We acknowledge Tubitak 2247A lead researcher award 120C139 providing funding. The authors would also like to thank PakTavuk G&#305;da. A. S., Istanbul, Turkey, for providing fertile eggs and supporting the cardiovascular research.

Fertile white Leghorn eggs are obtained from trusted suppliers and incubated according to university-approved guidelines. Chick embryos, stages 18 (day 3) to 24 (day 4) (the stages presented in this paper) are not considered live vertebrate animals by the European Union (EU) directive 2010/63/EU and the institutional animal care and use committee (IACUC) guidelines in the US. Chick embryos are considered &#34;live animals&#34; after day 19 of incubation according to US laws, but not for the EU. Each egg is labeled with the hatching start date and is scheduled to hatch no later than the 10th day of incubation. After the eggs hatch, the chicks are removed from the incubator. The protocol is performed in two bench-top operation stations (Station 1 and Station 2), focusing on specialized model generation stages.
1. Preparation before microsurgery
<ol>
	<li>Obtain fertilized eggs either from a vaccine development center with a specific pathogen-free (SPF) grade or through a trusted commercial supplier farm by a fragile delivery courier in dry polystyrene containers. Before incubation, gently clean the eggshell with lint-free wipes soaked in 70% ethanol to remove contamination.</li>
	<li>Embryo inclusion/exclusion criteria
	<ol>
		<li>Do not incubate eggs that were cracked or damaged during transport.</li>
		<li>If bleeding is noted during the LAL procedure or following re-incubation, do not use the embryo.</li>
		<li>Do not use embryos developing in the left side-up position, as hemodynamic blood flow may differ from normal orientation.</li>
		<li>D&#305;scard embryos developing with congenital defects in both pre- and post-surgical procedures.</li>
		<li>Include embryos that reach the targeted stage developing in their original location, mimicking HLHS as a LAL model.</li>
	</ol>
	</li>
	<li>Incubate the fertilized white-Leghorn chicken eggs (Gallus gallus domesticus L.), blunt end-up, to the desired stage typically at HH20-2115 (37.5 &#176;C, 60% humidity, 3.5 days) (Figure 1). 
	NOTE: It is important to keep the eggs at a constant temperature and humidity to increase the yield. Depending on the incubator model, the addition of a pan filled with distilled water will maintain stable humidity. Blueprints of an additional/auxiliary temperature and humidity control system that would fit most incubators are developed by the authors and recommended. The electronics, hardware, and code details of this in-house-built sensor/control unit are provided in a data repository26. Continuous gentle shaking (rotating) of the eggs during incubation may allow optimal positioning of the embryo and thus lead to a higher percentage of &#34;operable&#34; embryos. Shaking can also work with incubators with this capacity and increase productivity further.</li>
	<li>Before starting the procedure, prepare the required number of knots by tying a loose overhand knot into a 1.5 cm long 10-0 suture. Be sure the knot is not tight and is large enough to fit over the atria easily during the operation (Figure 2). 
	NOTE: Tie the knots ahead of time and keep them in a sterile chick ringer solution before using. The knot-tying operation requires the use of two hands to operate the tweezers synchronously. Since this is a critical stage in the protocol, a model of the atrium can be made from putty for practicing this step (Figure 3). This will improve the three-dimensional microsurgery skills needed to perform step 3.2.3 at Station 2 (Figure 4).</li>
</ol>
2. Operations at Station 1 (Figure 4A)
<ol>
	<li>Open a window from the blunt end of the egg and remove both the outer and inner membranes15 (Figure 5A-D).</li>
	<li>Open the eggshell by cracking gently with the reverse end of the tweezers, with the free fingers firmly supporting the egg to reduce unwanted crack propagation.</li>
	<li>Since LAL is a long procedure, conserve the temperature and humidity of the embryo, as the heart rate depends on the temperature. Thus, ensure the initial shell window is created as small as possible, just enough to execute the operations. 
	NOTE: No humidity or temperature control systems are employed during the operation, but these systems, if available, would benefit the yield. If possible, the air conditioning system in the lab is shut-down, and the procedure is performed at the highest room temperature (RT) possible. Optimization of the embryonic heart rate, which can be controlled by the temperature, during the operation is also recommended. Some laboratories maintain the heart rate at slightly lower rates than 120 bpm via temperature control during the LAL operation. As such, humidity control employed around the surgical zone would increase the yield further. The eggshell windows are created as small as possible, just big enough to allow surgical access. This is also applicable to the thick outer membrane, which is typically smaller than the eggshell only to the extent of the embryo&#39;s circumference. These ensure to maintain the temperature and humidity of the embryo. While opening a window from the blunt end of the egg, the small shell fragments are cleaned so that these pieces do not damage the vitelline vascular integrity or lead to unwanted artifacts. In addition, other laboratories use curved micro-serrated scissors for making windows. Also, two widths of Scotch tape can be used to stabilize the eggshell to control cracking.</li>
	<li>Remove only the necessary vitelline membrane using micro scissors (Supplementary Video 1).</li>
	<li>Normal embryonic development is right side-up. Once the embryo is free of the vitelline membrane, place the tweezer with closed tips under the dorsal segment of the embryo and gently flip the embryo to expose the left side (i.e., left side-up configuration) (Figure 6A,B; Supplementary Video 2).</li>
	<li>Ensure that the left atrial bud is now exposed but still covered by a complex membrane system, typically consisting of a double layer of the pericardium.</li>
	<li>Remove the membranes, including the fine ones, immediately around the atrial bud. This is another critical stage; perform membrane removal from coarse membranes and progress to the fine ones around the left atrial bud. Reserve the finest tweezers for removal of the fine membrane (Supplementary Video 3).</li>
	<li>During the membrane removal process, orient the embryo in a left side-up position, so that the knot placement operation in step 3.2 can be performed without further repositioning. To achieve this, lift the embryo using the membranes in step 2.6 and hang it from the eggshell, ensuring the left side faces up. 
	&#8203;NOTE: Some embryos can be located close to the eggshell periphery and can be challenging to operate. Still, these embryos will most likely be oriented right side-up and display normal behavior, and can be included in the study. In these cases, if needed, the obscured atrium can be made more accessible by gently cleaning the pericardial membrane with #4 fine tweezers and removing the eggshell in the reverse direction (toward the shell opening). These embryos can also be lifted using parts of the extraembryonic membranes and fixed at the desired position by attaching one end of the membrane (the tweezer end) to the eggshell, using its natural stickiness. Additionally, the space between the head and spinal cord region of the embryo can be expanded with the help of tweezers to reveal the obscured atrium.</li>
</ol>
3. Operations at Station 2 (Figure 4B)
<ol>
	<li>Under the stereomicroscope, position the pre-prepared knot from step 1.4&#160;close to the embryo at an accessible location (Figure 6B). The atrial bud is now ready to be tied off (Supplementary Video 4).</li>
	<li>Retrieve the pre-prepared open knot and orient it over the left atrial bud. For LAL to work, place the egg in a uniquely inclined three-dimensional orientation.
	<ol>
		<li>Correctly orient the knot to execute the tightening process without damaging the embryos.</li>
		<li>Clear the fine membranes optimally in step 2.7 to reduce the effect of a beating heart.</li>
		<li>Tighten the suture (Figure 6C). For this step, practicing with the putty is very useful. For sham embryos, tighten the knot just enough to hold the knot.</li>
	</ol>
	</li>
	<li>Next, use the micro scissors to cut the excess ends of the suture as close as possible to the bud (Supplementary Video 5).</li>
	<li>Be very careful that the newly cut ends of the tied suture are not in a position to puncture nearby vessels during rotation or due to the heartbeat.</li>
	<li>Using the tweezers, remove the excess suture pieces cut away in step 3.3.</li>
	<li>Finally, using closed tweezers, return the embryo to its original position, as in step 2.5&#160;(Supplementary Video 6).</li>
	<li>After completing the LAL process, cover the egg with a double layer of parafilm and incubate it again. A tight and sterile closing of the eggs is paramount for survival, especially past 24 h of incubation. If visual access is also desired, use paraffin wax with glass slides. 
	NOTE: As the early embryonic period is studied, the eggs are typically incubated for 24-48 h until they reach approximately HH25 or HH27. However, there is no limit, and later stages can be studied, as attempted by other researchers. For fast operation speeds, at least a two-person team is recommended. One person should be trained for and is responsible for egg opening, initial membrane cleanup, rotation, and membrane cleanup around the left atrial bud. The other person is responsible only for initial knot preparation, knot placement, and tightening. The final embryo rotation can be performed by Person 1. The surgical operation for a single embryo takes about 4-5 min.</li>
	<li>Before/after surgical operations, clean the bench-top surfaces and instruments with ethanol. Ensure to apply fresh chick ringer solution (NaCl, KCl, CaCl2, and NaHCO3)16,27 to the metal instruments touching the embryonic tissues.</li>
</ol>

Avian Embryonic Left Atrial Ligation Model for HLHS and Cardiovascular Research

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Y., 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=Induction+of+left+ventricular+hypoplasia+by+occluding+the+foramen+ovale+in+the+fetal+lamb.">Induction of left ventricular hypoplasia by occluding the foramen ovale in the fetal lamb.</a> Scientific Reports. 10 (1), 880 (2020).</li><li>Nie, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+frogs+as+a+model+to+study+the+etiology+of+HLHS.">Use of frogs as a model to study the etiology of HLHS.</a> Journal of Cardiovascular Development and Disease. 10 (2), 51 (2023).</li><li>Vilches-Moure, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Embryonic+chicken+(Gallus+gallus+domesticus)+as+a+model+of+cardiac+biology+and+development.">Embryonic chicken (Gallus gallus domesticus) as a model of cardiac biology and development.</a> Comparative Medicine. 69 (3), 184-203 (2019).</li><li>Kain, K. H., 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+chick+embryo+as+an+expanding+experimental+model+for+cancer+and+cardiovascular+research.">The chick embryo as an expanding experimental model for cancer and cardiovascular research.</a> Developmental Dynamics. 243 (2), 216-228 (2014).</li><li>Sukparangsi, W., Thongphakdee, A., Intarapat, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Avian+embryonic+culture:+A+perspective+of+in+ovo+to+ex+ovo+and+in+vitro+studies.">Avian embryonic culture: A perspective of in ovo to ex ovo and in vitro studies.</a> Frontiers in Physiology. 13, 903491 (2022).</li></ol>

The authors have nothing to disclose.

In HLHS, blood flow is altered due to structural defects, leading to abnormal morphology on the left side4,6. The present model provides a practical experimental system to better understand the progression of HLHS and may even mimic its pathogenesis8. However, establishing a fully clinically equivalent HLHS animal model is a challenging task. In addition to the avian LAL model presented here, recent studies in mice, fetal sheep, and frogs ...

Congenital heart defects (CHDs) are structural disorders that occur due to abnormal embryonic development1. In addition to genetic conditions, the pathogenesis is influenced by altered mechanical loading2,3. Hypoplastic left heart syndrome (HLHS), a congenital heart disease, results in an underdeveloped ventricle/aorta at birth4 with a high rate of mortality5,6. Despite the recent advances in its clinical management, the vascular growth and development dynamics of HLHS are still unclear7. In normal embryonic development, the left ventricle (LV) endocardium and myocardium originate from cardiac progenitors as the early embryonic heart tube formation progresses. The gradual presence of myocardial trabeculation, thickening layers, and cardiomyocyte proliferation is reported2. For HLHS, altered trabecular remodeling and left ventricular flattening are observed, further contributing to myocardial hypoplasia due to abnormal cardiomyocyte migration2,8,9,10
Among the widely used model organisms to study heart development and understand hemodynamic conditions11, the avian embryo is preferred due to its four-chambered mature heart and its ease of culture11,12,13,14. On the other hand, advanced imaging access of zebrafish embryos and transgenic/knockout mice provide distinct advantages11,12. Various mechanical interventions have been tested for the avian embryo that alter the intramural pressure and wall shear stress in developing cardiovascular components. These models include left vitelline ligation, conotruncal banding15, and left atrial ligation (LAL)11,12,16. The resulting phenotype due to the altered mechanical loading can be observed approximately 24-48 h after the surgical intervention in studies focusing on early prognosis11,13. The LAL intervention is a popular technique to narrow the functional volume of the left atrium (LA) by placing a suture loop around the atrioventricular opening. Likewise, microsurgical interventions have also been performed that target right atrial ligation (RAL)17,18. Similarly, some researchers target the left atrial appendage (LAA) using micro clips to reduce the volume of the LA19,20. In some studies, a surgical nylon thread is applied to the atrioventricular node19,21. One of the interventions used is LAL, which can mimic HLHS but is also the most difficult model to perform, with very small sample yields due to the extremely fine microsurgical operations required. In our laboratory, LAL is performed in ovo between Hamburger-Hamilton (HH) stages 20 and 21, before the common atrium is fully septate6,14,22,23. A surgical suture is placed around the LA, which alters the intracardiac blood flow streams. In LAL models of HLHS, increased ventricle wall stiffness, altered myofiber angles, and decreased LV cavity size are observed14,24.
In this video article, a detailed protocol and approach for in ovo LAL is provided. Briefly, the fertilized avian embryos were incubated for microsurgery, the eggshell was cracked open, and the outer and inner membranes were cleared. The embryo was then slowly rotated so that the LA was accessible. A 10-0 nylon surgical suture was tied to the atrial bud, and the embryo was returned to its original orientation, completing the LAL procedure25. LAL and normal ventricles are compared for tissue compaction and ventricle volume via optical coherence tomography and basic histology.
A successfully executed LAL model pipeline, as described here, will contribute to basic studies focusing on the embryonic development of cardiovascular components. This model can also be used together with genetic manipulations and advanced imaging modalities. Likewise, the acute LAL model is a stable source of diseased vascular cells for tissue culture experiments.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>10-0 nylon surgical suture</td>
			<td>Ethicon</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Elastica van Gieson staining kit</td>
			<td>Sigma-Aldrich</td>
			<td>115974</td>
			<td>For staining connective tissues in histological sections</td>
		</tr>
		<tr>
			<td>Ethanol absolute</td>
			<td>Interlab</td>
			<td>64-17-5</td>
			<td>For the sterilization step, 70% ethanol was obtained by diluting absolute ethanol with distilled water.</td>
		</tr>
		<tr>
			<td>Incubator</td>
			<td>KUHL, Flemington, New Jersey-U.S.A</td>
			<td>AZYSS600-110</td>
			<td></td>
		</tr>
		<tr>
			<td>Kimwipes</td>
			<td>Interlab</td>
			<td>080.65.002</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscissors</td>
			<td>World Precision Instruments (WPI), Sarasota FL</td>
			<td>555640S</td>
			<td>Vannas STR 82 mm</td>
		</tr>
		<tr>
			<td>Parafilm M</td>
			<td>Sigma-Aldrich</td>
			<td>P7793-1EA</td>
			<td>Sealing stage for egg reincubation</td>
		</tr>
		<tr>
			<td>Paraplast Bulk</td>
			<td>Leica Biosystems&#160;</td>
			<td>39602012</td>
			<td>Tissue embedding medium</td>
		</tr>
		<tr>
			<td>Stereo Microscope</td>
			<td>Zeiss Stemi 508</td>
			<td>&#160;Stemi 508</td>
			<td>Used at station 1</td>
		</tr>
		<tr>
			<td>Stereo Microscope</td>
			<td>Zeiss Stemi 2000-C</td>
			<td>Stemi 2000-C</td>
			<td>Used at station 2</td>
		</tr>
		<tr>
			<td>Tweezer (Dumont 4 INOX #F4)</td>
			<td>Adumont &#38; Fils, Switzerland</td>
			<td></td>
			<td>Used to return the embryo</td>
		</tr>
		<tr>
			<td>Tweezer (Super Fine Dumont #5SF)&#160;</td>
			<td>World Precision Instruments (WPI), Sarasota FL</td>
			<td>501985</td>
			<td>Used to remove the membranes on the embryo</td>
		</tr>
	</tbody>
</table>

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.

Advanced time-resolved imaging techniques can be employed to observe the structural and morphological changes due to LAL intervention10. Furthermore, LAL samples are also amenable to molecular and biological methods19,28. In Table 1, sample studies that employed LAL model results are provided. In this context, LAL intervention was performed in chick embryos that reached HH20-21. Both control (healthy) and LAL hearts were r...

Watch this Scientific Journal Video about Effect of Left Atrial Ligation on Avian Embryonic Hearts and HLHS Implications at JoVE.com

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.

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 ...

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Developmental Biology

1.0K Views.  Koc University. 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. 

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

Protocols for Obtaining Zygotic and Somatic Embryos for Studying the Regulation of Early Embryo Development in the Model Legume Medicago truncatula

Early embryogenesis starting from a single cell zygote goes through rapid cell division and morphogenesis, and is morphologically characterized by pre-globular, globular, heart, torpedo and cotyledon stages. This progressive development is under the tight regulation of a complex molecular network. Harvesting sufficient early embryos at a similar stage of development is essential for investigating the cellular and molecular regulation of early embryogenesis. This is not straightforward since early embryogenesis undergoes rapid morphogenesis in a short while e.g. 8 days for Medicago truncatula to reach the early cotyledon stage. Here, we address the issue by two approaches. The first one establishes a linkage between embryo development and pod morphology in helping indicate the stage of the zygotic embryo. This is particularly based on the number of pod spirals and development of the spines. An alternative way to complement the in vivo studies is via culturing leaf explants to produce somatic embryos. The medium includes an unusual hormone combination - an auxin (1-naphthaleneacetic acid), a cytokinin (6-benzylaminopurine), abscisic acid and gibberellic acid. The different stages can be discerned growing out of the callus without dissection.

Protocols for Obtaining Zygotic and Somatic Embryos for Studying the Regulation of Early Embryo Development in the Model Legume Medicago truncatula

The goal is to illustrate that the model legume Medicago truncatula can be readily utilized to investigate the regulation of early plant embryogenesis to complement the non-legume Arabidopsis model. Pod morphology is linked to zygotic embryogenesis stages and a protocol to collect embryos using tissue culture is also provided.

Early embryogenesis starting from a single cell zygote goes through rapid cell division and morphogenesis, and is morphologically characterized by ...

Microbead Implantation in the Zebrafish Embryo

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic conservation, as well as similarities in cellular, molecular, and physiological processes, with other vertebrates including humans. During early ontogeny, zebrafish embryos are optically transparent, allowing researchers to visualize the dynamics of organogenesis using a simple stereomicroscope. Microbead implantation is a method that enables tissue manipulation through the alteration of factors in local environments. This allows researchers to assay the effects of any number of signaling molecules of interest, such as secreted peptides, at specific spatial and temporal points within the developing embryo. Here, we detail a protocol for how to manipulate and implant beads during early zebrafish development.

The zebrafish is an excellent model system for genetic and developmental studies. Bead implantation is a valuable tissue manipulation technique that can be used to interrogate developmental mechanisms by introducing alterations in local cellular environments. This protocol describes how to perform microbead implantation in the zebrafish embryo.

The zebrafish has emerged as a valuable genetic model system for the study of developmental biology and disease. Zebrafish share a high degree of genomic ...

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Assessment of Maternal Vascular Remodeling During Pregnancy in the Mouse Uterus

The placenta mediates the exchange of factors such as gases and nutrients between mother and fetus and has specific demands for supply of blood from the maternal circulation. The maternal uterine vasculature needs to adapt to this temporary demand and the success of this arterial remodeling process has implications for fetal growth. Cells of the maternal immune system, especially natural killer (NK) cells, play a critical role in this process. Here we describe a method to assess the degree of remodeling of maternal spiral arteries during mouse pregnancy. Hematoxylin and eosin-stained tissue sections are scanned and the size of the vessels analysed. As a complementary validation method, we also present a qualitative assessment for the success of the remodeling process by immunohistochemical detection of smooth muscle actin (SMA), which normally disappears from within the arterial vascular media at mid-gestation. Together, these methods enable determination of an important parameter of the pregnancy phenotype. These results can be combined with other endpoints of mouse pregnancy to provide insight into the mechanisms underlying pregnancy-related complications.

This protocol describes a technique to assess changes in the maternal vasculature during pregnancy in mice. Using stereological methods, remodeling of the decidual spiral arteries is assessed quantitatively and the results confirmed qualitatively using immunohistochemistry.

The placenta mediates the exchange of factors such as gases and nutrients between mother and fetus and has specific demands for supply of blood from the ...

Two-step Approach to Explore Early- and Late-stages of Organ Formation in the Avian Model: The Thymus and Parathyroid Glands Organogenesis Paradigm

The avian embryo, as an experimental model, has been of utmost importance for seminal discoveries in developmental biology. Among several approaches, the formation of quail-chicken chimeras and the use of the chorioallantoic membrane (CAM) to sustain the development of ectopic tissues date back to the last century. Nowadays, the combination of these classical techniques with recent in vitro methodologies offers novel prospects to further explore organ formation.
Here we describe a two-step approach to study early- and late-stages of organogenesis. Briefly, the embryonic region containing the presumptive territory of the organ is isolated from quail embryos and grown in vitro in an organotypic system (up to 48 h). Cultured tissues are subsequently grafted onto the CAM of a chicken embryo. After 10 days of in ovo development, fully formed organs are obtained from grafted tissues. This method also allows the modulation of signaling pathways by the regular administration of pharmacological agents and tissue genetic manipulation throughout in vitro and in ovo developmental steps. Additionally, developing tissues can be collected at any time-window to analyze their gene-expression profile (using quantitative PCR (qPCR), microarrays, etc.) and morphology (assessed with conventional histology and immunochemistry).
The described experimental procedure can be used as a tool to follow organ formation outside the avian embryo, from the early stages of organogenesis to fully formed and functional organs.

This article provides an updated approach to the classical quail-chicken chimera system to study organ formation, by combining novel in vitro and in ovo experimental procedures.

The avian embryo, as an experimental model, has been of utmost importance for seminal discoveries in developmental biology. Among several approaches, the ...

Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model

Assisted reproductive techniques (ARTs), such as in vitro embryo culture or embryo cryopreservation, affect natural development patterns with perinatal and postnatal consequences. To ensure the innocuousness of ART applications, studies in animal models are necessary. In addition, as a last step, embryo development studies require evaluation of their capacity to develop full-term healthy offspring. Here, embryo transfer to the uterus is indispensable to perform any ARTs-related experiment.
The rabbit has been used as a model organism to study mammalian reproduction for over a century. In addition to its phylogenetic proximity to the human species and its small size and low maintenance cost, it has important reproductive characteristics such as induced ovulation, a chronology of early embryonic development similar to humans and a short gestation that allow us to study the consequences of ART application easily. Moreover, ARTs (such as intracytoplasmic sperm injection, embryo culture, or cryopreservation) are applied with suitable efficiency in this species.
Using the laparoscopic embryo transfer technique and the cryopreservation protocol presented in this article, we describe 1) how to transfer embryos through an easy, minimally invasive technique and 2) an effective protocol for long-term storage of rabbit embryos to provide time-flexible logistical capacities and the ability to transport the sample. The outcomes obtained after transferring rabbit embryos at different developmental stages indicate that morula is the ideal stage for rabbit embryo recovery and transfer. Thus, an oviductal embryo transfer is required, justifying the surgical procedure. Furthermore, rabbit morulae are successfully vitrified and laparoscopically transferred, proving the effectiveness of the described techniques.

Assisted reproductive techniques (ARTs) are in continuous evaluation to improve outcomes and reduce the associated risks. This manuscript describes a minimally invasive embryo transfer procedure with an efficient cryopreservation protocol that allows the use of rabbits as an ideal animal model of human reproduction.

Assisted reproductive techniques (ARTs), such as in vitro embryo culture or embryo cryopreservation, affect natural development patterns with perinatal ...

Microinjection-based System for In Vivo Implantation of Embryonic Cardiomyocytes in the Avian Embryo

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a general challenge in developmental biology research. In the embryonic heart, this can be particularly difficult as regional differences in the expression of transcriptional regulators, paracrine/juxtacrine signaling cues, and hemodynamic force are all known to influence cardiomyocyte maturation. A simplified method to alter a developing cardiomyocyte's molecular and biomechanical microenvironment would, therefore, serve as a powerful technique to examine how local conditions influence cell fate and function. To address this, we have optimized a method to physically transplant juvenile cardiomyocytes into ectopic locations in the heart or the surrounding embryonic tissue. This allows us to examine how microenvironmental conditions influence cardiomyocyte fate transitions at single cell resolution within the intact embryo. Here, we describe a protocol in which embryonic myocytes can be isolated from a variety of cardiac sub-domains, dissociated, fluorescently labeled, and microinjected into host embryos with high precision. Cells can then be directly analyzed in situ using a variety of imaging and histological techniques. This protocol is a powerful alternative to traditional grafting experiments that can be prohibitively difficult in a moving tissue such as the heart. The general outline of this method can also be adapted to a variety of donor tissues and host environments, and its ease of use, low cost, and speed make it a potentially useful application for a variety of developmental studies.

In this method, embryonic cardiac tissues are surgically microdissected, dissociated, fluorescently labeled, and implanted into host embryonic tissues. This provides a platform for studying the individual or tissue level developmental organization under ectopic hemodynamic conditions, and/or altered paracrine/juxtacrine environments.

Interpreting the relative impact of cell autonomous patterning versus extrinsic microenvironmental influence on cell lineage determination represents a ...

Genotyping of Sea Anemone during Early Development

Described here is a PCR-based protocol to genotype the gastrula stage embryo of the anthozoan cnidarian Nematostella vectensis without sacrificing the life of the animal. Following in vitro fertilization and de-jellying, zygotes are allowed to develop for 24 h at room temperature to reach the early- to mid-gastrula stage. The gastrula embryos are then placed on an agarose gel bed in a Petri dish containing seawater. Under the dissecting microscope, a tungsten needle is used to surgically separate an aboral tissue fragment from each embryo. Post-surgery embryos are then allowed to heal and continue development. Genomic DNA is extracted from the isolated tissue fragment and used as a template for locus-specific PCR. The genotype can be determined based on the size of PCR products or presence/absence of allele-specific PCR products. Post-surgery embryos are then sorted according to the genotype. The duration of the entire genotyping process depends on the number of embryos to be screened, but it minimally requires 4&#8211;5 h. This method can be used to identify knockout mutants from a genetically heterogeneous population of embryos and enables analyses of phenotypes during development.

The goal of this protocol is to genotype the sea anemone Nematostella vectensis during gastrulation without sacrificing the embryo.

Described here is a PCR-based protocol to genotype the gastrula stage embryo of the anthozoan cnidarian Nematostella vectensis without sacrificing the ...

Vascular Casting of Adult and Early Postnatal Mouse Lungs for Micro-CT Imaging

Blood vessels form intricate networks in 3-dimensional space. Consequently, it is difficult to visually appreciate how vascular networks interact and behave by observing the surface of a tissue. This method provides a means to visualize the complex 3-dimensional vascular architecture of the lung.
To accomplish this, a catheter is inserted into the pulmonary artery and the vasculature is simultaneously flushed of blood and chemically dilated to limit resistance. Lungs are then inflated through the trachea at a standard pressure and the polymer compound is infused into the vascular bed at a standard flow rate. Once the entire arterial network is filled and allowed to cure, the lung vasculature may be visualized directly or imaged on a micro-CT (&#181;CT) scanner.
When performed successfully, one can appreciate the pulmonary arterial network in mice ranging from early postnatal ages to adults. Additionally, while demonstrated in the pulmonary arterial bed, this method can be applied to any vascular bed with optimized catheter placement and endpoints.

The aim of this technique is ex vivo visualization of pulmonary arterial networks of early postnatal and adult mice through lung inflation and injection of a radio-opaque polymer-based compound via the pulmonary artery. Potential applications for casted tissues are also discussed.

Blood vessels form intricate networks in 3-dimensional space. Consequently, it is difficult to visually appreciate how vascular networks interact and ...

A Protocol for Immunohistochemistry and RNA In-situ Distribution within Early Drosophila Embryo

Calcium induced calcium release signaling (CICR) plays a critical role in many biological processes. Every cellular activity from cell proliferation and apoptosis, development and ageing, to neuronal synaptic plasticity and regeneration have been associated with Ryanodine receptors (RyRs). Despite the importance of calcium signaling, the exact mechanism of its function in early development is unclear. As an organism with a short gestational period, the embryos of Drosophila melanogaster are prime study subjects for investigating the distribution and localization of CICR associated proteins and their regulators during development. However, because of their lipid-rich embryos and chitin-rich chorion, their utility is limited by the difficulty of mounting embryos on glass surfaces. In this work, we introduce a practical protocol that significantly enhances the attachment of Drosophila embryo onto slides and detail methods for successful histochemistry, immunohistochemistry, and in-situ hybridization. The chrome alum gelatin slide-coating method and embryo pre-embedding method dramatically increases the yield in studying Drosophila embryo protein and RNA expression. To demonstrate this approach, we studied DmFKBP12/Calstabin, a well-known regulator of RyR during early embryonic development of Drosophila melanogaster. We identified DmFKBP12 in as early as the syncytial blastoderm stage and report the dynamic expression pattern of DmFKBP12 during development: initially as an evenly distributed protein in the syncytial blastoderm, then preliminarily localizing to the basement layer of the cortex during cellular blastoderm, before distributing in the primitive neuronal and digestion architecture during the three-gem layer phase in early gastrulation. This distribution may explain the critical role RyR plays in the vital organ systems that originate in from these layers: the suboesophageal and supraesophageal ganglion, ventral nervous system, and musculoskeletal system.

A Protocol for Immunohistochemistry and RNA In-situ Distribution within Early Drosophila Embryo

Here, we describe a protocol for detection and localization of Drosophila embryo protein and RNA from collection to pre-embedding and embedding, immunostaining, and mRNA in situ hybridization.

Calcium induced calcium release signaling (CICR) plays a critical role in many biological processes. Every cellular activity from cell proliferation and ...

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

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Chapters in this video

Protocols for Obtaining Zygotic and Somatic Embryos for Studying the Regulation of Early Embryo Development in the Model Legume Medicago truncatula

Microbead Implantation in the Zebrafish Embryo

Imaging Subcellular Structures in the Living Zebrafish Embryo

Assessment of Maternal Vascular Remodeling During Pregnancy in the Mouse Uterus

Two-step Approach to Explore Early- and Late-stages of Organ Formation in the Avian Model: The Thymus and Parathyroid Glands Organogenesis Paradigm

Minimally Invasive Embryo Transfer and Embryo Vitrification at the Optimal Embryo Stage in Rabbit Model

Microinjection-based System for In Vivo Implantation of Embryonic Cardiomyocytes in the Avian Embryo

Genotyping of Sea Anemone during Early Development

Vascular Casting of Adult and Early Postnatal Mouse Lungs for Micro-CT Imaging

A Protocol for Immunohistochemistry and RNA In-situ Distribution within Early Drosophila Embryo