			<td>Corning</td>
			<td>430052</td>
			<td></td>
		</tr>
		<tr>
			<td>Tweezers</td>
			<td>DR Instruments</td>
			<td>N/A</td>
			<td>Microdissection Forceps Set&#160;</td>
		</tr>
	</tbody>
</table>

monitoring the mechanical evolution of tissue during neural tube closure of chick embryo

Neural tube closure (NTC) is a critical process during embryonic development. Failure in this process can lead to neural tube defects, causing congenital malformations or even mortality. NTC involves a series of mechanisms on genetic, molecular, and mechanical levels. While mechanical regulation has become an increasingly attractive topic in recent years, it remains largely unexplored due to the lack of suitable technology for conducting mechanical testing of 3D embryonic tissue in situ. In response, we have developed a protocol for quantifying the mechanical properties of chicken embryonic tissue in a non-contact and non-invasive manner. This is achieved by integrating a confocal Brillouin microscope with an on-stage incubation system. To probe tissue mechanics, a pre-cultured embryo is collected and transferred to an on-stage incubator for ex ovo culture. Simultaneously, the mechanical images of the neural plate tissue are acquired by the Brillouin microscope at different time points during development. This protocol includes detailed descriptions of sample preparation, the implementation of Brillouin microscopy experiments, and data post-processing and analysis. By following this protocol, researchers can study the mechanical evolution of embryonic tissue during development longitudinally.

Figure 6 shows the schematic of the Brillouin microscope. The system employs a 660 nm laser as the light source. An isolator is placed right after the laser head to reject any back-reflected light, and a neutral density (ND) filter is used to adjust the laser power. A pair of lenses, L1 and L2, with focal lengths of f1 = 16 mm and f2 = 100 mm, respectively, are used to expand the laser beam. A half-wave plate (HWP) and a linear polarizer (Polarizer 1) are employed to adjust the power of the ...

Watch this Scientific Journal Video about Monitoring the Mechanical Evolution of Tissue During Neural Tube Closure of Chick Embryo at JoVE.com

Author Spotlight: Non-Contact Measurement of Tissue Mechanics in Live Chick Embryos Using Brillouin Microscopy

This protocol was developed to longitudinally monitor the mechanical properties of neural plate tissue during chick embryo neurulation. It is based on the integration of a Brillouin microscope and an on-stage incubation system, enabling live mechanical imaging of neural plate tissue in ex ovo cultured chick embryos.

Monitoring the Mechanical Evolution of Tissue During Neural Tube Closure of Chick Embryo

Neural tube closure (NTC) is a critical process during embryonic development. Failure in this process can lead to neural tube defects, causing congenital ...

monitoring-mechanical-evolution-tissue-during-neural-tube-closure

Research

JoVE Journal

Developmental Biology

541 Views.  Wayne State University. This protocol was developed to longitudinally monitor the mechanical properties of neural plate tissue during chick embryo neurulation. It is based on the integration of a Brillouin microscope and an on-stage incubation system, enabling live mechanical imaging of neural plate tissue in ex ovo cultured chick embryos.

Video: Author Spotlight: Non-Contact Measurement of Tissue Mechanics in Live Chick Embryos Using Brillouin Microscopy

Dual Labeling of Neural Crest Cells and Blood Vessels Within Chicken Embryos Using ChickGFP Neural Tube Grafting and Carbocyanine Dye DiI Injection

All developing organs need to be connected to both the nervous system (for sensory and motor control) as well as the vascular system (for gas exchange, fluid and nutrient supply). Consequently both the nervous and vascular systems develop alongside each other and share striking similarities in their branching architecture. Here we report embryonic manipulations that allow us to study the simultaneous development of neural crest-derived nervous tissue (in this case the enteric nervous system), and the vascular system. This is achieved by generating chicken chimeras via transplantation of discrete segments of the neural tube, and associated neural crest, combined with vascular DiI injection in the same embryo. Our method uses transgenic chickGFP embryos for intraspecies grafting, making the transplant technique more powerful than the classical quail-chick interspecies grafting protocol used with great effect since the 1970s. ChickGFP-chick intraspecies grafting facilitates imaging of transplanted cells and their projections in intact tissues, and eliminates any potential bias in cell development linked to species differences. This method takes full advantage of the ease of access of the avian embryo (compared with other vertebrate embryos) to study the co-development of the enteric nervous system and the vascular system.

Dual Labeling of Neural Crest Cells and Blood Vessels Within Chicken Embryos Using ChickGFP Neural Tube Grafting and Carbocyanine Dye DiI Injection

Here we report dual labeling of neural crest cells and blood vessels using chickGFP neural tube intraspecies grafting combined with intra-vascular DiI injection. This experimental technique allows us to simultaneously visualize and study development of the NCC-derived (enteric) nervous system and the vascular system, during organogenesis.

All developing organs need to be connected to both the nervous system (for sensory and motor control) as well as the vascular system (for gas exchange, ...

Forward Genetic Approach to Uncover Stress Resistance Genes in Mice &#8212; A High-throughput Screen in ES Cells

Phenotype-driven genetic screens in mice is a powerful technique to uncover gene functions, but are often hampered by extremely high costs, which severely limits its potential. We describe here the use of mouse embryonic stem (ES) cells as surrogate cells to screen for a phenotype of interest and subsequently introduce these cells into a host embryo to develop into a living mouse carrying the phenotype. This method provides (1) a cost effective, high-throughput platform for genetic screen in mammalian cells; (2) a rapid way to identify the mutated genes and verify causality; and (3) a short-cut to develop mouse mutants directly from these selected ES cells for whole animal studies. We demonstrated the use of paraquat (PQ) to select resistant mutants and identify mutations that confer oxidative stress resistance. Other stressors or cytotoxic compounds may also be used to screen for resistant mutants to uncover novel genetic determinants of a variety of cellular stress resistance.

Forward Genetic Approach to Uncover Stress Resistance Genes in Mice &amp;#8212; A High-throughput Screen in ES Cells

Stress resistance is one of the hallmarks for longevity and is known to be genetically governed. Here, we developed an unbiased high-throughput method to screen for mutations that confer stress resistance in ES cells with which to develop mouse models for longevity studies.

Phenotype-driven genetic screens in mice is a powerful technique to uncover gene functions, but are often hampered by extremely high costs, which severely ...

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance to man of conventional invertebrate model systems and the complexity of genomes in vertebrates are critical challenges to modeling human normal and pathological conditions. The chordate Ciona intestinalis is an invertebrate chordate that emerged from a common ancestor with the vertebrates and may represent features at the interface between invertebrates and vertebrates. A common body plan with much simpler cellular and genomic composition should unveil gene regulatory network (GRN) links and functional genomics readouts explaining phenomena in the vertebrate condition. The compact genome of Ciona, a fixed embryonic lineage with few divisions and large cells, combined with versatile community tools foster efficient gene functional analyses in this organism. Here, we present several crucial methods for this promising model organism, which belongs to the closest sister group to vertebrates. We present protocols for transient transgenesis by electroporation, along with microinjection-mediated gene knockdown, which together provide the means to study gene function and genomic regulatory elements. We extend our protocols to provide information on how community databases are utilized for in silico design of gene regulatory or gene functional experiments. An example study demonstrates how novel information can be gained on the interplay, and its quantification, of selected neural factors conserved between Ciona and man. Furthermore, we show examples of differential subcellular localization in embryonic cells, following DNA electroporation in Ciona zygotes. Finally, we discuss the potential of these protocols to be adapted for tissue specific gene interference with emerging gene editing methods. The in vivo approaches in Ciona overcome major shortcomings of classical model organisms in the quest of unraveling conserved mechanisms in the chordate developmental program, relevant to stem cell research, drug discovery, and subsequent clinical application.

Embryo Microinjection and Electroporation in the Chordate Ciona intestinalis

We present transient transgenesis and gene knockdown in Ciona intestinalis, a chordate sister group to vertebrates, using microinjection and electroporation techniques. Such methods facilitate functional genomics in this simple invertebrate that features rudimentary characteristics of vertebrates, including notochord and head sensory epithelia, and many orthologs of human disease associated genes.

Simple model organisms are instrumental for in vivo studies of developmental and cellular differentiation processes. Currently, the evolutionary distance ...

Effectiveness of the Air Stripping in Two Salmonid Fish, Rainbow Trout (Oncorhynchus Mykiss) and Brown Trout (Salmo Trutta Morpha fario)

Egg collection is one of the most crucial procedures during fish reproduction in salmonid hatcheries. Classic methods involve the use of hand massage on fish abdomens to expel the eggs. An alternative method uses the pressure of gas injected into the body cavity, which causes the subsequent release of the eggs. This method is believed to have less negative effects on both the welfare and egg quality of the broodstocks. Herein, we compare the results of air and hand stripping methods with respect to one-year survival and egg quantity and quality in two salmonid fish, rainbow trout (Oncorhynchus mykiss) and brown trout (Salmo trutta morpha fario). Our results indicate that air stripping yielded a better quality of eggs and higher one-year survival rate in rainbow trout. In addition, air stripping resulted in lower mortality rate than the group subjected to hand stripping (25% vs. 35%). The pH and hatching rate of the hand stripped group was lower than those of the air stripped group. In the case of brown trout, the quality of eggs obtained by both hand and air-stripping methods was similar; however, the one-year losses in fish were higher in air stripped group (15% compared to 0% in hand stripped fish). Although the advantages of air stripping method over hand stripping in terms of egg quality might not be observed in all salmonid species, the air-stripping procedure might be a promising option to be adopted in hatcheries as it ensures a high level of reproducibility and efficiency.

Effectiveness of the Air Stripping in Two Salmonid Fish, Rainbow Trout Oncorhynchus Mykiss and Brown Trout Salmo Trutta Morpha fario

The principal aim of this study is to standardize and test the pneumatic method (air stripping) of collecting eggs in rainbow trout and brown trout. This method allows effective and simple collection of the eggs without the necessity of fish abdomen massage.

Egg collection is one of the most crucial procedures during fish reproduction in salmonid hatcheries. Classic methods involve the use of hand massage on ...

Following Endocardial Tissue Movements via Cell Photoconversion in the Zebrafish Embryo

During embryogenesis, cells undergo dynamic changes in cell behavior, and deciphering the cellular logic behind these changes is a fundamental goal in the field of developmental biology. The discovery and development of photoconvertible proteins have greatly aided our understanding of these dynamic changes by providing a method to optically highlight cells and tissues. However, while photoconversion, time-lapse microscopy, and subsequent image analysis have proven to be very successful in uncovering cellular dynamics in organs such as the brain or the eye, this approach is generally not used in the developing heart due to challenges posed by the rapid movement of the heart during the cardiac cycle. This protocol consists of two parts. The first part describes a method for photoconverting and subsequently tracking endocardial cells (EdCs) during zebrafish atrioventricular canal (AVC) and atrioventricular heart valve development. The method involves temporally stopping the heart with a drug in order for accurate photoconversion to take place. Hearts are allowed to resume beating upon removal of the drug and embryonic development continues normally until the heart is stopped again for high-resolution imaging of photoconverted EdCs at a later developmental time point. The second part of the protocol describes an image analysis method to quantify the length of a photoconverted or non-photoconverted region in the AVC in young embryos by mapping the fluorescent signal from the three-dimensional structure onto a two-dimensional map. Together, the two parts of the protocol allows one to examine the origin and behavior of cells that make up the zebrafish AVC and atrioventricular heart valve, and can potentially be applied for studying mutants, morphants, or embryos that have been treated with reagents that disrupt AVC and/or valve development.

This protocol describes a method for the photoconversion of Kaede fluorescent protein in endocardial cells of the living zebrafish embryo that enables the tracking of endocardial cells during atrioventricular canal and atrioventricular heart valve development.

During embryogenesis, cells undergo dynamic changes in cell behavior, and deciphering the cellular logic behind these changes is a fundamental goal in the ...

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A remarkable example is given by sensory epithelia, where cells of the same or distinct identities are brought together via cell-cell adhesion showing highly organized planar patterns. Cells align to one another in the same direction and display equivalent polarity over large distances. This organization of the mature epithelia is established over the course of morphogenesis. To understand how the planar arrangement of the mature epithelia is achieved, it is crucial to track cell orientation and growth dynamics with high spatiotemporal fidelity during development in vivo. Robust analytical tools are also essential to identify and characterize local-to-global transitions. The Drosophila pupa is an ideal system to evaluate oriented cell shape changes underlying epithelial morphogenesis. The pupal developing epithelium constitutes the external surface of the immobile body, allowing long-term imaging of intact animals. The protocol described here is designed to image and analyze cell behaviors at both global and local levels in the pupal abdominal epidermis as it grows. The methodology described can be easily adapted to the imaging of cell behaviors at other developmental stages, tissues, subcellular structures, or model organisms.

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

This protocol is designed for the imaging and analysis of the dynamics of cell orientation and tissue growth in the Drosophila abdominal epithelia as the fruit fly undergoes metamorphosis. The methodology described here can be applied to the study of different developmental stages, tissues, and subcellular structures in Drosophila or other model organisms.

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A ...

An Optimized O9-1/Hydrogel System for Studying Mechanical Signals in Neural Crest Cells

Neural crest cells (NCCs) are vertebrate embryonic multipotent cells that can migrate and differentiate into a wide array of cell types that give rise to various organs and tissues. Tissue stiffness produces mechanical force, a physical cue that plays a critical role in NCC differentiation; however, the mechanism remains unclear. The method described here provides detailed information for the optimized generation of polyacrylamide hydrogels of varying stiffness, the accurate measurement of such stiffness, and the evaluation of the impact of mechanical signals in O9-1 cells, a NCC line that mimics in vivo NCCs.
Hydrogel stiffness was measured using atomic force microscopy (AFM) and indicated different stiffness levels accordingly. O9-1 NCCs cultured on hydrogels of varying stiffness showed different cell morphology and gene expression of stress fibers, which indicated varying biological effects caused by mechanical signal changes. Moreover, this established that varying the hydrogel stiffness resulted in an efficient in vitro system to manipulate mechanical signaling by altering gel stiffness and analyzing the molecular and genetic regulation in NCCs. O9-1 NCCs can differentiate into a wide range of cell types under the influence of the corresponding differentiation media, and it is convenient to manipulate chemical signals in vitro. Therefore, this in vitro system is a powerful tool to study the role of mechanical signaling in NCCs and its interaction with chemical signals, which will help researchers better understand the molecular and genetic mechanisms of neural crest development and diseases.

Detailed step-by-step protocols are described here for studying mechanical signals in vitro using multipotent O9-1 neural crest cells and polyacrylamide hydrogels of varying stiffness.

Neural crest cells (NCCs) are vertebrate embryonic multipotent cells that can migrate and differentiate into a wide array of cell types that give rise to ...

Investigating Scarless Tissue Regeneration in Embryonic Wounded Chick Corneas

Chick embryonic corneal wounds display a remarkable capacity to fully and rapidly regenerate, whereas adult wounded corneas experience a loss of transparency due to fibrotic scarring. The tissue integrity of injured embryonic corneas is intrinsically restored with no detectable scar formation. Given its accessibility and ease of manipulation, the chick embryo is an ideal model for studying scarless corneal wound repair. This protocol demonstrates the different steps involved in wounding the cornea of an embryonic chick in ovo. First, eggs are windowed at early embryonic ages to access the eye. Second, a series of in ovo physical manipulations to the extraembryonic membranes are conducted to ensure access to the eye is maintained through later stages of development, corresponding to when the three cellular layers of the cornea are formed. Third, linear cornea wounds that penetrate the outer epithelial layer and the anterior stroma are made using a microsurgical knife. The regeneration process or fully restored corneas can be analyzed for regenerative potential using various cellular and molecular techniques following the wounding procedure. Studies to date using this model have revealed that wounded embryonic corneas display activation of keratocyte differentiation, undergo coordinated remodeling of ECM proteins to their native three-dimensional macrostructure, and become adequately re-innervated by corneal sensory nerves. In the future, the potential impact of endogenous or exogenous factors on the regenerative process could be analyzed in healing corneas by using developmental biology techniques, such as tissue grafting, electroporation, retroviral infection, or bead implantation. The current strategy identifies the embryonic chick as a crucial experimental paradigm for elucidating the molecular and cellular factors coordinating scarless corneal wound healing.

The present protocol demonstrates the different steps involved in wounding the cornea of an embryonic chick in ovo. The regenerating or fully restored corneas can be analyzed for regenerative potential using various cellular and molecular techniques following the wounding procedure.&#160;

Chick embryonic corneal wounds display a remarkable capacity to fully and rapidly regenerate, whereas adult wounded corneas experience a loss of ...

Preparation and Morphological Analysis of Chick Cranial Neural Crest Cell Cultures

During vertebrate development, neural crest cells (NCCs) migrate extensively and differentiate into various cell types that contribute to structures like the craniofacial skeleton and the peripheral nervous system. While it is critical to understand NCC migration in the context of a 3D embryo, isolating migratory cells in 2D culture facilitates visualization and functional characterization, complementing embryonic studies. The present protocol demonstrates a method for isolating chick cranial neural folds to generate primary NCC cultures. Migratory NCCs emerge from neural fold explants plated onto a fibronectin-coated substrate. This results in dispersed, adherent NCC populations that can be assessed by staining and quantitative morphological analyses. This simplified culture approach is highly adaptable and can be combined with other techniques. For example, NCC emigration and migratory behaviors can be evaluated by time-lapse imaging or functionally queried by including inhibitors or experimental manipulations of gene expression (e.g., DNA, morpholino, or CRISPR electroporation). Because of its versatility, this method provides a powerful system for investigating cranial NCC development.

This versatile protocol describes the isolation of premigratory neural crest cells (NCCs) through the excision of cranial neural folds from chick embryos. Upon plating and incubation, migratory NCCs emerge from neural fold explants, allowing for assessment of cell morphology and migration in a simplified 2D environment.

During vertebrate development, neural crest cells (NCCs) migrate extensively and differentiate into various cell types that contribute to structures like ...

Avian Embryonic Left Atrial Ligation Model for HLHS and Cardiovascular Research

Left Atrial Ligation in the Avian Embryo as a Model for Altered Hemodynamic Loading During Early Vascular Development

Due to its four-chambered mature ventricular configuration, ease of culture, imaging access, and efficiency, the avian embryo is a preferred vertebrate animal model for studying cardiovascular development. Studies aiming to understand the normal development and congenital heart defect prognosis widely adopt this model. Microscopic surgical techniques are introduced to alter the normal mechanical loading patterns at a specific embryonic time point and track the downstream molecular and genetic cascade. The most common mechanical interventions are left vitelline vein ligation, conotruncal banding, and left atrial ligation (LAL), modulating the intramural vascular pressure and wall shear stress due to blood flow. LAL, particularly if performed in ovo, is the most challenging intervention, with very small sample yields due to the extremely fine sequential microsurgical operations. Despite its high risk, in ovo LAL is very valuable scientifically as it mimics hypoplastic left heart syndrome (HLHS) pathogenesis. HLHS is a clinically relevant, complex congenital heart disease observed in human newborns. A detailed protocol for in ovo LAL is documented in this paper. Briefly, fertilized avian embryos were incubated at 37.5 &#176;C and 60% constant humidity typically until they reached Hamburger-Hamilton (HH) stages 20 to 21. The egg shells were cracked open, and the outer and inner membranes were removed. The embryo was gently rotated to expose the left atrial bulb of the common atrium. Pre-assembled micro-knots from 10-0 nylon sutures were gently positioned and tied around the left atrial bud. Finally, the embryo was returned to its original position, and LAL was completed. Normal and LAL-instrumented ventricles demonstrated statistically significant differences in tissue compaction. An efficient LAL model generation pipeline would contribute to studies focusing on synchronized mechanical and genetic manipulation during the embryonic development of cardiovascular components. Likewise, this model will provide a perturbed cell source for tissue culture research and vascular biology.

Author Spotlight: Effect of Left Atrial Ligation on Avian Embryonic Hearts and HLHS Implications 

Here, we present a detailed visual protocol for executing the left atrial ligation (LAL) model in the avian embryo. The LAL model alters the intracardiac flow, which changes wall shear stress loading, mimicking hypoplastic left heart syndrome. An approach to overcome the challenges of this difficult microsurgery model is presented.

Due to its four-chambered mature ventricular configuration, ease of culture, imaging access, and efficiency, the avian embryo is a preferred vertebrate ...

Methods Article

Monitoring the Mechanical Evolution of Tissue During Neural Tube Closure of Chick Embryo