The authors thank their funding sources, Howard Hughes Medical Institute Undergraduate Science Education Program to the College of William and Mary; Grant sponsor: NIH (M.S.S); Grant number: R15NS067566. They also acknowledge support from the College of William and Mary, Department of Biology and College of Arts and Sciences for assistance with animal care.

Ethics Statement: The methods were conducted with domesticated zebra finches from the breeding colony at the College of William and Mary. All procedures followed RSPCA guidelines23 and were approved by the College of William and Mary&#8217;s OLAW (Office of Laboratory Animal Welfare) Animal Welfare Assurance (#A3713-01) and had Institutional Animal Care and Use Committee (IACUC) approval (#2013-06-02-8721-dacris).
1. Egg Collection and Incubation
<ol>
	<li>Establish zebra finch pairs on a 14:10 light: dark cycle. Supply food, water, hay, and a nest box ad libitum. Note: Routine care and breeding for zebra finches has been well described23.</li>
	<li>Prepare a standard chick incubator with tilting shelves. Note: Maintain the artificial incubator at 37.5 (+/- 1) &#176;C with 80 - 95% humidity by adding water to the bottom of the incubator on a daily basis. Allow the incubator to equilibrate for two days before using.
	<ol>
		<li>Select small feed cups (5 x 7.6 cm) at least 2.5 cm deep to hold eggs in the incubator. Line the bottom and lower edges of the cup with two layers of paper towel so that the cup is padded, while still allowing the eggs to easily roll. Place these cups on the tilting shelves of the incubator. Note: Too much padding can inhibit rolling, which will prevent successful incubation due to embryonic adhesion to the shell interior.</li>
	</ol>
	</li>
	<li>Collect the eggs at two hours following the onset of the light cycle. Note: This minimizes discrepancies in stage development for a given incubation time due to parental incubation.
	<ol>
		<li>Pick up eggs gently with thumb and forefinger at the tip and base along the length of the egg. Use a dull, soft 4B graphite pencil to label the date laid and time collected from the nest. Note: The softer 4B pencil reduces the risk of the pencil breaking the fragile eggshell.</li>
		<li>1.3.2) Place 1 - 5 labeled eggs in each cup in the incubator so that eggs can freely roll to prevent embryonic adhesion to the interior of the shell. Note: Placing more than 5 eggs in a cup may prevent rolling of the individual eggs and reduce viability of embryos.</li>
	</ol>
	</li>
</ol>
2. Removal of Embryo from Egg
<ol>
	<li>To prepare for dissection, assemble the following materials: a clean scalpel, fine tipped forceps, and two extra fine tipped forceps. Place 10 x 10 cm weigh paper on the base of the dissecting scope to provide a clean, non-absorbent surface for dissection. Note: The weigh paper is crucial because it allows easy manipulation of the yolk and is the best surface for cutting the delicate membranes with a scalpel.</li>
	<li>Prepare an aliquot of phosphate buffer saline (1X PBS) in a 50 ml polypropylene tube, and acquire at least three transfer pipettes and a small waste bucket. If fixing the embryos, prepare an aliquot of 4% paraformaldehyde (PFA).Caution: PFA vapors are toxic, and all steps involving PFA should be done inside a fume hood to minimize exposure. If flash freezing the embryos, obtain liquid nitrogen and keep it near the dissecting scope. Sterility is not an issue for this dissection protocol, but ensure all materials are clean.</li>
	<li>Remove the egg from the incubator at the time point needed for the desired stage as described in the zebra finch staging guide22. Note: Remove eggs after 36 hr of incubation to dissect stage 6 embryos (Figure&#160;4A)22 and remove eggs after 56 hr of incubation to dissect stage 12 embryos (Figure&#160;3A)22.</li>
	<li>Using a fiber optic illuminator lamp, candle the egg by holding it along its vertical axis and shine light through the egg to illuminate the interior. Place the tip of the light behind the egg and locate the yolk and the developing embryo.
	<ol>
		<li>Position the scalpel on the side opposite the yolk and cut along the egg from tip to base with faint pressure.</li>
	</ol>
	</li>
	<li>Remove the contents of the egg by carefully applying pressure on the tip and base of the egg with the thumb and the forefinger, or by gently prying apart the eggshell along the cut with forceps. Open the egg directly above the weigh paper so that the yolk gently rolls out.</li>
	<li>Examine the yolk for the white zone of junction, which is visible as a faint white ring encircling embryonic development and orient the yolk using extra fine tipped forceps so that the embryo is located in the center. 
	Note: If a faint white circle on the surface of the yolk is not observed, take the fine forceps and gently roll the yolk over until the embryo is on top of the yolk. Note: For stages 1 - 922, the embryonic disk is less than 6 mm in diameter and is visible as a faint, slightly opaque disk on the surface of the yolk - see Figure 1 for reference. For stages 10 and older22, blood pools allow improved visibility of the embryo, but take care to avoid puncturing the yolk sac, as this makes locating the embryo difficult.</li>
</ol>
3. Separation of Embryo from Extra-embryonic Tissue
<ol>
	<li>Puncture the edges of the yolk to relieve pressure (Figure 1B), and make single cuts across the length of the yolk diameter alongside the embryonic disk. Repeat making these diagonal lines until the section of the yolk containing the embryo has been successfully separated (Figure 1B). Note: This step allows the yolk mass to remain intact, but the reduced pressure on the surface of the yolk allows for more precision when cutting around the embryo, preventing damage to embryonic structures.</li>
	<li>Remove the edges of the yolk with a transfer pipette. Note: Remove as much yolk as possible, but leave a small amount so that the embryo does not adhere to the dry surface of the weigh paper and tear.</li>
	<li>Wash the embryonic disk by dispensing 1X PBS at a 45&#176; angle towards the bottom of the embryo. Place the tip of the transfer pipette next to&#8212;not above&#8212;the embryonic disk. If additional yolk needs to be removed, separate any yolk with the scalpel on the weigh paper before transferring the embryo to the Petri dish. Note: This method removes the embryo from the surface of the weigh paper.</li>
	<li>Transfer the embryo using a transfer pipette along with a minimal volume of 1X PBS to a small, plastic Petri dish. Wash the embryo by adding more 1X PBS into the Petri dish and dripping 1X PBS near, but not directly onto, the embryo. Swirl the Petri dish to wash and remove residual yolk, tilt the Petri dish, and remove waste 1X PBS with the transfer pipette. 
	NOTE: When the 1X PBS is removed, the embryo typically does not adhere to the bottom of the Petri dish because the plastic surface is slippery with residual 1X PBS. However, more 1X PBS can be added if the embryo sticks to the dish. 
	If fixing the embryo, follow steps 3.5 and 3.6. If flash freezing the embryo, skip to step 3.8 immediately.</li>
	<li>If fixing the embryo for in situ hybridization, immediately add 4% PFA to the Petri dish to submerge the embryo. Drip 4% PFA directly onto the top of the embryo in order to flatten it; this prevents the embryo from curling. Fix the embryo in 4% PFA at 4 &#176;C for 12 hr.
	<ol>
		<li>After fixation, dehydrate embryos in graded methanol (MeOH) solutions and store at -20 &#176;C in 100% MeOH.</li>
	</ol>
	</li>
	<li>If the embryo is between stages 1 -&#160;822,&#160;remove the vitelline membrane adhered to the embryo.&#160;Note: This step can be done during or after fixation. Embryos younger than stage 822 are not easily visualized because they adhere to the vitelline membrane, obscuring key structures as seen in Figure 2A.
	<ol>
		<li>Grip the edge of the membrane that extends beyond the zone of junction with extra fine forceps. Carefully flip the embryo over multiple times to wash away residual yolk granules and to loosen the adherence of the embryonic disk to the vitelline membrane. Note: Do not touch the center of the embryo, as this will damage embryonic structures.</li>
		<li>If a gap does not appear between the zone of junction and the vitelline membrane, gently scratch the zone of junction with the extra fine tipped forceps to loosen it from the vitelline membrane. Grip the vitelline membrane with the extra fine tipped forceps and gently pull it away from the embryo. If necessary, gently pull the embryo away from the vitelline membrane by gripping the peripheral edge of the embryonic disk at the zone of junction.</li>
		<li>Discard the vitelline membrane in biohazard 1 waste (biosafety level 1) after removal.</li>
	</ol>
	</li>
	<li>When conducting an ISH assay on young embryonic stages, follow standard whole mount ISH protocols for chick embryos24, 25, but consider the following suggestions.
	<ol>
		<li>To reduce damage to the embryos during the ISH procedure, use a single 5 ml glass vial with a screw cap for each embryo. Only fill vials with 2 - 3 ml of solution, ensuring that the embryos are fully submerged. Nutate vials vertically by placing 5 ml vials into a styrofoam rack (or any rack that will keep the vials secure) that is secured to a nutator. Note: This precaution prevents the embryo from being torn, which can occur when it comes into contact with the lid of the vial during horizontal nutation.</li>
		<li>For embryos stage 0 - 622, treat with 5 &#956;g/ml proteinase K in 1X PTw for 5 min at room temperature. Treat embryos staged 7 - 1222 with 10 &#956;g/ml proteinase K in 1X PTw for 10 min at 37 &#176;C to reduce background staining.</li>
		<li>To produce a sufficient color reaction to detect low levels of gene expression in stages 1 - 1022, incubate with 10 &#956;g/ml probe concentration for 12 hr. For older stages, incubate with 1 &#956;g/ml probe concentration at 60 &#176;C.</li>
	</ol>
	</li>
	<li>If flash-freezing the embryo, quickly add 2 - 3 ml 1X PBS to the Petri dish following dissection and tilt the Petri dish to separate the embryo from the yolk granules.&#160;Remove liquid and repeat 2 - 3 times to remove all yolk. Using fine tipped forceps, transfer embryo to a pre-labeled microcentrifuge tube and flash freeze in liquid nitrogen before storing at -80 &#176;C.</li>
</ol>
4. EdU Cell Proliferation Assay. Incorporation and Detection of EdU in Zebra Finch Embryos.
<ol>
	<li>Candle the egg using a fiber optic illuminator lamp to locate the embryo or yolk.</li>
	<li>Mark the side of the egg opposite to the location of the embryo or yolk within the egg to ensure that the embryo is not damaged during the microinjection process.</li>
	<li>Line a 60 mm plastic dish with modeling clay and mold it in the shape of a bowl to hold the egg in the desired orientation. The marked spot should be oriented to allow insertion of the microinjection needle using the micromanipulator. Note: The clay will stabilize the egg during microinjection and throughout the incubation in step 4.11.</li>
	<li>Pull two glass capillary microinjection needles. Blunt one needle to poke a hole into the egg at the marked spot. Prepare the second microinjection needle for injection by backloading it with mineral oil and inserting it into the microinjector.</li>
	<li>Set the volume of solution released per injection to 59.8 nl on the microinjector.</li>
	<li>Load the microinjection needle with the 10 mM stock EdU solution.</li>
	<li>Create a hole in the shell on the marked spot with the blunted glass capillary, taking care to not shatter the shell or drop pieces of the shell into the egg cavity. Orient the egg so that the hole is at a 45&#176;&#160;angle, allowing the microinjection needle to be inserted directly through the cavity to the yolk or embryo.</li>
	<li>Using the micromanipulator, insert the loaded needle approximately 1.0 cm into the cavity.</li>
	<li>Inject the desired amount of the EdU solution directly onto the developing embryo or yolk. Note: A maximum of 478 nl of EdU can be injected without resulting in embryonic death.</li>
	<li>Immediately wrap the egg and the clay holder with multiple layers of plastic wrap to cover the egg and the dish to prevent desiccation of the embryo during incubation. Tape the edges of the plastic wrap to the bottom of the dish to prevent the plastic from unwrapping during incubation. Note: The plastic wrap should only be removed immediately before dissection.</li>
	<li>Allow the newly proliferating cells to incorporate the EdU by incubating the egg at 37 &#176;C until the desired stage is reached. After injection, do not return egg to tilting shelves in incubator. Place the plastic-wrapped clay lined dish on stable surface inside the incubator.</li>
	<li>Remove the plastic wrap and dissect the embryo following the aforementioned protocol. Fix the embryo in 4% PFA and incubate 12 hr at 4 &#176;C as described above. After fixation, wash with 100% ethanol (EtOH) for 5 min and store in fresh 100% EtOH at&#160;-20 &#176;C until further analysis.</li>
</ol>
5. EdU &#8220;Click&#8221; Reaction Protocol
The following steps are all performed in glass vials.
<ol>
	<li>Rehydrate embryos with successive 5 min washes of the following:</li>
	<li>EtOH, 75% EtOH and 25% sterile deionized distilled (sdd) water, 50% EtOH and 50% sdd water, 25% EtOH and 75% 1X PTw, 100% 1X Ptw</li>
	<li>Wash three times in 100% 1X PTw for 10 min each.</li>
	<li>Dilute the reaction buffer additive (Kit Component F) by combining 1 part of the 10X stock buffer solution (10 &#956;l) to 9 parts sdd water (90 &#956;l).</li>
	<li>Prepare the reaction mix in a separate tube by mixing the following: 875 &#956;l 1X PBS, 20 &#956;l CuSO4, 5 &#956;l azide, 100 &#956;l diluted reaction buffer additive. Note: Volume of the reaction mix can be scaled down. The reaction will work if the embryo is completely covered in the reaction mixture. Add the diluted reaction buffer additive (step 5.3) immediately before use. All subsequent steps are performed in the dark. Cover the embryos with foil to protect from the light.</li>
	<li>Cover the vials with aluminum foil, and incubate them vertically at room temperature for two hours by placing them in a styrofoam rack attached to a nutator.</li>
	<li>Wash the embryos 3 times in fresh 1X PBS for 10 min each.</li>
	<li>Incubate the embryos in fresh 4% PFA overnight at 4 &#176;C.</li>
	<li>Wash 3 times in 1X PBS for 10 min each and store in fresh 1X PBS at 4 &#176;C. Cover the embryos with foil until further analysis.</li>
</ol>

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B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Adverse+effects+of+fipronil+on+avian+reproduction+and+development:+maternal+transfer+of+fipronil+to+eggs+in+zebra+finch+Taeinopygia+guttata+and+in+ovo+exposure+in+chickens+Gallus+domesticus.">Adverse effects of fipronil on avian reproduction and development: maternal transfer of fipronil to eggs in zebra finch Taeinopygia guttata and in ovo exposure in chickens Gallus domesticus.</a> Ecotoxicology. 20, 653-660 (2011).</li><li>Hallinger, K. K., Zabransky, D. J., Kazmer, K. A., Cristol, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Birdsong+differs+between+mercury-polluted+and+reference+sites.">Birdsong differs between mercury-polluted and reference sites.</a> The Auk. 127, 156-161 (2010).</li><li>Chen, C. C., Wada, K., Jarvis, E. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Radioactive+in+situ+Hybridization+for+Detecting+Diverse+Gene+Expression+Patterns+in+Tissue.">Radioactive in situ Hybridization for Detecting Diverse Gene Expression Patterns in Tissue.</a> Journal of Visualized Experiments. , (2012).</li><li>Chen, C. C., Balaban, E., Jarvis, E. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Interspecies+Avian+Brain+Chimeras+Reveal+That+Large+Brain+Size+Differences+Are+Influenced+by+Cell&#8211;Interdependent+Processes.">Interspecies Avian Brain Chimeras Reveal That Large Brain Size Differences Are Influenced by Cell&#8211;Interdependent Processes.</a> PLoS One. 7, (2012).</li><li>Chen, C. C., Winkler, C. M., Pfenning, A. R., Jarvis, E. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Molecular+profiling+of+the+developing+avian+telencephalon:+Regional+timing+and+brain+subdivision+continuities.">Molecular profiling of the developing avian telencephalon: Regional timing and brain subdivision continuities.</a> The Journal of Comparative Neurology. 521, 3666-3701 (2013).</li></ol>

The authors have nothing to disclose.

Recent development of an embryological staging guide22 and genome annotation make the zebra finch a desirable model organism for developmental studies. However, the small size and fragility of the zebra finch embryos, which range from 3 to 7 mm in stages 1 - 1022, can make dissections difficult11,14. Locating and cleanly removing embryos from the surface of the yolk can be challenging. This protocol provides sufficient detail to perform the procedure with ease. This protocol demonstrates the critical steps that are not typically known, but are necessary to ensure a successful dissection. For example, it is essential to leave a small layer of yolk between the embryo and sheet of weigh paper to preclude sticking.
Both the identification and removal of the embryo can be difficult. To troubleshoot identifying the embryo on the surface of the yolk, shine light directly above the yolk after it has been removed from the egg, and look at the yolk at a 45&#176; angle to find the embryo. Once the embryo is located, cut the yolk on weigh paper taking care to not tear the embryo.
If further applications include imaging for anatomical differences, in situ hybridization, or cell proliferation assays, it is important to remove the vitelline membrane in stage 1 - 822 for better visualization of structures. If experiencing difficulties when removing the yolk or the vitelline membrane in early stages, fix the embryo in 4 % PFA before washing it in 1X PBS to reduce embryonic fragility. By first removing the vitelline membrane during dissection as described, structures are clearly visible and intact in zebra finch embryos after performing an in situ hybridization.
A limitation of EdU is that administration of dosage volumes over 478 nl leads to embryonic fatality. However, the vast dosage range allows varying levels of proliferative cell tagging.
The &#8220;click&#8221; reaction used in this kit is the Copper(I)-Catalyzed-Alkyne-Azide-Cycloaddition (Cu(I)AAC). In this specific reaction, an alkyne-containing thymidine analogue molecule (EdU) is incorporated by actively dividing cells. The alkyne group in the EdU protrudes from the helical structure of the DNA, and is detected by exposure to an azide molecule conjugated to a green, fluorescent molecule which binds to the free alkyne group. The green fluorescence shows the newly proliferating cells in the embryo. The bio-orthogonality of the azide and the alkyne groups prevents non-specific staining because these reactive species are not naturally present in organisms. Also, because the DNA does not need to be denatured in order for the reaction to occur, further DNA-dependent analysis can be easily performed27.
An inherent limitation of this method is the small size and fragility of zebra finch embryos. Removing the vitelline membrane of embryos stages 1 - 1522 can result in damaging embryonic structures if not performed with caution. However, this protocol simplifies the dissection method, allowing investigators to use early embryonic stages to examine structural anomalies and gene expression that have not been previously studied in depth. This protocol opens the way for a plethora of cellular-molecular assays that will allow investigators to determine the developmental origins of adult phenotypes. For example, it will be possible to examine gene expression implicated in vocal learning under various environmental conditions or following pharmacological treatments at the earliest stages of development28,29,30,31. Although not demonstrated in this paper, this method potentially allows for other procedures such as radioactive in situ hybridization on zebra finch tissue sections and electroporation/in ovosurgery32,33,34. Given that the zebra finch has been established as an important model organism within a vast body of literature, such studies provide untapped opportunities to link developmental mechanisms with adult physiology and behavior, in particular the development of language7,9.

The overall goal of this technique is to obtain zebra finch (Taeniopygiaguttata) embryos from the earliest stages of embryogenesis for use in a wide range of developmental studies. Zebra finch have become the predominant songbird model organism and have been used extensively in a variety of fields, including toxicology1,2, behavior3, memory and learning4,5,6, comparative neuroanatomy7,8, and language development9,10. As the only songbird with a sequenced genome, the zebra finch allows molecular and genetic study of the Passeriformes order, which represents over 50% of known bird species11,12,13.
Despite the use of adult and juvenile zebra finch in a diverse array of fields, few studies have been performed on zebra finch embryos, particularly during early stages of development. This can be attributed to the small size of their eggs and embryos, and their newer status as a model organism14,15,16 for studies in which the chicken (Gallus gallusdomesticus) was previously used as a predominant model system17,18,19,20,21. However, as non-vocal learners, chickens are not an appropriate model system for studying the genetic basis of vocal learning, development of vocal learning, heritability, behavior, and the cortical-basal ganglia circuitry involved in motor learning10.
It is important to note that zebra finch embryos are much more delicate and more easily damaged than chick embryos during dissection and molecular procedures. In particular, greater care is required when performing permeabilization steps on zebra finch embryos. Strong detergents and enzymes that would not harm a chick embryo can damage zebra finch embryos. In terms of general care, it is necessary to put zebra finch eggs in small cups before placement in an incubator to prevent them from breaking when rolling during incubation.
Zebra finch are amenable to behavioral studies, easily and prolifically breed year-round in captivity, and are vocal learners. These characteristics allow the use of zebra finch to address the need for a model organism that integrates development, genetics, and behavioral aspects of language. The dissections methods detailed below, combined with a recently developed staging guide specific to zebra finch22,&#160;make the zebra finch an increasingly useful standardized developmental model organism. However, obtaining embryos at early stages can be daunting. This protocol allows investigators to easily obtain early stage embryos. Studies investigating the early development and the molecular developmental basis of complex behaviors in zebra finch, or the toxicological effects on development in other small, passerine birds will find this dissection methodology useful.

<table><tbody><tr><td>Chicken egg incubator</td><td></td><td></td><td>We use a Picture Window Hova-Bator Incubator, Circulated Air Model&amp;nbsp;&amp;nbsp;&amp;nbsp;</td></tr><tr><td>Click-iT EdU Alexa Fluor 488 Imaging kit</td><td>Invitrogen</td><td>C10337</td><td>Detection of cell proliferation.&amp;nbsp;</td></tr><tr><td>Dissection microscope&amp;nbsp;</td><td></td><td></td><td>We use Olympus SZ61</td></tr><tr><td>7&#039;&#039; Drummond capillary for Nanoject II injector</td><td>Drummond</td><td>3-00-203-G/X</td><td></td></tr><tr><td>Drummond Nanoject II microinjector</td><td>Drummond</td><td></td><td></td></tr><tr><td>Dumont Tweezers #55</td><td>World Precision Instruments</td><td>14099</td><td></td></tr><tr><td>Ethanol (EtOH)</td><td></td><td></td><td></td></tr><tr><td>50 ml Falcon tube (polypropylene)</td><td>Fisher Scientific</td><td>06-443-18&amp;nbsp;</td><td></td></tr><tr><td>FastPrep Lysis Matrix H tubes</td><td>MP biomedicals</td><td>6917-100</td><td>Used for RNA extraction, used to homogenize embryos</td></tr><tr><td>Fiber optic illuminator lamps (High Intensity Illuminator)</td><td>Dolan-Jenner Industries&amp;nbsp;</td><td>Fiber-Lite MI-150&amp;nbsp;</td><td></td></tr><tr><td>Glass vials with screw cap (DEPC treated)</td><td>Fisher Scientific</td><td>03-338A</td><td></td></tr><tr><td>Microcentrifuge eppe tube (1.5 ml)</td><td>Fisher Scientific</td><td>05-408-129</td><td></td></tr><tr><td>Mineral oil</td><td>Sigma-Aldrich</td><td>M-8410</td><td></td></tr><tr><td>Modeling Clay</td><td></td><td></td><td>Any brand</td></tr><tr><td>Narshige PB-7 needle puller</td><td>Narshige</td><td></td><td></td></tr><tr><td>Omni Bead Ruptor Homogenizer</td><td>OMNI International</td><td>19-010</td><td>Used for RNA extraction</td></tr><tr><td>4% Paraformaldehyde (4% PFA): 20 ml 8% PFA (32 g paraformaldehyde, 350 ml sdd H&lt;sub&gt;2&lt;/sub&gt;O, adjust pH to 7.6, 400 ml sdd H&lt;sub&gt;2&lt;/sub&gt;O), 20 ml 2x PBS</td><td>Fixation of embryos. Caution, is harmful and do not inhale.&amp;nbsp;</td><td></td><td></td></tr><tr><td>Phosphate buffered saline (1x PBS): 200 ml 10x PBS, 1,800 ml Barnstead H&lt;sub&gt;2&lt;/sub&gt;O, adjust pH to 7.4</td><td></td><td></td><td></td></tr><tr><td>Phosphate buffered saline (10x PBS): 800 ml Barnstead, 2.013 g KCL, 80.063 g NaCl, 2.722 g KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt; monobasic, 14.196 g Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt; dibasic, 200 ml Barnstead H&lt;sub&gt;2&lt;/sub&gt;O, adjust pH to 6.5</td><td></td><td></td><td></td></tr><tr><td>Phosphate buffered saline with 0.1% Tween-20 (1x PTw): 1,800 ml Barnstead H&lt;sub&gt;2&lt;/sub&gt;O, 200 ml 10x PBS, adjust pH to 7.4, 2 ml Tween-20</td><td></td><td></td><td></td></tr><tr><td>35 ml Plastic Petri dish&amp;nbsp;</td><td>Fisher Scientific</td><td>08-757-100A</td><td></td></tr><tr><td>plastic wrap</td><td></td><td></td><td>Any brand</td></tr><tr><td>Prepease RNA Spin Kit</td><td>PrepEase, Affymetrix</td><td>78766 1 KT</td><td>Used for RNA extraction, used to homogenize embryos</td></tr><tr><td>Reaction Mix: 875 &amp;mu;l 1xPBS, 100 mM 20 &amp;mu;l CuSO&lt;sub&gt;4&lt;/sub&gt; (Kit Component E), 5 &amp;mu;l Alexa Fluor 488 azide (Kit Component B), 100 &amp;mu;l diluted reaction buffer additive (Kit Component F)</td><td>Invitrogen</td><td>C10337</td><td>Detection of cell proliferation.&amp;nbsp;</td></tr><tr><td>sdd H&lt;sub&gt;2&lt;/sub&gt;O</td><td></td><td></td><td></td></tr><tr><td>Seed cup&amp;nbsp;</td><td></td><td></td><td>We use it as a container when incubating eggs</td></tr><tr><td>Stainless steel scalpel&amp;nbsp;</td><td></td><td></td><td></td></tr><tr><td>Standard nest box</td><td></td><td></td><td>We use Abba Plastic Finch Nestboxes</td></tr><tr><td>4 ml, Teflon Lined Cap, Glass Vials</td><td>Fisher Scientific</td><td>02-912-352</td><td></td></tr><tr><td>Transfer pipettes (polyethylene)</td><td>Fisher Scientific</td><td>13-711-7M&amp;nbsp;</td><td></td></tr><tr><td>4 x 4 weigh paper</td><td>Fisher Scientific</td><td>09-898-12B</td><td></td></tr></tbody></table>

dissection and downstream analysis of zebra finch embryos at early stages of development

The zebra finch (Taeniopygiaguttata) has become an increasingly important model organism in many areas of research including toxicology1,2, behavior3, and memory and learning4,5,6. As the only songbird with a sequenced genome, the zebra finch has great potential for use in developmental studies; however, the early stages of zebra finch development have not been well studied. Lack of research in zebra finch development can be attributed to the difficulty of dissecting the small egg and embryo. The following dissection method minimizes embryonic tissue damage, which allows for investigation of morphology and gene expression at all stages of embryonic development. This permits both bright field and fluorescence quality imaging of embryos, use in molecular procedures such as in situ hybridization (ISH), cell proliferation assays, and RNA extraction for quantitative assays such as quantitative real-time PCR (qtRT-PCR).&#160;This technique allows investigators to study early stages of development that were previously difficult to access.

The steps diagrammed in Figure 1 indicate the appearance of the embryo while attached to the vitelline membrane (A) and demonstrate the proper method to separate the embryo from the yolk (B). The embryo can be identified by the zone of junction, which is much lighter than the vitelline membrane. The embryo itself is often difficult to distinguish until the yolk is cut away. Once the embryo is dissected from the egg, it can be fixed or flash frozen for future use. If an in situ hybridization is planned for the dissected embryo, it is necessary to remove the vitelline membrane that is adhered to the embryo via the zone of junction. Figure 2 illustrates the improved visibility of the embryo once this membrane is removed (C), and the proper way to peel the vitelline membrane (A, B). After dissection and fixation, whole mount in situ hybridization was performed as seen in Figure 3 (A, A&#8217;, B, B&#8217;) and Figure 4 (A, A&#39;, B, B&#8217;) and Figure 5 (A, B, C) to detect differences in orthodenticlehomeobox 2 (Otx2) expression in embryos developmentally exposed to low doses of methylmercury. Figure 5 shows sense probe results, demonstrating lack of background. In Figure 4, despite being dissected from the egg at the same time point, the embryo developmentally exposed to methylmercury (MeHg) progressed to stage 522 (B, B&#8217;), while the control embryo developed to stage 622 (A, A&#8217;). The group of embryos dissected and shown in Figure 4 were collected from the nest and taken from the incubator at the same times. Although some natural variation is present in development, based on previous dissection data, it is unlikely that temperature fluctuations in the incubator would cause only the 2.4 ppm methylmercury embryos to be developmentally delayed. The differences in stages indicate changes in cell proliferation in embryos developmentally exposed to methylmercury.
Before dissection, EdU was injected into a day 2 egg and allowed to incubate overnight. After dissection and fixation of the stage 1622 embryo, EdU was visualized using &#8220;click&#8221; chemistry, allowing detection of proliferating cells as seen in Figure 6 (A, B, C). It is important to carefully monitor time points when placing eggs in the incubator and during dissections, as exposure to methylmercury or performing the EdU assay may disrupt developmental progression. The earliest injection was performed on day 0, which was the day of collection as specified in step 1.3. This embryo was dissected approximately 38 hours later (stage 722). The survival rate was found to be approximately 90% (same rate as control embryos) as long as the injection amount was under 478 nl.
This dissection methodology also allows for high quality RNA extraction. After dissecting stage 1622 embryos, an RNA extraction was performed according to manufacturer&#8217;s protocol with no optimization required, as seen in Figure 7. The removal of the vitelline membrane was unnecessary for RNA extraction and later qRT-PCR applications.
Note: All embryo figures are oriented so that the anterior and posterior regions are at the top and bottom of the images, respectively. 
<img alt="Figure 1" src="/files/ftp_upload/51596/51596fig1highres.jpg" /> 
Figure 1.&#160;Procedure for locating and dissecting zebra finch embryos, stages 1-1022. Locate embryo by gently rolling the yolk until the faint white disk is apparent (A). Once the embryo is located at the center of the yolk, the yolk is dissected in a stepwise fashion (B) where the first cut relieves pressure of the vitelline membrane (1) and subsequent cuts (2) border the zone of junction (zj) which is adhered to the vitelline membrane. Scale bars represent 1 mm.
<img alt="Figure 2" src="/files/ftp_upload/51596/51596fig2highres.jpg" /> 
Figure 2. Removal of vitelline membrane and visibility of embryonic structures. Following removal of the embryo from the yolk, place embryo in a Petri dish containing 4% PFA. If an in situ hybridization needs to be performed, visibility of the embryonic structures is essential and can be achieved by removing the vitelline membrane (A). Grip the vitelline membrane with extra fine tipped forceps and gently peel it away from the embryo by handling the embryo directly at the outermost edge, if necessary (B). Vitelline membrane removal increases clarity of embryonic structures, and allows embryos to be imaged or processed with in situ hybridization (C). Scale bars represent 1 mm. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/51596/51596fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/51596/51596fig3highres.jpg" /> 
Figure 3. Whole mount in situ hybridization performed on zebra finch embryos developmentally exposed to methylmercury. Expression patterns of orthodenticlehomeobox 2 (Otx2) were characterized in embryos exposed to 0.0 ppm methylmercury (A, A&#8217;) and 2.4 ppm methylmercury (B, B&#8217;) via parental diet. The dorsal (A) and ventral (A&#8217;) expression of Otx2 is visible throughout the midbrain and optic vesicles during stage 1222. The treatment group embryos were dissected at the same time point, but were developmentally delayed as seen in the dorsal (B) and ventral (B&#8217;) view of head structures, which are characteristic of stage 1122. Abbreviations: mb, midbrain; op, optic vesicle. Scale bars represent 1 mm.
<img alt="Figure 4" src="/files/ftp_upload/51596/51596fig4highres.jpg" /> 
Figure 4.&#160;Whole mount&#160;in situ&#160;hybridization performed on zebra finch embryos developmentally exposed to methylmercury. Expression patterns of&#160;orthodenticlehomeobox 2 (Otx2)&#160;were characterized in stage 622 embryos exposed to 0.0 ppm methylmercury (A, A&#8217;) and stage 522 embryos exposed to 2.4 ppm methylmercury (B, B&#8217;) via parental diet. Abbreviations: am, anterior margin of mesoderm; no, notochord, notochord mesoderm; po, proamnion, anterior blastopore; ps, primitive streak22. Scale bars represent 1 mm.
<img alt="Figure 5" fo:content-width="6in" src="/files/ftp_upload/51596/51596fig5highres.jpg" width="600" /> 
Figure 5.&#160;Whole mount in situ&#160;hybridization performed on zebra finch embryos using sense probe. (A) Stage 522&#160;embryo. (B) Early stage 622 embryo. (C) Stage 1122 embryo. Scale bars represent 1 mm.
<img alt="Figure 6" src="/files/ftp_upload/51596/51596fig6highres.jpg" /> 
Figure 6.EdU incorporation and detection in zebra finch embryos. EdU &#8220;click&#8221; chemistry was used to detect proliferating cells in a stage 1622 embryo (A, B, C). EdU is incorporated into the DNA in the place of thymidine26,27 and is detected using click chemistry27. Proliferation is clearly visible in the lateral edges of the somites, and the tailbud. Panel A shows proliferation occurring exclusively in the posterior of the embryo and also shows individual proliferative cells. Panel B shows the proliferative locations in the whole embryo. Panel C shows the anterior region, and shows the highly proliferative telencephalon (te) in greater detail. Abbreviations: af, amniotic fold; flb, forelimb bud; hlb, hindlimb bud; le, lens vesicle; ms, mesencephalon; mt, metencephalon; opc, optic cup; pa, pharyngeal arch; sm, somite mesoderm; tb, tailbud; te, telencephalon. Scale bars represent 1 mm. <a href="https://jove.unicartagenaproxy.elogim.com/files/ftp_upload/51596/51596fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" src="/files/ftp_upload/51596/51596fig7highres.jpg" /> 
Figure 7.&#160;Quality of RNA extracted from dissected zebra finch embryos. Control (0.0 ppm) and 1.2 ppm methylmercury embryos were dissected and flash frozen as described in Step 3.8. Each lane shows RNA extracted from two homogenized embryos from each treatment group.

Watch this Scientific Journal Video about Dissection and Downstream Analysis of Zebra Finch Embryos at Early Stages of Development at JoVE.com

Dissection and Downstream Analysis of Zebra Finch Embryos at Early Stages of Development

The zebra finch (Taeniopygiaguttata) is a valuable model organism; however, early stages of zebra finch development have not been extensively studied. The protocol describes how to dissect early embryos for developmental and molecular applications.

The zebra finch (Taeniopygiaguttata) has become an increasingly important model organism in many areas of research including toxicology1,2, behavior3, and ...

dissection-downstream-analysis-zebra-finch-embryos-at-early-stages

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

10.9K Views.  College of William and Mary. The zebra finch (Taeniopygiaguttata) is a valuable model organism; however, early stages of zebra finch development have not been extensively studied. The protocol describes how to dissect early embryos for developmental and molecular applications. 

Video: Dissection and Downstream Analysis of Zebra Finch Embryos at Early Stages of Development

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method described here allows time-lapse analysis of normal and impaired kidney development in zebrafish embryos by using a fluorescence dissecting microscope equipped for structured illumination and z-stack acquisition. To visualize nephrogenesis, transgenic zebrafish (Tg(wt1b:GFP)) with fluorescently labeled kidney structures were used. Renal defects were triggered by injection of an antisense morpholino oligonucleotide against the Wilms tumor gene wt1a, a factor known to be crucial for kidney development.
The advantage of the experimental setup is the combination of a zoom microscope with simple strategies for re-adjusting movements in x, y or z direction without additional equipment. To circumvent focal drift that is induced by temperature variations and mechanical vibrations, an autofocus strategy was applied instead of utilizing a usually required environmental chamber. In order to re-adjust the positional changes due to a xy-drift, imaging chambers with imprinted relocation grids were employed.
In comparison to more complex setups for time-lapse recording with optical sectioning such as confocal laser scanning&#160;or light sheet microscopes, a zoom microscope is easy to handle. Besides, it offers dissecting microscope-specific benefits such as high depth of field and an extended working distance.
The method to study organogenesis presented here can also be used with fluorescence stereo microscopes not capable of optical sectioning. Although limited for high-throughput, this technique offers an alternative to more complex equipment that is normally used for time-lapse recording of developing tissues and organ dynamics.

The method described here allows time-lapse analysis of organ development in zebrafish embryos by using a fluorescence dissecting microscope capable of performing optical sectioning and simple strategies of readjustment to correct focal and planar drift.

In order to understand organogenesis, the spatial and temporal alterations that occur during development of tissues need to be recorded. The method ...

Developmental Biology

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 ...

Dissection of Larval Zebrafish Gonadal Tissue

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex determination system, where several genes distributed throughout the genome collectively determine the sex identities of individual fish. Currently, the genes involved in regulating gonad development and how they work remain elusive. Normally, isolating gonadal tissue is the first step to examine the sex developmental processes. Here, we present a procedure to isolate gonadal tissue from 17 dpf (days post fertilization) and 25 dpf zebrafish larvae. The isolated gonadal tissue may be subsequently examined by morphology and gene expression profiling.

Here, we present a protocol for isolating gonadal tissue of larval zebrafish, which will facilitate investigations of zebrafish sex differentiation and maintenance.

Although wild zebrafish possess a ZZ/ZW sex-determination system, domesticated zebrafish have lost the sex chromosome. They utilize a polygenic sex ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Dissection and Lateral Mounting of Zebrafish Embryos: Analysis of Spinal Cord Development

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene knockdown approaches can be utilized to examine the molecular mechanisms underlying nervous system development. Second, large clutches of developmentally synchronized embryos provide large experimental sample sizes. Third, the optical clarity of the zebrafish embryo permits researchers to visualize progenitor, glial, and neuronal populations. Although zebrafish embryos are transparent, specimen thickness can impede effective microscopic visualization. One reason for this is the tandem development of the spinal cord and overlying somite tissue. Another reason is the large yolk ball, which is still present during periods of early neurogenesis. In this article, we demonstrate microdissection and removal of the yolk in fixed embryos, which allows microscopic visualization while preserving surrounding somite tissue. We also demonstrate semipermanent mounting of zebrafish embryos. This permits observation of neurodevelopment in the dorso-ventral and anterior-posterior axes, as it preserves the three-dimensionality of the tissue.

Developmental processes such as proliferation, patterning, differentiation, and axon guidance can be readily modeled in the zebrafish spinal cord. In this article, we describe a mounting procedure for zebrafish embryos, which optimizes visualization of these events.

The zebrafish spinal cord is an effective investigative model for nervous system research for several reasons. First, genetic, transgenic and gene ...

Neuroscience

A 3-D Tail Explant Culture to Study Vertebrate Segmentation in Zebrafish

Vertebrate embryos pattern their major body axis as repetitive somites, the precursors of vertebrae, muscle, and skin. Somites progressively segment from the presomitic mesoderm (PSM) as the tail end of the embryo elongates posteriorly. Somites form with regular periodicity and scale in size. Zebrafish is a popular model organism as it is genetically tractable and has transparent embryos that allow for live imaging. Nevertheless, during somitogenesis, fish embryos are wrapped around a large, rounding yolk. This geometry limits live imaging of PSM tissue in zebrafish embryos, particularly at higher resolutions that require a close objective working distance. Here, we present a flattened 3-D tissue culture method for live imaging of zebrafish tail explants. Tail explants mimic intact embryos by displaying a proportional slowdown of axis elongation and shortening of rostrocaudal somite lengths. We are further able to stall axis elongation speed through explant culture. This, for the first time, enables us to untangle the chemical input of signaling gradients from the mechanistic input of axial elongation. In future studies, this method can be combined with a microfluidic setup to allow time-controlled pharmaceutical perturbations or screening of vertebrate segmentation without any drug penetration concerns.

Here, we present the protocol for 3-D tissue culture of the zebrafish posterior body axis, enabling live study of vertebrate segmentation. This explant model provides control over axis elongation, alteration of morphogen sources, and subcellular resolution tissue-level live imaging.

Vertebrate embryos pattern their major body axis as repetitive somites, the precursors of vertebrae, muscle, and skin. Somites progressively segment from ...

Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology

Whole mount in situ hybridization (WISH) is a common technique in molecular biology laboratories used to study gene expression through the localization of specific mRNA transcripts within whole mount specimen. This technique (adapted from Albertson and Yelick, 2005) was used in an upper level undergraduate Comparative Vertebrate Biology laboratory classroom at Syracuse University. The first two thirds of the Comparative Vertebrate Biology lab course gave students the opportunity to study the embryology and gross anatomy of several organisms representing various chordate taxa primarily via traditional dissections and the use of models. The final portion of the course involved an innovative approach to teaching anatomy through observation of vertebrate development employing molecular techniques in which WISH was performed on zebrafish embryos. A heterozygous fibroblast growth factor 8 a (fgf8a) mutant line, ace, was used. Due to Mendelian inheritance, ace intercrosses produced wild type, heterozygous, and homozygous ace/fgf8a mutants in a 1:2:1 ratio. RNA probes with known expression patterns in the midline and in developing anatomical structures such as the heart, somites, tailbud, myotome, and brain were used. WISH was performed using zebrafish at the 13 somite and prim-6 stages, with students performing the staining reaction in class. The study of zebrafish embryos at different stages of development gave students the ability to observe how these anatomical structures changed over ontogeny. In addition, some ace/fgf8a mutants displayed improper heart looping, and defects in somite and brain development. The students in this lab observed the normal development of various organ systems using both external anatomy as well as gene expression patterns. They also identified and described embryos displaying improper anatomical development and gene expression (i.e., putative mutants).
For instructors at institutions that do not already own the necessary equipment or where funds for lab and curricular innovation are limited, the financial cost of the reagents and apparatus may be a factor to consider, as will the time and effort required on the part of the instructor regardless of the setting. Nevertheless, we contend that the use of WISH in this type of classroom laboratory setting can provide an important link between developmental genetics and anatomy. As technology advances and the ability to study organismal development at the molecular level becomes easier, cheaper, and increasingly popular, many evolutionary biologists, ecologists, and physiologists are turning to research strategies in the field of molecular biology. Using WISH in a Comparative Vertebrate Biology laboratory classroom is one example of how molecules and anatomy can converge within a single course. This gives upper level college students the opportunity to practice modern biological research techniques, leading to a more diversified education and the promotion of future interdisciplinary scientific research.

Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology

Whole mount in situ hybridization (WISH) was used in an upper level undergraduate Comparative Vertebrate Biology course in addition to vertebrate dissections. This gave students the opportunity to study gene expression patterns as well as gross anatomy, linking the study of molecular and organismal biology within one course.

Whole mount in situ hybridization (WISH) is a common technique in molecular biology laboratories used to study gene expression through the localization of ...

Refined Multiview Light Sheet Microscopy Protocol for Zebrafish Embryos

Light Sheet Microscopy Imaging and Mounting Strategies for Early Zebrafish Embryos

Light sheet microscopy has become the methodology of choice for live imaging of zebrafish embryos over long time scales with minimal phototoxicity. In particular, a multiview system, which allows sample rotation, enables imaging of entire embryos from different angles. However, in most imaging sessions with a multiview system, sample mounting is a troublesome process as samples are usually prepared in a polymer tube. To aid in this process, this protocol describes basic mounting strategies for imaging early zebrafish development between the 70% epiboly and early somite stages. Specifically, the study provides statistics on the various positions the embryos default to at the 70% epiboly and bud stages within the chorion. Furthermore, it discusses the optimum number of angles and the interval between angles required for imaging whole zebrafish embryos at the early stages of development so that cellular-scale information can be extracted by fusing the different views. Finally, since the embryo covers the entire field of view of the camera, which is required to obtain a cellular-scale resolution, this protocol details the process of using bead information from above or below the embryo for the registration of the different views.

Author Spotlight: Strategies for Mounting Zebrafish Embryos for High-Resolution Multiview Light-Sheet Microscopy &#8212; Techniques for Imaging and Image Reconstruction

A sample preparation strategy for imaging early zebrafish embryos within an intact chorion using a light-sheet microscope is described. It analyzes the different orientations that embryos acquire within the chorion at the 70% epiboly and bud stages and details imaging strategies for obtaining cellular-scale resolution throughout the embryo using the light-sheet system.

Light sheet microscopy has become the methodology of choice for live imaging of zebrafish embryos over long time scales with minimal phototoxicity. In ...

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical clarity and amenability to genetic manipulation, the zebrafish embryo has become a popular model organism with which to perform time-lapse analysis of morphogenesis in living embryos. Confocal imaging of a live zebrafish embryo requires that a tissue of interest is persistently labeled with a fluorescent marker, such as a transgene or injected dye. The process demands that the embryo is anesthetized and held in place in such a way that healthy development proceeds normally. Parameters for imaging must be set to account for three-dimensional growth and to balance the demands of resolving individual cells while getting quick snapshots of development. Our results demonstrate the ability to perform long-term in vivo imaging of fluorescence-labeled zebrafish embryos and to detect varied tissue behaviors in the cranial neural crest that cause craniofacial abnormalities. Developmental delays caused by anesthesia and mounting are minimal, and embryos are unharmed by the process. Time-lapse imaged embryos can be returned to liquid medium and subsequently imaged or fixed at later points in development. With an increasing abundance of transgenic zebrafish lines and well-characterized fate mapping and transplantation techniques, imaging any desired tissue is possible. As such, time-lapse in vivo imaging combines powerfully with zebrafish genetic methods, including analyses of mutant and microinjected embryos.

Time-lapse confocal imaging is a powerful technique useful for characterizing embryonic development. Here, we describe the methodology and characterize craniofacial morphogenesis in wild-type, as well as pdgfra, smad5, and smo mutant embryos.

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical ...

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model congenital abnormalities. During the first day of life, the zebrafish embryo progresses through many developmental stages including fertilization, cleavage, gastrulation, segmentation, and the organogenesis of structures such as the kidney, heart, and central nervous system. The anatomy of a young zebrafish embryo presents several challenges for the visualization and analysis of the tissues involved in many of these events because the embryo develops in association with a round yolk mass. Thus, for accurate analysis and imaging of experimental phenotypes in fixed embryonic specimens between the tailbud and 20 somite stage (10 and 19 hours post fertilization (hpf), respectively), such as those stained using whole mount in situ hybridization (WISH), it is often desirable to remove the embryo from the yolk ball and to position it flat on a glass slide. However, performing a flat mount procedure can be tedious. Therefore, successful and efficient flat mount preparation is greatly facilitated through the visual demonstration of the dissection technique, and also helped by using reagents that assist in optimal tissue handling. Here, we provide our WISH protocol for one or two-color detection of gene expression in the zebrafish embryo, and demonstrate how the flat mounting procedure can be performed on this example of a stained fixed specimen. This flat mounting protocol is broadly applicable to the study of many embryonic structures that emerge during early zebrafish development, and can be implemented in conjunction with other staining methods performed on fixed embryo samples.

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization

The zebrafish embryo is an excellent model for developmental biology research. During embryogenesis, zebrafish develop with a yolk mass, which presents three-dimensional challenges for sample observation and analysis. This protocol describes how to create two-dimensional flat mount preparations of whole mount in situ (WISH) stained zebrafish embryo specimens.

The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model ...

Dissection and Downstream Analysis of Zebra Finch Embryos at Early Stages of Development

Summary

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Chapters in this video

Analysis of Zebrafish Kidney Development with Time-lapse Imaging Using a Dissecting Microscope Equipped for Optical Sectioning

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Dissection of Larval Zebrafish Gonadal Tissue

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

Dissection and Lateral Mounting of Zebrafish Embryos: Analysis of Spinal Cord Development

A 3-D Tail Explant Culture to Study Vertebrate Segmentation in Zebrafish

Using Whole Mount in situ Hybridization to Link Molecular and Organismal Biology

Light Sheet Microscopy Imaging and Mounting Strategies for Early Zebrafish Embryos

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization