This work was supported by National Natural Science Foundation of China (Grant No. 91643203, 91543208, 81502835).

All procedures described were approved by the Institutional Animal Care and Use Committee (IACUC) of Qingdao University. In our lab, the eggs were incubated in two incubators. Eggs were held upright in the incubator and randomly placed on the shelves. The incubation conditions for the eggs were as follows: incubation temperature started at 37.9 &#176;C, and gradually decreased to 37.1 &#176;C as incubation proceeded; the humidity started at 50% and gradually increased to 70%.
1. Exposure methods
NOTE: Exposure of environmental contaminants to chicken embryos may be achieved in several ways. In this section, three routinely used methods are described in detail.
<ol>
	<li>Air cell injection (Figure 1) 
	NOTE: This is the classical exposure method to chicken embryos4,5,6, suitable for a wide range of materials, and may be performed at a very wide time window, from the beginning of development (embryonic day zero, ED0) all the way to the day prior to hatch (ED20). Sunflower oil is used as the vehicle. Previous studies have shown that no significant changes in mortality, hatchability, or body weight have been observed between untreated embryos and embryos injected with sunflower oil7.
	<ol>
		<li>Prepare the following necessary reagents/tools: 75% ethanol, tissue paper, metal probe (can substitute with any sharp metal needle/stick/awl), melted paraffin, brush, povidone iodide solution, pipette, pipette tips, candling lamp, dosing mixture. Prepare the dosing mixture with sunflower oil (recommended)4. To use other diluents, perform vehicle control (versus untreated embryos).</li>
		<li>Clean the surface of the eggs with povidone iodide solution (commercially available povidone iodide solution 1:5 diluted with deionized water), and dip-dry the egg shell with tissue paper without scrubbing. Scrubbing will break the protective layer coating the outside of the shell.</li>
		<li>Candle the eggs in a dark room and mark the air cell with pencil. Exclude eggs with cracks on the shell. Exclude eggs with air cells on the side instead of blunt tip, as those are highly unlikely to hatch normally.</li>
		<li>Sanitize the air cell area with 75% ethanol, and then drill a small hole at the center of air cell area with the metal probe. Do not stick the probe deep into the air cell or the inner membrane may be damaged, instead, only break the shell with the tip of the probe. If the hole is not large enough to fit in a fine pipette tip, break the shell again at the vicinity of existing hole, until the hole is large enough to allow insertion of 10 &#181;L pipette tip.</li>
		<li>Vortex the dosing mixture vigorously, and immediately draw solution to pipette tip. The recommended injection volume is 1 &#181;L per 10 gram of egg (e.g., 5 &#181;L injection volume for a 50-gram egg) as larger injection volumes may create hypoxic or anoxic conditions for the developing embryo. Calculate the concentration of the dosing solution for desired mg/egg kg dose.</li>
		<li>Insert the pipette tip into the hole, with the tip touching the inner membrane. Slowly eject the dosing mixture, hold for at least ten seconds (allow the viscous oil to be fully dispensed), and then remove the tip.</li>
		<li>Seal the hole with a brush and a drop of melted paraffin. Be careful not to drip melted paraffin onto the inner membrane.</li>
		<li>Once sealed, place the eggs in the incubator until they reach the desired embryonic stage. On already developing embryos, perform the whole process as soon as possible to prevent potential embryo loss due to low environmental temperatures.</li>
	</ol>
	</li>
	<li>Microinjection (Figure 2) 
	NOTE: This is a more direct exposure method, resulting in definitive exposure to the substance-of-interest, and especially suitable for compounds with a short duration of action (e.g., lentivirus), since the classical air cell injection requires time for the compounds to penetrate the inner membrane. This method may also be tried if satisfying results could not be achieved by air cell injection. This method is best suitable for early embryos (up to ED2), but also can be performed on older embryos (with higher risk of embryo loss).
	<ol>
		<li>Prepare the following necessary reagents/tools: 75% ethanol, povidone iodide solution, micro-injector (5 &#181;L), metal probe (can substitute with any sharp metal needle/stick/awl should work), fine forceps, tape. Prepare the dosing mixture with sterile saline, which also serves as an injection control without affecting hatchability significantly. Ensure the sterility of the saline, as a contaminated injection will dramatically increase mortality.</li>
		<li>Clean the eggs as described in 1.1.2.</li>
		<li>Candle the eggs as described in 1.1.3.</li>
		<li>Sanitize the air cell area with 75% ethanol, and then drill a small hole at the center of air cell area with the metal probe. Do not stick the probe deep into the air cell or the inner membrane may be damaged, instead, only break the shell with the tip of the probe. Then use the fine forceps to carefully enlarge the hole until the diameter is approximately 2 mm, allowing visual confirmation of the inner membrane.</li>
		<li>Load the solution into microinjector (maximum injection volume: 0.5 &#181;L/10 g egg. (e.g., 2.5 &#181;L for a 50 g egg) and carefully insert the needle through the hole into the inner membrane for approximately 2-3 mm. Gently dispense the solution and remove the needle. Keep the needle as perpendicular to the membrane as possible.</li>
		<li>Seal the hole with a small piece of tape. Completely cover the hole to prevent embryo dehydration and death during subsequent incubation. Nevertheless, avoid pieces of tape that are too large to prevent hypoxia.</li>
		<li>Once sealed, place the eggs in the incubator until they reach the desired embryonic stage. On already developing embryos, perform the whole process as soon as possible to prevent potential embryo loss due to low environmental temperature.</li>
	</ol>
	</li>
	<li>Air cell inhalation (Figure 3) 
	NOTE: This is a novel inhalation method taking advantage of the air cell, from which the late-stage chicken embryo will start to breathe air. It is suitable for gas or aerosol exposures and may achieve very early-in-life inhalation exposures, and fill the lungs with the target gas/aerosol when they open for the first time in life.
	<ol>
		<li>Prepare the following necessary reagents/tools: Sampling bag (PVF bag, for storage of gas/aerosol sample before exposure), catheter needle, syringe, metal probe (can substitute with any sharp metal needle/stick/awl should work), tape, fume hood.</li>
		<li>Clean the eggs as described in 1.1.2 and candle them as described in 1.1.3 (it is not necessary to mark air cell prior to incubation), and then incubate eggs without treatment until ED17.</li>
		<li>Candle the eggs at ED17 to mark the air cell area.</li>
		<li>At ED18, take an egg out from the incubator, sanitize the air cell area with 75% ethanol, and then carefully drill two small holes on two sides of the air cell. One is for injection of gas/aerosol, the other is for expelling of air. Carefully control the size of the holes so that the size of the injection hole is just enough for the catheter needle to be inserted, while the diameter of the expelling hole is a bit larger (approximately 1 mm).</li>
		<li>Gently inject 10 mL of target gas/aerosol from the injection hole with a syringe attached to the catheter needle. Inject air for an inhalation control group, which should have no significant differences to a negative control group8. Apply pressure against the catheter needle (with appropriate amount of pressure the elastic needle can be pushed against the shell) to minimize the leakage from injection hole. Seal both holes immediately with tape afterwards, and return the egg to incubator. 
		&#8203;NOTE: This procedure should be performed in a fume hood to prevent inhalation of gas/aerosol by the operator.</li>
		<li>Repeat the described procedure after one hour to further ensure the whole air cell is filled up with target gas/aerosol (optional).</li>
		<li>Repeat the described procedure at ED19 again (optional). Repeating the exposure helps to ensure consistent exposure until hatch. Record the hatch time for an approximate estimation of exposure duration.</li>
		<li>Once desired exposures have been performed and sealed, place the eggs in the incubator for hatching. Minimize the time the egg spends outside of the incubator to prevent death from low environmental temperature.</li>
	</ol>
	</li>
</ol>
2. Endpoint assessment methods
NOTE: Following exposure of contaminant-of-interest to the developing embryo, several toxicity parameters can be evaluated, including cardiotoxicity. In this section, two frequently used specific methods are described in detail.
<ol>
	<li>Electrocardiography (Figure 4) 
	NOTE: It is impossible to do non-invasive electrocardiography in hatchling chickens due to the presence of feathers. Thus, subcutaneous implantation of electrodes is essential, requiring anesthesia. The dose used in the lab is 33 mg/kg pentobarbital via intraperitoneal injection (some chickens may require up to 50% dose increase). This method will result in stable electrocardiography in over 90% of animals, allowing for analyzation of heart rate.
	<ol>
		<li>Prepare the following necessary reagents/tools: 1% (10 mg/mL) pentobarbital solution in saline, syringe, electrical balance, heater (if necessary), an electrocardiography instrument attached (e.g., BL-420E+) with two-channel metal needle electrodes.</li>
		<li>Weigh the chickens with a balance and calculate the necessary volume of pentobarbital solution and inject the chickens. For a chicken that weighs 30 g, 0.1 mL of pentobarbital solution is needed. Make sure that the injection is done on the lateral side of the abdomen, as the yolk is located at the middle and injection there may not be effective.</li>
		<li>Wait until the injected chickens are anesthetized (hold the chicken in hand, if the neck has no tension and the head can be swung freely, the anesthetization is sufficient). Place chickens on the operation table (heaters are necessary if room temperature is below 20 &#176;C).</li>
		<li>Insert two needle electrodes from two sides of the abdomen, subcutaneously. Make sure that the needles do not enter the abdominal cavity by lifting the skin a little bit and inserting needle from there. Once inserted, carefully push the needle forward until it reaches the side of chest cavity. Make sure that the needle does not go deep into the body or stick out from the skin.</li>
		<li>Make the measurements with the electrocardiography instrument. Other similar instruments capable of electrocardiography can be used.</li>
		<li>If the chickens are to be sacrificed, perform euthanasia after the electrocardiography since they are already under anesthesia. If chickens are to survive, place them back into their cages and warm until waking up. Returning them to the incubator is another option.</li>
	</ol>
	</li>
	<li>Histomorphometry (Figure 5) 
	NOTE: A specific method is developed to assess the right ventricular wall thickness in transverse sections of the heart. Morphological assessment of right ventricle dimension via echocardiography is not 100% accurate due to the irregular shape of right ventricle, and this method may serve as a good supplement in the morphological assessments for the right ventricle.
	<ol>
		<li>Prepare the following necessary reagents/tools: 4% phosphate buffered formaldehyde, sharp blade, phosphate buffered saline, paper towel, electrical balance, small scissors, general histological processing agents (graded ethanol, xylene, paraffin).</li>
		<li>Once animals are sacrificed, use water to wet the feathers. This is to minimize the potential contamination due to flying feathers while opening up the chest.</li>
		<li>Open the chest cavity carefully without damaging the heart. Use small scissors to cut the vasculature and gently remove the heart from the chest cavity. Leave a small piece (approximately 1-2 mm) of vasculature attached to the heart as this may be convenient for subsequent handling of the heart without damaging the heart.</li>
		<li>Once removed, rinse the heart in cold phosphate buffered saline to remove blood and relax the muscle. Then dip dry the heart on paper towel before weighing for accurate weight reading. Put the heart into 10x volume fixative (4% phosphate buffered formaldehyde) for 24 hours at room temperature. Fixed tissues may be subsequently processed to paraffin blocks, or be stored at 4 &#176;C for years (not recommended if immunohistochemistry is planned).</li>
		<li>Before embedding, cut the tissues at approximately 60% length of the heart counting from the apex (Figure 5A), for easier subsequent processing. A microtome blade is recommended for a quick and vertical clean cut. For chickens older than one day, make another cut at approximately 25-30% length from the apex for easier paraffin penetration and to allow the tissue to fit in tissue cassettes.</li>
		<li>Process the tissues with the following conditions (adjust as needed): 70% ethanol for 1 h, 80% ethanol for 1 h, 95% ethanol for 1 h x2, 100% ethanol for 30 min x2, xylene for 5 min x2, paraffin (melting point 52-54 &#176;C) for 1.5 h, paraffin (melting point 62-64 &#176;C) for 1 h, and then embed the tissues in a 3:1 mixture of paraffin (melting point 62-64 &#176;C) and paraffin (melting point 52-54 &#176;C).</li>
		<li>Section the tissue at 6 &#181;m thickness. Carefully maintain identical relative positions of the cross-sections by confirming the presence and size of an anatomical landmark (septomarginal trabecula) in the right ventricle. Confirm a landmark with moderate length on each section (Figure 5B, arrow).</li>
		<li>Make two electronic rulers with Logo programmer: Ruler 1 is a straight line with 7 radius measure lines attached to the middle point, with 22.5&#176; in between two adjacent measure lines. Ruler 2 is just two perpendicular lines in a T shape (Figure 5B).</li>
		<li>Measure with two software programs: Adobe Photoshop and ImageJ.
		<ol>
			<li>In Photoshop, resize ruler 1 (NO reshaping) to place the two ends of the ruler on the two ends of free right ventricular wall, so that the seven measure lines on ruler 1 will each meet the inner right ventricular wall. Then use ruler 2 to make perpendicular measurements from the inner to external ventricular wall (Figure 5B).</li>
			<li>Use ImageJ to make the seven measurements for each heart.</li>
		</ol>
		</li>
		<li>Depending on specific needs, analyze the seven measurements or average for one representative right ventricular wall thickness. Normalize to whole heart weight for specific ventricular wall thickness changes.</li>
	</ol>
	</li>
</ol>

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The authors declare no conflict of interest.

The chicken embryo has been a classical model in developmental studies for 200 years1. Our methods presented in this manuscript have been used in the assessment of several environmental contaminants, including perfluorooctanoic acid, particulate matter, and diesel exhaust with success5, 7,8,9,10,11<sup...

The chicken embryo is a classic developmental model, which has been used for over two hundred years1. The chicken embryo model has various advantages compared to traditional models. First of all, as early as over 70 years ago, the normal development of the chicken embryo had been illustrated very clearly in the Hamburger-Hamilton staging guide2, in which a total of 46 stages during chicken embryo development were defined with precise time and morphological characteristics, facilitating detections of abnormal development. Additionally, the chicken embryo model has other features such as being relatively low-cost and redundant in quantity, relatively accurate exposure-dose controls, an independent, closed system within the shell and easy manipulation of the developing embryo, all of which guarantees its potential to be used as a powerful toxicological assessment model. 
In cardiotoxicity, the chicken embryo features a four chambered heart, similar to mammalian hearts but with thicker walls, allowing easier morphological assessments. Additionally, the chicken embryo allows for developmental inhalation exposure, which is not possible in mammalian models: during the later stage of development, the chicken embryo will transition from internal respiration to external respiration (getting oxygen via the lung); the latter requires that the embryo penetrates the air cell membrane with the beak, and starts to breathe air3, making the air cell a mini-inhalation chamber. Utilizing this phenomenon, the toxicological effects of gas contaminants on the heart (and other organs) may be assessed without the need of dedicated inhalation chamber instruments. 
In this manuscript, several exposure/endpoint assessment methods are described, all of which serve to make the chicken embryo a powerful tool in the assessment of development cardiotoxicity following exposure to environmental contaminants.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>4% phosphate buffered formaldehydefixative</td>
			<td>Biosharp, Hefei, China</td>
			<td>REF: BL539A</td>
			<td></td>
		</tr>
		<tr>
			<td>75% ethanol</td>
			<td>Guoyao,Shanghai,China</td>
			<td>CAS:64-17-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Biosignaling monitor BL-420E+</td>
			<td>Taimeng, Chengdu, China</td>
			<td>BL-420E+</td>
			<td></td>
		</tr>
		<tr>
			<td>Candling lamp</td>
			<td>Zhenwei, Dezhou, China</td>
			<td>WZ-001</td>
			<td></td>
		</tr>
		<tr>
			<td>Disposable syringe</td>
			<td>Zhiyu, Jiangsu, China</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Egg incubator</td>
			<td>Keyu,Dezhou, China</td>
			<td>KFX</td>
			<td></td>
		</tr>
		<tr>
			<td>Electrical balance</td>
			<td>OHAUS, Shanghai, China</td>
			<td>AR 224CN</td>
			<td></td>
		</tr>
		<tr>
			<td>Electro-thermal incubator</td>
			<td>Shenxian, Shanghai, China</td>
			<td>DHP-9022</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol absolute</td>
			<td>Guoyao,Shanghai,China</td>
			<td>CAS:64-17-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Fertile chicken egg</td>
			<td>Jianuo, Jining, China</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hematoxylin and Eosin Staining Kit</td>
			<td>Beyotime, Bejing, China</td>
			<td>C0105</td>
			<td></td>
		</tr>
		<tr>
			<td>Histology paraffin</td>
			<td>Aladdin, Shanghai, China</td>
			<td>P100928-500g</td>
			<td>Melt point 52~54&#176;C</td>
		</tr>
		<tr>
			<td>Histology paraffin</td>
			<td>Aladdin, Shanghai, China</td>
			<td>P100936-500g</td>
			<td>Melt point 62~64&#176;C</td>
		</tr>
		<tr>
			<td>IV catheter</td>
			<td>KDL, Zhejiang, China</td>
			<td></td>
			<td>The catheters have to be soft, plastic ones.</td>
		</tr>
		<tr>
			<td>Lentivirus</td>
			<td>Genechem, Shanghai, China</td>
			<td></td>
			<td>The lentivirus were individually designed/synthesized by Genechem.</td>
		</tr>
		<tr>
			<td>Masson&#39;s trichrome staining kit</td>
			<td>Solarbio, Beijing, China</td>
			<td>G1340</td>
			<td></td>
		</tr>
		<tr>
			<td>Metal probe</td>
			<td>Jinuotai, Beijing, China</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microinjector (5 uL)</td>
			<td>Anting,Shanghai, China</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>CAIKON, Shanghai, China</td>
			<td>XSP-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Microtome</td>
			<td>Leica, Germany</td>
			<td>HistoCore BIOCUT</td>
			<td></td>
		</tr>
		<tr>
			<td>Microtome blade</td>
			<td>Leica,Germany</td>
			<td>Leica 819</td>
			<td></td>
		</tr>
		<tr>
			<td>Pentobarbitual sodium</td>
			<td>Yitai Technology Co. Ltd.,&#160; Wuhan, China</td>
			<td>CAS: 57-33-0</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipetter(10ul)</td>
			<td>Sartorius, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Povidone iodide</td>
			<td>Longyuquan, Taian, China</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Scissor</td>
			<td>Anqisheng,Suzhou, China</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile saline</td>
			<td>Kelun,Chengdu, China</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sunflower oil</td>
			<td>Mighty Jiage, Jiangsu, China</td>
			<td></td>
			<td>Any commerical sunflower oil for human consumption should work</td>
		</tr>
		<tr>
			<td>Tape</td>
			<td>M&#38;G, Shanghai, China</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tedlar PVF Bag (5L)</td>
			<td>Delin, Dalian, China</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Vortex mixer</td>
			<td>SCILOGEX, Rocky Hill, CT, US</td>
			<td>MX-F</td>
			<td></td>
		</tr>
		<tr>
			<td>Xylene</td>
			<td>Guoyao,Shanghai,China</td>
			<td>CAS:1330-20-7</td>
			<td></td>
		</tr>
	</tbody>
</table>

using chicken embryo as a powerful tool in assessment of developmental cardiotoxicities

Chicken embryos are a classical model in developmental studies. During the development of chicken embryos, the time window of heart development is well-defined, and it is relatively easy to achieve precise and timely exposure via multiple methods. Moreover, the process of heart development in chicken embryos is similar to mammals, also resulting in a four-chambered heart, making it a valuable alternative model in the assessment of developmental cardiotoxicities. In our lab, the chicken embryo model is routinely used in the assessment of developmental cardiotoxicities following exposure to various environmental pollutants, including per- and polyfluoroalkyl substances (PFAS), particulate matter (PMs), diesel exhaust (DE) and nano materials. The exposure time can be freely selected based on the need, from the beginning of development (embryonic day 0, ED0) all the way to the day prior to hatch. The major exposure methods include air-cell injection, direct microinjection, and air-cell inhalation (originally developed in our lab), and the currently available endpoints include cardiac function (electrocardiography), morphology (histological assessments) and molecular biological assessments (immunohistochemistry, qRT-PCR, western blotting, etc.). Of course, the chicken embryo model has its own limitations, such as limited availability of antibodies. Nevertheless, with more laboratories starting to utilize this model, it can be used to make significant contributions to the study of developmental cardiotoxicities.

Exposure results 
Air cell injection 
Air cell injection can effectively expose developing chicken embryos to various agents, which may be subsequently detected in the collected samples (serum, tissue, etc.) of embryos/hatchling chickens. Here is an example, in which perfluorooctanoic acid (PFOA) was air-cell injected, and serum PFOA concentrations were then determined with Ultra-performance liquid chromatography-mass spectrometry. The serum concentrations corresponded with the injected doses, indicating the ef...

Watch this Scientific Journal Video about Using Chicken Embryo as a Powerful Tool in Assessment of Developmental Cardiotoxicities at JoVE.com

Using Chicken Embryo as a Powerful Tool in Assessment of Developmental Cardiotoxicities

Chicken embryos, as a classical developmental model, are used in our lab to assess developmental cardiotoxicities following exposure to various environmental contaminants. Exposure methods and morphological/functional assessment methods established are described in this manuscript.

Chicken embryos are a classical model in developmental studies. During the development of chicken embryos, the time window of heart development is ...

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JoVE Journal

Developmental Biology

4.1K Views.  Qingdao University. Chicken embryos, as a classical developmental model, are used in our lab to assess developmental cardiotoxicities following exposure to various environmental contaminants. Exposure methods and morphological/functional assessment methods established are described in this manuscript. 

Video: Using Chicken Embryo as a Powerful Tool in Assessment of Developmental Cardiotoxicities

Comprehensive Assessment of Germline Chemical Toxicity Using the Nematode Caenorhabditis elegans

Identifying the reproductive toxicity of the thousands of chemicals present in our environment has been one of the most tantalizing challenges in the field of environmental health. This is due in part to the paucity of model systems that can (1) accurately recapitulate keys features of reproductive processes and (2) do so in a medium- to high-throughput fashion, without the need for a high number of vertebrate animals.
We describe here an assay in the nematode C. elegans that allows the rapid identification of germline toxicants by monitoring the induction of aneuploid embryos. By making use of a GFP reporter line, errors in chromosome segregation resulting from germline disruption are easily visualized and quantified by automated fluorescence microscopy. Thus the screening of a particular set of compounds for its toxicity can be performed in a 96- to 384-well plate format in a matter of days. Secondary analysis of positive hits can be performed to determine whether the chromosome abnormalities originated from meiotic disruption or from early embryonic chromosome segregation errors. Altogether, this assay represents a fast first-pass strategy for the rapid assessment of germline dysfunction following chemical exposure.

Comprehensive Assessment of Germline Chemical Toxicity Using the Nematode Caenorhabditis elegans

We describe the detailed steps of a high-throughput chemical assay in the nematode Caenorhabditis elegans used to assess germline toxicity. In this assay, disruption of germline function following chemical exposure is monitored using a fluorescent reporter specific to aneuploid embryos.

Identifying the reproductive toxicity of the thousands of chemicals present in our environment has been one of the most tantalizing challenges in the ...

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 ...

Grafting of Beads into Developing Chicken Embryo Limbs to Identify Signal Transduction Pathways Affecting Gene Expression

Using chicken embryos it is possible to test directly the effects of either growth factors or specific inhibitors of signaling pathways on gene expression and activation of signal transduction pathways. This technique allows the delivery of signaling molecules at precisely defined developmental stages for specific times. After this embryos can be harvested and gene expression examined, for example by in situ hybridization, or activation of signal transduction pathways observed with immunostaining.
In this video heparin beads soaked in FGF18 or AG 1-X2 beads soaked in U0126, a MEK inhibitor, are grafted into the limb bud in ovo. This shows that FGF18 induces expression of MyoD and ERK phosphorylation and both endogenous and FGF18 induced MyoD expression is inhibited by U0126. Beads soaked in a retinoic acid antagonist can potentiate premature MyoD induction by FGF18.
This approach can be used with a wide range of different growth factors and inhibitors and is easily adapted to other tissues in the developing embryo.

By grafting beads soaked in growth factors or specific inhibitors of signaling pathways into developing embryos it is possible to directly test their effects in vivo. In this protocol beads are grafted into the limb bud to determine the effects of these molecules on gene expression and signal transduction.

Using chicken embryos it is possible to test directly the effects of either growth factors or specific inhibitors of signaling pathways on gene expression ...

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin condensation, microtubule-kinetochore attachment, chromosome segregation, and cytokinesis in a small time frame. Errors in the delicate process can result in human disease, including birth defects and cancer. Traditional approaches investigating human mitotic disease states often rely on cell culture systems, which lack the natural physiology and developmental/tissue-specific context advantageous when studying human disease. This protocol overcomes many obstacles by providing a way to visualize, with high resolution, chromosome dynamics in a vertebrate system, the zebrafish. This protocol will detail an approach that can be used to obtain dynamic images of dividing cells, which include: in vitro transcription, zebrafish breeding/collecting, embryo embedding, and time-lapse imaging. Optimization and modifications of this protocol are also explored. Using H2A.F/Z-EGFP (labels chromatin) and mCherry-CAAX (labels cell membrane) mRNA-injected embryos, mitosis in AB wild-type, auroraBhi1045, and esco2hi2865 mutant zebrafish is visualized. High resolution live imaging in zebrafish allows one to observe multiple mitoses to statistically quantify mitotic defects and timing of mitotic progression. In addition, observation of qualitative aspects that define improper mitotic processes (i.e., congression defects, missegregation of chromosomes, etc.) and improper chromosomal outcomes (i.e., aneuploidy, polyploidy, micronuclei, etc.) are observed. This assay can be applied to the observation of tissue differentiation/development and is amenable to the use of mutant zebrafish and pharmacological agents. Visualization of how defects in mitosis lead to cancer and developmental disorders will greatly enhance understanding of the pathogenesis of disease.

Mitosis is critical to every living organism and defects often lead to cancer and developmental disorders. Using this imaging protocol and zebrafish as a model system, researchers can visualize mitosis in a live vertebrate organism and the multitude of defects that arise when mitotic processes are defective.

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin ...

In Ovo Electroporation in the Chicken Auditory Brainstem

Electroporation is a method that introduces genes of interest into biologically relevant organisms like the chicken embryo. It is long established that the chicken embryo is an effective research model for studying basic biological functions of auditory system development. More recently, the chicken embryo has become particularly valuable in studying gene expression, regulation and function associated with hearing. In ovo electroporation can be used to target auditory brainstem regions responsible for highly specialized auditory functions. These regions include the chicken nucleus magnocellularis (NM) and nucleus laminaris (NL). NM and NL neurons arise from distinct precursors of rhombomeres 5 and 6 (R5/R6). Here, we present in ovo electroporation of plasmid-encoded genes to study gene-related properties in these regions. We show a method for spatial and temporal control of gene expression that promote either gain or loss of functional phenotypes. By targeting auditory neural progenitor regions associated with R5/R6, we show plasmid transfection in NM and NL. Temporal regulation of gene expression can be achieved by adopting a tet-on vector system. This is a drug inducible procedure that expresses the genes of interest in the presence of doxycycline (Dox). The in ovo electroporation technique &#8211; together with either biochemical, pharmacological, and or in vivo functional assays &#8211; provides an innovative approach to study auditory neuron development and associated pathophysiological phenomena.

In Ovo Electroporation in the Chicken Auditory Brainstem

Auditory brainstem neurons of avians and mammals are specialized for fast neural encoding, a fundamental process for normal hearing functions. These neurons arise from distinct precursors of embryonic hindbrain. We present techniques utilizing electroporation to express genes in the hindbrain of chicken embryos to study gene function during auditory development.

Electroporation is a method that introduces genes of interest into biologically relevant organisms like the chicken embryo. It is long established that ...

Organotypic Culture Method to Study the Development Of Embryonic Chicken Tissues

The embryonic chicken is commonly used as a reliable model organism for vertebrate development. Its accessibility and short incubation period makes it ideal for experimentation. Currently, the study of these developmental pathways in the chicken embryo is conducted by applying inhibitors and drugs at localized sites and at low concentrations using a variety of methods. In vitro tissue&#160;culturing is a technique that enables the study of tissues separated from the host organism, while simultaneously bypassing many of the physical limitations present when working with whole embryos, such as the susceptibility of embryos to high doses of potentially lethal chemicals. Here, we present an organotypic culturing protocol for culturing the embryonic chicken half head in vitro, which presents new opportunities for the examination of developmental processes beyond the currently established methods.

Here, we present an organotypic culturing protocol to grow embryonic chicken organs in vitro. Using this method, the development of embryonic chicken tissue can be studied, while maintaining a high degree of control over the culture environment.

The embryonic chicken is commonly used as a reliable model organism for vertebrate development. Its accessibility and short incubation period makes it ...

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 ...

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits are required to control this behavior. Failure to produce the rhythmic pattern of contractions can be indicative of neurological abnormalities. We previously found that defects in protein O-mannosylation, a posttranslational protein modification, affect the axon morphology of sensory neurons and result in abnormal coordinated muscle contractions in embryos. Here, we present a relatively simple method for recording and analyzing the pattern of peristaltic muscle contractions by live imaging of late stage embryos up to the point of hatching, which we used to characterize the muscle contraction phenotype of protein O-mannosyltransferase mutants. Data obtained from these recordings can be used to analyze muscle contraction waves, including frequency, direction of propagation and relative amplitude of muscle contractions at different body segments. We have also examined body posture and taken advantage of a fluorescent marker expressed specifically in muscles to accurately determine the position of the embryo midline. A similar approach can also be utilized to study various other behaviors during development, such as embryo rolling and hatching.

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Here, we present a method to record embryonic muscle contractions in Drosophila embryos in a non-invasive and detail-oriented manner.

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits ...

X-ray Diffraction of Intact Murine Skeletal Muscle as a Tool for Studying the Structural Basis of Muscle Disease

Transgenic mouse models have been important tools for studying the relationship of genotype to phenotype for human diseases including those of skeletal muscle. Mouse skeletal muscle has been shown to produce high quality X-ray diffraction patterns on third generation synchrotron beamlines providing an opportunity to link changes at the level of the genotype to functional phenotypes in health and disease by determining the structural consequences of genetic changes. We present detailed protocols for preparation of specimens, collecting the X-ray patterns and extracting relevant structural parameters from the X-ray patterns that may help guide experimenters wishing to perform such experiments for themselves.

We present detailed protocols for performing small-angle X-ray diffraction experiments using intact mouse skeletal muscles. With the wide availability of transgenic mouse models for human diseases, this experimental platform can form a useful test bed for elucidating the structural basis of genetic muscle diseases

Transgenic mouse models have been important tools for studying the relationship of genotype to phenotype for human diseases including those of skeletal ...

Assessment of Oxidative Damage in the Primary Mouse Ocular Surface Cells/Stem Cells in Response to Ultraviolet-C (UV-C) Damage

The ocular surface is subjected to regular wear and tear due to various environmental factors. Exposure to UV-C radiation constitutes an occupational health hazard. Here, we demonstrate the exposure of primary stem cells from the mouse ocular surface to UV-C radiation. Reactive oxygen species (ROS) formation is the readout of the extent of oxidative stress/damage. In an experimental in vitro setting, it is also essential to assess the percentage of dead cells generated due to oxidative stress. In this article, we will demonstrate the 2&#39;,7&#39;-Dichlorofluoresceindiacetate (DCFDA) staining of UV-C exposed mouse primary ocular surface stem cells and their quantification based on the fluorescent images of DCFDA staining. DCFDA staining directly corresponds to ROS generation. We also demonstrate the quantification of dead and live cells by simultaneous staining with propidium iodide (PI) and Hoechst 3332 respectively and the percentage of DCFDA (ROS positive) and PI positive cells.

Assessment of Oxidative Damage in the Primary Mouse Ocular Surface Cells/Stem Cells in Response to Ultraviolet-C UV-C Damage

This protocol demonstrates the simultaneous detection of reactive oxygen species (ROS), live cells, and dead cells in live primary cultures from mouse ocular surface cells. 2&#39;,7&#39;-Dichlorofluoresceindiacetate, propidium iodide, and Hoechst staining are used to assess the ROS, dead cells, and live cells, respectively, followed by imaging and analysis.

The ocular surface is subjected to regular wear and tear due to various environmental factors. Exposure to UV-C radiation constitutes an occupational ...