This work was supported by grant R00HL122360 from the National Institutes of Health, National Heart, Lung, and Blood Institute (NHLBI).

All methods described adhere to animal care guidelines of The University of North Carolina at Chapel Hill.
1. Preparation of micro-injection pipettes
<ol>
	<li>Pull glass capillaries using a micropipette puller. For some injections, needle beveling is recommended as it provides an extremely sharp surface devoid of structural impurities. To do this, polish the end of a pulled needle on a beveling wheel at an angle of 45&#176; for 15&#8722;20 min. 
	NOTE: Exact settings for pulling will vary based on the puller being used. The final inner diameter of the bevel should be between 20&#8722;40 &#181;m.</li>
	<li>Coat the inner and external surfaces of the glass capillary with silicone. First dip the needles in the siliconizing agent to coat the external surface of the needle and then backload the siliconizing agent solution into each micropipette to coat the inner surface. 
	NOTE: Coating of the glass capillaries should be done 24 h before the implantation experiment. Coating the glass capillaries with silicone provides a chemically inert surface to the glass. If the capillaries are left untreated, the cell suspension generated in later steps will adhere to the glass and plug the needle. Therefore, coating is necessary and vital to the success of the method. 
	CAUTION: The siliconizing agent is a ready-made commercially available mixture of heptane and 1,7-dichloro-1,1,3,3,5,5,7,7-octamethyltetrasiloxane (Table of Materials). It is extremely flammable and acutely toxic. Always handle with proper PPE inside a fume hood.
	<ol>
		<li>To backload the needle, load ~5&#8722;10 &#181;L of the siliconizing agent into a micro-injection pipette tip (Table of Materials). Place the micro-injection tip in the wide end of the pulled glass capillary and position the tip as far down as possible (close to the glass needle tip). Eject the siliconizing agent while slowly removing the loading pipette in order to minimize air bubbles in the needle.</li>
		<li>Leave the siliconizing agent in the glass needle for 10 min, remove by aspirating with a new loading pipette and allow needles dry overnight in a fume hood.</li>
	</ol>
	</li>
	<li>The morning of the experiment, rinse the glass capillaries with deionized water following the procedure in step 1.2 and allow to dry for 3&#8722;4 h.</li>
</ol>
2. Preparation of solutions
<ol>
	<li>Prepare 5 mL of trypsin neutralizing solution by supplementing 4.2 mL of Dulbecco&#39;s modified Eagle&#39;s medium and Ham&#39;s nutrient mixture F12 (DMEM/F12) with 750 &#181;L of Fetal Bovine Serum (FBS) and 50 &#181;L of Penicillin/Streptomycin. Store at 37 &#176;C until use.</li>
	<li>Prepare 5 &#181;M labeling dye solution by pipetting 5 &#181;L of 1 mM stock labeling dye (in dimethyl sulfoxide [DMSO], Table of Materials) into 995 &#181;L of Hank&#39;s balanced salt solution (HBSS). Vortex for 1 min and store at 37 &#176;C until use.</li>
	<li>Prepare fresh paraformaldehyde (PFA) by combining 10 mL of 32% PFA stock solution with 62 mL of molecular biology grade water and 8 mL of 10x Dulbecco&#39;s phosphate-buffered saline (DPBS). The final concentration is 4% PFA in 1x DPBS.</li>
</ol>
3. Preparation of host embryos
<ol>
	<li>Incubate fertile chicken eggs in a horizontal orientation in a humidified incubator at 38 &#176;C until Hamburger and Hamilton (HH) Stage 1910. 
	NOTE: The stage chosen for manipulation is flexible and entirely dependent upon the aims of each individual experiment.</li>
	<li>Score the &#34;flat&#34; end of the egg shell along the egg equator using angled forceps to make a small puncture &#62;1 mm in diameter. Insert an 18 G needle with attached 10 mL syringe through the puncture and remove ~5 mL of albumin. 
	NOTE: This anatomical location is external to the &#34;air cell&#34; inside the egg and keeps albumin from leaking once the puncture is made. This step is recommended as it &#34;drops&#34; the embryo away from the egg shell, preventing potential damage in the subsequent steps.</li>
	<li>Apply transparent tape to the top of the egg shell. Score with angled forceps and cut a ~2.5 cm window using curved tenotomy scissors.</li>
	<li>Inspect and stage the embryo based on criteria established by Hamburger and Hamilton10 and seal the puncture from step 3.2 with transparent tape. 
	NOTE: Here, stage HH 19 embryos are used which have 37&#8722;40 somites extending into the tail bud. The puncture should not be sealed until after the window is opened along the top of the egg (step 3.3).</li>
	<li>Inject ~200 &#181;L of India Ink/HBSS mixture (1:5) beneath the embryo using a 32 G needle with attached 1 mL syringe. 
	NOTE: India ink provides visual contrast between the embryo and the yolk beneath. Alternative dyes such as neutral red or commercially available cyan fluorescent protein (CFP) or blue fluorescent protein (BFP) fluorescence filter sets can be used to improve contrast.</li>
	<li>Add 1 mL of HBSS dropwise onto the embryonic disc and seal windowed shells with paraffin film. Place eggs back in the humidified incubator until ready for injection.</li>
</ol>
4. Isolation of donor tissue
<ol>
	<li>Incubate donor fertile chicken eggs in a humidified incubator at 38 &#176;C until Stage HH 19 (or desired stage).</li>
	<li>Remove the embryo from the egg and place in a 100 mm x 15 mm petri dish containing sterile HBSS at room temperature (RT).</li>
	<li>Surgically microdissect atrial donor tissue from each embryo by first isolating the entire embryonic heart from the embryo and then by isolating the atria from the heart using forceps, tenotomy scissors, and microspatula under a stereo dissecting microscope. Pool in a sterile 1.5 mL microcentrifuge tube containing 1 mL of HBSS on ice.</li>
	<li>Once all donor tissue has been collected, pellet the tissue by centrifugation at 1000 x g for 5 min at 4 &#176;C in a fixed-angle microcentrifuge.</li>
</ol>
5. Trypsin digestion of donor tissue
<ol>
	<li>Resuspend cell pellets in 1 mL of prewarmed 0.05% trypsin-EDTA and incubate at 37 &#176;C for 15 min in a shaking heat block at 300 rpm. Alternatively, use a water bath with periodical agitation of the sample.</li>
	<li>Pipette the digestion solution up and down to break up any remaining tissue, and pellet as in step 4.4.</li>
	<li>Resuspend the pellet in 1 mL of the trypsin neutralizing solution and centrifuge as in step 4.4.</li>
	<li>Resuspend the cells in 400 &#181;L of red fluorescent labeling dye solution and incubate at 37 &#176;C for 20 min in a heat block. Alternatively, use a water bath.</li>
	<li>Once the labeling reaction is finished, pellet the cells as in step 4.4 and wash with 1 mL of HBSS (number of wash steps can be varied between 1 and 3).</li>
	<li>Resuspend the labeled, pelleted cells at a concentration of ~50,000 cells/&#181;L, which generally results in a 5&#8722;10 &#181;L working volume depending on the total cell yield. 
	NOTE: Cell concentrations below ~50,000 cells/&#181;L can result in poor injection efficiency.</li>
</ol>
6. In-vivo injection
<ol>
	<li>Backload the cell suspension into a silicone treated glass capillary pipette following the procedures in step 1.2. Mount the pipette into the pressure microinjector apparatus.</li>
	<li>Remove host embryos from humidified incubator and place in an egg holder underneath the fluorescent stereo dissecting microscope.
	<ol>
		<li>Open the vitelline membrane using sterile fine forceps and make a small incision (~0.5&#8722;1.0 mm in length) in the pericardium. Additional manipulation/dissection may be needed depending on target region for injection.</li>
	</ol>
	</li>
	<li>Position the microinjector such that the tip of the microinjection needle penetrates the target tissue.
	<ol>
		<li>Pressure inject cells, and use the fluorescent label to determine that implanted cells are present in desired tissue. For typical injections, apply single pulses less than 0.5 s in duration ranging from 100-400 hectopascals in pressure. 
		NOTE: Pulse length and absolute pressure will vary depending on the number of cells to be injected and can be modified to suit individual needs.</li>
		<li>Retract the microinjector apparatus and remove the egg from the holder after pressure injecting.</li>
	</ol>
	</li>
	<li>Add 1 mL of warm HBSS dropwise onto the embryo, seal eggs using transparent packing tape, and incubate in the humidified incubator at 38 &#176;C for 24 h post implantation.</li>
</ol>
7. Isolation and analysis
<ol>
	<li>Isolate host embryos in RT HBSS using forceps, tenotomy scissors, and microspatula similar to step 4.3, and fix in 4% PFA overnight at 4 &#176;C with gentle rocking.</li>
	<li>Wash embryos 3 x 5 min in HBSS at RT with gentle rocking, and store in HBSS at 4 &#176;C for further downstream analysis (microscopy, immunohistochemistry, in-situ hybridization, etc.).</li>
</ol>

<ol>
	<li>Guo, Y. X., Pu, W. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+Mosaics+for+Greater+Precision+in+Cardiovascular+Research.">Genetic Mosaics for Greater Precision in Cardiovascular Research.</a> Circulation Research. 123, 27-29 (2018).</li><li>Guo, Y. X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+Cardiac+Myocyte+Maturation+Using+CASAAV,+a+Platform+for+Rapid+Dissection+of+Cardiac+Myocyte+Gene+Function+In+Vivo.">Analysis of Cardiac Myocyte Maturation Using CASAAV, a Platform for Rapid Dissection of Cardiac Myocyte Gene Function In Vivo.</a> Circulation Research. 120 (12), 1874-1888 (2017).</li><li>Rugh, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Experimental embryology; techniques and procedures. 3rd edition. , (1962).</li><li>Slack, J. M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Essential developmental biology. 3rd edition. , (2013).</li><li>Reinecke, H., Zhang, M., Bartosek, T., Murray, C. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Survival,+integration,+and+differentation+of+cardiomyocyte+grafts:+a+study+in+normal+and+injured+rat+hearts.">Survival, integration, and differentation of cardiomyocyte grafts: a study in normal and injured rat hearts.</a> Circulation. 100, 193-202 (1999).</li><li>Rojas, S. V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transplantation+of+purified+iPSC-derived+cardiomyocytes+in+myocardial+infarction.">Transplantation of purified iPSC-derived cardiomyocytes in myocardial infarction.</a> PLOS ONE. 12, 0173222 (2017).</li><li>Zhang, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiomyocyte+Grafting+for+Cardiac+Repair:+Graft+Cell+Death+and+Anti-Death+Strategies.">Cardiomyocyte Grafting for Cardiac Repair: Graft Cell Death and Anti-Death Strategies.</a> Journal of Molecular and Cellular Cardiology. 33 (5), 907-921 (2001).</li><li>Bressan, M., Liu, G., Mikawa, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+mesodermal+cues+assign+avian+cardiac+pacemaker+fate+potential+in+a+tertiary+heart+field.">Early mesodermal cues assign avian cardiac pacemaker fate potential in a tertiary heart field.</a> Science. 340 (6133), 744-748 (2013).</li><li>Bressan, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reciprocal+myocardial-endocardial+interactions+pattern+the+delay+in+atrioventricular+junction+conduction.">Reciprocal myocardial-endocardial interactions pattern the delay in atrioventricular junction conduction.</a> Development. 141 (21), 4149-4157 (2014).</li><li>Hamburger, V., Hamilton, H. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+series+of+normal+stages+in+the+development+of+the+chick+embryo.">A series of normal stages in the development of the chick embryo.</a> Journal of Morphology. 88 (1), 49-92 (1951).</li></ol>

The authors have nothing to disclose.

The ability to define how microenvironmental conditions impact cardiac cell fate specification and lineage stabilization is fundamental to creating a compressive understanding of congenital heart disease as well as to developing efficient protocols for proper maturation of stem cell or somatic cell reprograming-based cardiomyocytes. The protocol outlined above gives investigators the ability to directly assay cardiac cell development under altered in vivo conditions, allowing for cell autonomous maturation processes to be separated from paracrine/juxtacrine and/or hemodynamic cues. When combined with high resolution imaging, genetic analysis, and physiological assays, this technique can serve as a powerful complement to existing transgenic models.
Form a technical stand point, the protocol presented here relies on efficient isolation, labeling, and precise implantation of donor heart cells into host embryonic tissues. The use of a microinjection system greatly aids in the targeting of the donor cells and allows for successful implantation without the need for creating a large engraftment site in the host tissue. Some operational skill is required to perform this technique however, as reduced viability can result if the injection needle is not carefully placed in the target tissue (causing rupture of the heart or local vasculature). Care and thought should also be given to the isolation and labeling steps. Over digestion of the donor tissue can lead to poor implantation efficiency, and transient labeling techniques can limit the time window over which donor cells can be tracked (as cell division can dilute the label).
This technique is highly modifiable and can be adapted for a variety of purposes. For example, donor cells from a large range of tissues and stages can be isolated (though optimization of the enzymatic digestion is required) and can similarly be injected into a variety of host tissues across different stages of development. Similarly, the labeling approach can be modified to track cells across different temporal windows, including the use of fluorescent inorganic semiconductor nanocrystals for longer transient labeling and implantation of quail cells or cells from green fluorescent protein transgenic (GFP+) donor embryos11 for permeant labeling.
While we currently use this technique for avian implantation studies, we feel that it could be used for a large range of chimeric studies in the future. For example, genetically altered cardiac cells from transgenic organisms could be isolated and microinjected into the avian heart using a very similar protocol. Furthermore, cells differentiated into cardiomyocytes from stems cells or via somatic cell reprograming approaches could be microinjected into the embryonic heart to evaluate their integration into the tissue and/or maturation under in vivo biomechanical conditions.

Cardiac developmental research has benefitted enormously from the advent of germline transgenic model systems which have identified many of the gene regulatory networks that pattern different cell lineages and functional domains in the heart. However, identifying how these gene networks interact with and respond to microenvironmental conditions, including paracrine/juxtacrine signals and biophysical inputs (stretch, strain, hemodynamic flow), can be challenging. As such, it is not always easy to determine whether a cellular phenotype arises as a direct consequence of a genetic perturbation or as a secondary result of an adaption to changes in cardiac biomechanics or higher order tissue composition1,2.
Grafting experiments, which have classically been used to address concepts of fate specification, commitment, induction, and competence3,4, would represent an ideal approach to circumvent some of the challenges inherent in defining cell autonomous versus environmental influence in the heart. Unfortunately, heart contractions make standard grafting approaches difficult. Rapid movement of the tissue often prevents grafted cells from adhering to the heart and large tissue punctures (normally required for grafting) frequently lead to heart failure and embryonic lethality5,6,7. Therefore, we have developed a pressure-based, microinjection system for precision cellular implantation into the developing chick heart, circumventing the technical hurdles of tissue grafting described previously8,9. Using this technique, individual or small groups of cardiac cells isolated from a donor embryo can be microinjected into a variety of regions of a host embryonic heart eliminating the need for extensive host preparation and the large tissue insults that arise using standard grafting techniques. The microinjection needles used for these implantation studies have an outer diameter of ~30&#8722;40 &#181;m, which means that the needle can be placed directly in the target tissue (i.e., can penetrate the embryonic myocardial wall) and cells can be focally delivered with minimal damage to the surrounding tissue. The protocol can be used to perform a variety of isotopic, heterotopic, isochoric, and heterochronic manipulations, providing a rapid, flexible, and low-cost approach to directly examine classical experimental embryological paradigms in the developing four-chambered heart.
In the protocol outlined below, we label donor cells with a cell permeant fluorescent dye, which allows for the success of a microinjection experiment to be monitored in real-time and the location of engrafted cells to be documented without the need for any additional staining. However, it should be noted that this approach is best suited for short term experiments (approximately 48 h) as the fluorescent dye can be lost through cell division. Alternative approaches can be used for longer term experiments.
While we are presenting this technique in the context of cardiac development, we have used it to great effect for cell implantation experiments into the mesoderm, head, limbs, and somites. As such the basic approach described below is highly tractable and can be used in a variety of organ systems.

<table><tbody><tr><td>1 mL Insulin Syringe</td><td>BD</td><td>329654</td><td></td></tr><tr><td>1.7 mL Microtubes, Clear</td><td>Genesee Scientific</td><td>24-282</td><td></td></tr><tr><td>10 ml Syringe</td><td>BD</td><td>305482</td><td></td></tr><tr><td>1000ul Reach Barrier Tip Racked, Sterile</td><td>Genesee Scientific</td><td>24-430</td><td></td></tr><tr><td>15 mL Centriguge Tubes, Racked</td><td>Genesee Scientific</td><td>28-101</td><td></td></tr><tr><td>1588 Genesis Hova-Bator Incubator</td><td>GQF</td><td>813927021221</td><td></td></tr><tr><td>18G x 1 1/2 Needle</td><td>BD</td><td>305196</td><td></td></tr><tr><td>200ul Barrier Tip Low Binding, Racked, Sterile</td><td>Genesee Scientific</td><td>24-412</td><td></td></tr><tr><td>32G x 1/2&quot; Needle</td><td>TSK Steriject Air-Tite</td><td>TSK3213</td><td></td></tr><tr><td>Alchohol Wipes 70%</td><td>Thermo Fisher Scientific</td><td>19015744</td><td></td></tr><tr><td>Angled Forceps</td><td>Fine Scientific Tools</td><td>11260-20</td><td></td></tr><tr><td>Backloading Tips</td><td>Eppendorf</td><td>930001007</td><td></td></tr><tr><td>Black India Ink</td><td>KOH-I-NOOR</td><td>3084-F</td><td></td></tr><tr><td>CellTracker Green CMF</td><td>Thermo Fisher Scientific</td><td>C7025</td><td>1 mM in DMSO</td></tr><tr><td>CellTracker Red CMTPX</td><td>Thermo Fisher Scientific</td><td>C34552</td><td>1 mM in DMSO</td></tr><tr><td>Centrifuge</td><td>Eppendorf</td><td>5424R</td><td></td></tr><tr><td>Commercial Grade Packing Tape</td><td>Staples</td><td>2619001</td><td></td></tr><tr><td>Curved Tenotomy Scissors</td><td>Fine Scientific Tools</td><td>14067-11</td><td></td></tr><tr><td>DMEM/F12</td><td>Thermo Fisher Scientific</td><td>11330-032</td><td></td></tr><tr><td>DMSO, anhydrous</td><td>Thermo Fisher Scientific</td><td>D12345</td><td></td></tr><tr><td>DPBS (10X), no calcium, no magnesium</td><td>Thermo Fisher Scientific</td><td>14025092</td><td></td></tr><tr><td>Femtojet 4i</td><td>Eppendorf</td><td>5252000021</td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>Thermo Fisher Scientific</td><td>10437-028</td><td></td></tr><tr><td>Hatching Eggs</td><td>Pilgrim&#039;s Hatchery</td><td>--</td><td></td></tr><tr><td>HBSS, calcium, magnesium, no phenol red</td><td>Thermo Fisher Scientific</td><td>14025-092</td><td></td></tr><tr><td>Injectman 4</td><td>Eppendorf</td><td>5192000027D</td><td></td></tr><tr><td>Micromanipulator</td><td>Leica Microsystems&amp;nbsp;</td><td>--</td><td></td></tr><tr><td>Parafilm</td><td>SIGMA</td><td>P6543-1EA</td><td></td></tr><tr><td>Paraformaldehyde 32% in aqueous solution, EM Grade</td><td>VWR</td><td>100496-496</td><td></td></tr><tr><td>Penicillin/Streptomycin</td><td>Thermo Fisher Scientific</td><td>15140-022</td><td></td></tr><tr><td>Petri Dish</td><td>Genesee Scientific</td><td>32-107</td><td></td></tr><tr><td>Pipette Grinder</td><td>Narishige</td><td>EG-44</td><td></td></tr><tr><td>Pipette Puller</td><td>HEKA</td><td>PIP 6</td><td></td></tr><tr><td>Scotch Transparent Tape</td><td>Staples</td><td>487909</td><td></td></tr><tr><td>Sigmacote</td><td>SIGMA</td><td>SL2-25ML</td><td></td></tr><tr><td>Stereo Microscope</td><td>Leica</td><td>--</td><td></td></tr><tr><td>ThermoMixer C</td><td>Eppendorf</td><td>5382000023</td><td></td></tr><tr><td>Thin Wall Glass Capillaries</td><td>World Precision Instruments</td><td>TW100F-4&amp;nbsp;</td><td></td></tr><tr><td>Transfer Pipette</td><td>Thermo Fisher Scientific</td><td>273</td><td></td></tr><tr><td>Trypsin-EDTA (0.05%), phenol red</td><td>Thermo Fisher Scientific</td><td>25300-054</td><td></td></tr></tbody></table>

microinjection-based system for in vivo implantation of embryonic cardiomyocytes in the avian embryo

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

After 24 h incubation, the heart and surround tissue of host embryos were isolated, photographed (Figure 1A), and processed for immunofluorescent analysis. In this example, donor atrial myocytes were microinjected into the proepicardium of a similar staged host embryo. The host embryo was then stained with the muscle marker (MF20 green) and 4&#39;,6-diamidino-2-phenylindole (DAPI; blue). Injected cells (red) are clearly visible (Figure 1B). Possible adjustments to consider if cells are not visible include: donor tissue was over digested (cells would be unable to attach), labeling dye solution was too dilute, cells were over-washed, or multiple cell divisions resulted in loss of the label.
To confirm that the injected cells in this example were myocardial, we optically sectioned this embryo using a confocal microscope (Figure 1C-E). The only MF20 positive cells within the proepicardium (PE) are the fluorescent red positive cells that were focally implanted.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59267/59267fig1.jpg" /> 
Figure 1: Representative images of embryos isolated 24 hours post injection. (A) Low magnification brightfield image of the trunk region of an E3.5 (HH Stage 19) chick embryo. (B) Merged image showing injected cells (red), cardiomyocytes (green), and DAPI. Cells were isolated from the atria and microinjected into the proepicardium. (C) High magnification confocal imaging showing labeled cells in the core of the proepicardium. (D) High magnification confocal imaging confirming CT Red labeled cells are cardiomyocytes. (E) Three-dimensional (3D) reconstruction of injected cells from panels D and E. At, atria; OFT, outflow tract; PE, proepicardium; Vt, ventricle; MF20, Myosin 4. <a href="https://www.jove.com/files/ftp_upload/59267/59267fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Microinjection-based System for In Vivo Implantation of Embryonic Cardiomyocytes in the Avian Embryo at JoVE.com

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

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

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

microinjection-based-system-for-vivo-implantation-embryonic

Research

JoVE Journal

Developmental Biology

6.2K Views.  University of North Carolina at Chapel Hill. In this method, embryonic cardiac tissues are surgically microdissected, dissociated, fluorescently labeled, and implanted into host embryonic tissues. This provides a platform for studying the individual or tissue level developmental organization under ectopic hemodynamic conditions, and/or altered paracrine/juxtacrine environments.

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

Intramyocardial Cell Delivery: Observations in Murine Hearts

Previous studies showed that cell delivery promotes cardiac function amelioration by release of cytokines and factors that increase cardiac tissue revascularization and cell survival. In addition, further observations revealed that specific stem cells, such as cardiac stem cells, mesenchymal stem cells and cardiospheres have the ability to integrate within the surrounding myocardium by differentiating into cardiomyocytes, smooth muscle cells and endothelial cells.
Here, we present the materials and methods to reliably deliver noncontractile cells into the left ventricular wall of immunodepleted mice. The salient steps of this microsurgical procedure involve anesthesia and analgesia injection, intratracheal intubation, incision to open the chest and expose the heart and delivery of cells by a sterile 30-gauge needle and a precision microliter syringe.
Tissue processing consisting of heart harvesting, embedding, sectioning and histological staining showed that intramyocardial cell injection produced a small damage in the epicardial area, as well as in the ventricular wall. Noncontractile cells were retained into the myocardial wall of immunocompromised mice and were surrounded by a layer of fibrotic tissue, likely to protect from cardiac pressure and mechanical load.

Intramyocardial cell delivery in murine models of cardiovascular diseases, such as hypertension or myocardial infarction, is widely used to test the therapeutic potential of different cell types in regenerative studies. Therefore, a detailed description and a clear visualization of this surgical procedure will help to define the limits and advantages of cardiovascular cell therapeutic analyses in small rodents.

Previous studies showed that cell delivery promotes cardiac function amelioration by release of cytokines and factors that increase cardiac tissue ...

Medicine

Microinjection of Medaka Embryos for use as a Model Genetic Organism

In this video, we demonstrate the technique of microinjection into one-cell stage medaka embryos. Medaka is a small egg-laying freshwater fish that allows both genetic and embryological analyses and is one of the vertebrate model organisms in which genome-wide phenotype-driven mutant screens were carried out 1, as in zebrafish and the mouse. Divergence of functional overlap of related genes between medaka and zebrafish allows identification of novel phenotypes that are unidentifiable in a single species 2, thus medaka and zebrafish are complementary for genetic dissection of vertebrate genome functions.
To take advantage of medaka fish whose embryos are transparent and develop externally, microinjection is an essential technique to inject cell-tracers for labeling cells, mRNAs or anti-sense oligonucleotides for over-expressing and knocking-down genes of interest, and DNAs for making transgenic lines.

Medaka and zebrafish are complementary for genetic dissection of vertebrate genome functions. This protocol highlights the key points for successful microinjection into medaka embryos, an important technique for embryological and genetic analysis using medaka and zebrafish in a laboratory.

In this video, we demonstrate the technique of microinjection into one-cell stage medaka embryos. Medaka is a small egg-laying freshwater fish that allows ...

Biology

Single Cell Transfection in Chick Embryos

A central theme in developmental biology is the diversification of lineages and the elucidation of underlying molecular mechanisms. This entails a thorough analysis of the fates of single cells under normal and experimental conditions. To this end, transfection methods that target single progenitors are a prerequisite. We describe here a technically straightforward method for transfecting single cells in chicken tissues in-ovo, allowing reliable lineage tracing as well as genetic manipulation. Specific tissue domains are targeted within the somite or neural tube, and DNA is injected directly into the epithelium of interest, resulting in sporadic transfection of single cells. Using reporters, clonal populations may consequently be traced for up to three days, and behavior of genetically manipulated clonal populations can be compared with that of controls. This method takes advantage of the accessibility of the chick embryo along with emerging tools for genetic manipulation. We compare and discuss its advantages over the widely-used electroporation method, and possible applications and use in additional in-vivo models are also suggested. We advocate the use of this method as a significant addition and complement for existing lineage tracing and genetic interference tools.

Using fine tip micropipettes we inject plasmid DNA into subdomains of chicken somites or neural tubes. The concentration of the plasmid is adjusted to generate single transfected cells. We then allow the cells to develop into clonal populations.

A central theme in developmental biology is the diversification of lineages and the elucidation of underlying molecular mechanisms. This entails a ...

An ex-ovo Chicken Embryo Culture System Suitable for Imaging and Microsurgery Applications

Understanding the relationships between genetic and microenvironmental factors that drive normal and malformed embryonic development is fundamental for discovering new therapeutic strategies. Advancements in imaging technology have enabled quantitative investigation of the organization and maturing of the body plan, but later stage embryonic morphogenesis is less clear. Chicken embryos are an attractive vertebrate animal model system for this application because of its ease of culture and surgical manipulation. Early embryos can be cultured for a short time on filter paper rings, which enables complete optical access for cell patterning and fate studies1,2. Studying advanced developmental processes such as cardiac morphogenesis are traditionally performed through a window of the eggshell3-5, but this technique limits optical access due to window size. We previously developed a simple method to culture whole embryos ex-ovo on hexagonal weigh boats for up to 10 days, which enabled high resolution imaging via ultrasonography6,7. These cultures were difficult to transport, limiting the types of imaging tools available for live experiments. We here present an improved shell-less culture system with a cost-effective, portable environmental chamber. Eggs were cracked onto a hammock created by a polyurethane membrane (cling wrap) affixed circumferentially to a plastic cup partially filled with sterile water. The dimensions of the circumference and depth of the hammock were both critical to maintain surface tension, while the mechanics of the hammock and water beneath helped dampen vibrations induced by transportation. A small footprint circulating water bath was also developed to enable continuous temperature control during experimentation. We demonstrate the ability to culture embryos in this way for at least 14 days without morphogenic defect or delay and employ this system in several microsurgical and imaging applications.

An ex-ovo Chicken Embryo Culture System Suitable for Imaging and Microsurgery Applications

In this article, we present a simple methodology to enable long-term ex-ovo avian embryo culture. This technique is ideal for longitudinal experimentation requiring complete optical accessibility and/or sterile transportation in avian embryos.

Understanding the relationships between genetic and microenvironmental factors that drive normal and malformed embryonic development is fundamental for ...

Avian Embryonic Left Atrial Ligation Model for HLHS and Cardiovascular Research

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

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

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

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

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

Percutaneous Contrast Echocardiography-guided Intramyocardial Injection and Cell Delivery in a Large Preclinical Model

Cell and gene therapy are exciting and promising strategies for the purpose of cardiac regeneration in the setting of heart failure with reduced ejection fraction (HFrEF). Before they can be considered for use, and implemented in humans, extensive preclinical studies are required in large animal models to evaluate the safety, efficacy, and fate of the injectate (e.g., stem cells) once delivered into the myocardium. Small rodent models offer advantages (e.g., cost effectiveness, amenability for genetic manipulation); however, given inherent limitations of these models, the findings in these rarely translate into the clinic. Conversely, large animal models such as rabbits, have advantages (e.g., similar cardiac electrophysiology compared to humans and other large animals), whilst retaining a good cost-effective balance. Here, we demonstrate how to perform a percutaneous contrast echocardiography-guided intramyocardial injection (IMI) technique, which is minimally invasive, safe, well tolerated, and very effective in the targeted delivery of injectates, including cells, into several locations within the myocardium of a rabbit model. For the implementation of this technique, we also have taken advantage of a widely available clinical echocardiography system. After putting in practice the protocol described here, a researcher with basic ultrasound knowledge will become competent in the performance of this versatile and minimally invasive technique for routine use in experiments, aimed at hypothesis testing of the capabilities of cardiac regenerative therapeutics in the rabbit model. Once competency is achieved, the whole procedure can be performed within 25 min after anaesthetizing the rabbit.

Novel therapeutic strategies in cardiac regenerative medicine require extensive and detailed studies in large preclinical animal models before they can be considered for use in humans. Here, we demonstrate a percutaneous contrast echocardiography-guided intramyocardial injection technique in rabbits, which is valuable for hypothesis testing the efficacy of such novel therapies.

Cell and gene therapy are exciting and promising strategies for the purpose of cardiac regeneration in the setting of heart failure with reduced ejection ...

A Novel Minimally Invasive Slow Injection Method for Reducing Tissue Injury

A Novel Cell Injection Method with Minimum Invasion

Directly injecting cells into tissues is a necessary process in cell administration and/or replacement therapy. The cell injection requires a sufficient amount of suspension solution to allow the cells to enter the tissue. The volume of the suspension solution affects the tissue, and this can cause major invasive injury as a result of the cell injection. This paper reports on a novel cell injection method, called slow injection, that aims to avoid this injury. However, pushing out the cells from the needle tip requires a sufficiently high injection speed according to Newton's law of shear force. To solve the above contradiction, a non-Newtonian fluid, such as gelatin solution, was used as the cell suspension solution in this work. Gelatins solution have temperature sensitivity, as their form changes from gel to sol at approximately 20 &#176;C. Therefore, to maintain the cell suspension solution in the gel form, the syringe was kept cooled in this protocol; however, once the solution was injected into the body, the body temperature converted it to a sol. The interstitial tissue fluid flow can absorb excess solution. In this work, the slow injection technique allowed cardiomyocyte balls to enter the host myocardium and engraft without surrounding fibrosis. This study employed a slow injection method to inject purified and ball-formed neonatal rat cardiomyocytes into a remote area of myocardial infarction in the adult rat heart. At 2 months following the injection, the hearts of the transplanted groups showed significantly improved contractile function. Furthermore, histological analyses of the slow-injected hearts revealed seamless connections between the host and graft cardiomyocytes via intercalated disks containing gap junction connections. This method could contribute to next-generation cell therapies, particularly in cardiac regenerative medicine.

Author Spotlight: A Novel Cell Injection Method with Minimum Invasion  

This method eliminates any major invasion during cell injections caused by the cell suspension solution.

Directly injecting cells into tissues is a necessary process in cell administration and/or replacement therapy. The cell injection requires a sufficient ...

Bioengineering

Cell Labeling and Injection in Developing Embryonic Mouse Hearts

Testing the fate of embryonic or pluripotent stem cell-derivatives in in vitro protocols has led to controversial outcomes that do not necessarily reflect their in vivo potential. Preferably, these cells should be placed in a proper embryonic environment in order to acquire their definite phenotype. Furthermore, cell lineage tracing studies in the mouse after labeling cells with dyes or retroviral vectors has remained mostly limited to early stage mouse embryos with still poorly developed organs. To overcome these limitations, we designed standard and ultrasound-mediated microinjection protocols to inject various agents in targeted regions of the heart in mouse embryos at E9.5 and later stages of development. &#160;Embryonic explant or embryos are then cultured or left to further develop in utero. These agents include fluorescent dyes, virus, shRNAs, or stem cell-derived progenitor cells. Our approaches allow for preservation of the function of the organ while monitoring migration and fate of labeled and/or injected cells. These technologies can be extended to other organs and will be very helpful to address key biological questions in biology of development.

We describe a series of methods to inject dyes, DNA vectors, virus, and cells in order to monitor both cell fate and phenotype of endogenous and grafted cells derived from embryonic or pluripotent cells within mouse embryos at embryonic day (E)9.5 and later stages of development.

Testing the fate of embryonic or pluripotent stem cell-derivatives in in vitro protocols has led to controversial outcomes that do not necessarily reflect ...

Loss-of-Function Approach in the Embryonic Chick Retina by Using Tol2 Transposon-Mediated Transgenic Expression of Artificial microRNAs

The chick retina has long been an important model system in developmental neurobiology, with advantages including its large size, rapid development, and accessibility for visualization and experimental manipulations. However, its major technical limitation had been the lack of robust loss-of-function approaches for gene function analyses. This protocol describes a methodology of gene silencing in the developing chick retina that involves transgenic expression of artificial microRNAs (miRNAs) by using the Tol2 transposon system. In this approach, a Tol2 transposon plasmid that contains an expression cassette for the EmGFP (emerald green fluorescent protein) marker and artificial pre-miRNA sequences against a target gene is introduced into the embryonic chick retina with a Tol2 transposase expression construct by in ovo electroporation. In the transfected retinal cells, the transposase catalyzes the excision of the expression cassette from the transposon vector and its integration into host chromosomes, leading to the stable expression of miRNAs and the EmGFP protein. In our previous study, we have demonstrated that the expression of Nel, a glycoprotein that exerts multiple functions in neural development, can be significantly suppressed in the developing chick retina by using this technique. Our results indicate that this methodology induces a stable and robust suppression of gene expression and thus provides an efficient loss-of-function approach for studies of retinal development.

We have developed a novel loss-of-function approach that involves the introduction and genomic integration of artificial micro-RNA sequences into chick embryos by using in ovo electroporation and the Tol2 transposon system. This technique provides a robust and stable gene knockdown methodology for studies of gene function during development.

The chick retina has long been an important model system in developmental neurobiology, with advantages including its large size, rapid development, and ...

Neuroscience

Ultrasound-Guided Induced Pluripotent Stem Cell-Derived Cardiomyocyte Implantation in Myocardial Infarcted Mice

The key objective of cell therapy after myocardial infarction (MI) is to effectively enhance the cell grafted rate, and human induced pluripotent stem cell-derived cardiomyocytes (hiPSC-CMs) are a promising cell source for cardiac repair after ischemic damage. However, a low grafted rate is a significant obstacle for effective cardiac tissue regeneration after transplantation. This protocol shows that multiple hiPSC-CM ultrasound-guided percutaneous injections into an MI area effectively increase cell transplantation rates. The study also describes the entire hiPSC-CM culture process, pretreatment, and ultrasound-guided percutaneous delivery methods. In addition, the use of human mitochondrial DNA help detect the absence of hiPSC-CMs in other mouse organs. Lastly, this paper describes the changes in cardiac function, angiogenesis, cell size, and apoptosis at the infarcted border zone in mice 4 weeks after cell delivery. It can be concluded that echocardiography-guided percutaneous injection of the left ventricular myocardium is a feasible, relatively invasive, satisfactory, repeatable, and effective cellular therapy.

Ultrasound-guided cell delivery around the site of myocardial infarction in mice is a safe, effective, and convenient way of cell transplantation.

The key objective of cell therapy after myocardial infarction (MI) is to effectively enhance the cell grafted rate, and human induced pluripotent stem ...

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

Summary

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Intramyocardial Cell Delivery: Observations in Murine Hearts

Microinjection of Medaka Embryos for use as a Model Genetic Organism

Single Cell Transfection in Chick Embryos

An ex-ovo Chicken Embryo Culture System Suitable for Imaging and Microsurgery Applications

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

Percutaneous Contrast Echocardiography-guided Intramyocardial Injection and Cell Delivery in a Large Preclinical Model

A Novel Cell Injection Method with Minimum Invasion

Cell Labeling and Injection in Developing Embryonic Mouse Hearts

Loss-of-Function Approach in the Embryonic Chick Retina by Using Tol2 Transposon-Mediated Transgenic Expression of Artificial microRNAs

Ultrasound-Guided Induced Pluripotent Stem Cell-Derived Cardiomyocyte Implantation in Myocardial Infarcted Mice