This work was funded by a National Institute of Dental and Craniofacial Research (NIDCR) F32 grant (DE021929) to J.L.F. and a NIDCR R01 grant DE016402 to R.A.S.

1. Prepare Tungsten Needles
<ol>
	<li>Cut a tungsten rod in half using wire cutters so that the rod does not bend.</li>
	<li>Thread the rod through the tapered end of a 5&#160;3/4 in&#160;borosilicate glass Pasteur pipette.</li>
	<li>Leaving approximately 3/4 in&#160;of the rod sticking out of the glass, adhere the rod to the pipette tip with a small bead of &#8220;hot melt adhesive&#8221; (HMA), which is the glue stick used for a glue gun. The HMA should be quickly melted in an alcohol flame, taking care not to burn the glue and generate black smoke.</li>
	<li>Using a pair of forceps, bend the tip of the tungsten rod (about 1/4 in&#160;from the top end) until a 45&#176;&#160;angle is reached.</li>
	<li>Holding the Pasteur pipette, place about 1/8 in&#160;of the tip of the tungsten rod into the flame of a propane torch. Hold the tip steady and exactly perpendicular to the flame.</li>
	<li>Pull the needle out of the flame the moment a tiny orange speck of tungsten flies off of the needle.</li>
</ol>
2. Prepare Spemann Pipettes
<ol>
	<li>Using a Bunsen burner, heat the narrow end of a Pasteur pipette until the glass shortly beyond the taper begins to melt. Remove from flame and pull the tip until the pipette becomes sealed. Be sure that there remains a portion of the tapered part that has an O.D. of approximately 0.7-1.0 mm. Break the sealed end off about 4 cm from the taper.</li>
	<li>Attach around 2 ft&#160;(60 cm) of rubber tubing to the wide (open) end of the pipette. Blow on the other end of the rubber tubing to ensure that no air can pass through the tapered part of the Pasteur pipette.</li>
	<li>Using a propane fuel cylinder with an attached pencil flame torch, heat an oval area on one side of the pipette just below the start of the taper. Blow air very gently and constantly through the rubber tubing with a slight amount of pressure (approximately the amount of pressure needed to say the beginning of the letter &#8220;P&#8221; at conversation level). When the glass becomes soft on one side and begins to buckle outwards, quickly remove the pipette from the flame and blow hard and steady through the rubber tubing. This will cause the heated part of the glass to form a sausage-shaped bubble that may pop, which is fine. If the bubble is too small, try melting and blowing the glass again. The final open window in the glass should be around 0.25 in&#160;(6.35 mm) wide by 0.5 in&#160;(12.7 mm) long.</li>
	<li>Pointing the tapered end upwards, scrape the bubble into a glass waste container while gently blowing through the tubing to prevent glass fragments from entering the pipette.</li>
	<li>Using the propane flame, burn off the remaining edges of the bubble and fire-polish the sides of the open window. Take care not to melt the shaft of the pipette.</li>
	<li>Using a diamond point pencil, score the tapered end of the pipette approximately 1.25 in&#160;(30 mm) from the window and break off the tip with forceps so that the pipette opening is cut cleanly (discard pipettes with broken or uneven tips).</li>
	<li>Using forceps and the edge of a flame from an alcohol burner, bend the narrow portion of the pipette 0.375 in&#160;(9.5 mm) from the tip to an angle of approximately 60&#176; so that the bent portion is in a vertical plane when the window opening is at the 1:00 position for right-handed usage and 11:00 for left-handed usage. This step is best conducted in a draft-free enclosure.</li>
	<li>Fire-polish the tip using the alcohol flame under a dissecting microscope. Be sure not to close off the tip but rather keep the opening round and smooth without any constriction.</li>
	<li>Cut a 1&#160;in&#160;(25 mm) piece of rubber tubing, spray the piece of tubing with 70% ethanol, and slide the tubing over the shaft of the pipette until the window opening is completely covered. This functions as the &#8220;diaphragm&#8221; of the Spemann pipette.</li>
	<li>Place a latex rubber 2 ml bulb over the open bottom of the pipette. The bulb maintains negative pressure in the Spemann pipette.</li>
	<li>To practice using a Spemann pipette, transfer chromatography beads of around 100-200 &#181;m in diameter from a marked spot in one Petri dish to a marked spot in another under a microscope.</li>
	<li>To clean the Spemann pipette, remove the rubber bulb and place pipette, tip down, in a tall beaker lined with cotton or gauze on the bottom and filled with distilled water and glassware detergent.</li>
</ol>
3. Sterilize and Incubate Eggs
<ol>
	<li>Wipe eggs with 70% ethanol and place the eggs on plastic egg trays.</li>
	<li>Set eggs in a heated incubator at 37.5 &#176;C and 85-87%.</li>
	<li>Incubate eggs until they reach HH9.5, approximately 26 hr for quail and 48 hr for duck.</li>
</ol>
4. Window Eggs
<ol>
	<li>Remove a small piece of outer shell from the top of the egg with forceps, taking care not to puncture the inner shell membrane.</li>
	<li>Using a syringe with an 18 G by 1-inch needle, poke a hole at the narrowest tip of the egg and withdraw approximately 1-2 ml of albumen.</li>
	<li>Place transparent tape over the hole and puncture mark in the shell.</li>
	<li>Cut a &#8220;window&#8221; along the top surface of the egg (through the transparent tape) using curved scissors.</li>
</ol>
5. Visualize Embryos and Prepare for Surgery
<ol>
	<li>Using a glass rod, gently brush a small amount of Neutral Red (0.02 g/ml in Hank&#8217;s BSS) over the embryo.</li>
	<li>Cut the vitelline membrane and pull it over the embryo using a flame-sharpened tungsten needle.</li>
	<li>Re-seal the egg by placing transparent tape over the window.</li>
</ol>
6. Separate the Donor Tissue from the Donor Embryo
<ol>
	<li>Remove tape from the window on the egg of the donor embryo.</li>
	<li>To cut the neural fold from the adjacent ectoderm of the donor embryo, make slits on both sides of the neural tube, using a flame-sharpened tungsten needle (manufactured as described above).</li>
	<li>Then, cut across the neural tube at the desired anterior and posterior levels of the graft and separate the graft from the rest of the neural tube.</li>
	<li>Remove the neural fold from the donor embryo using either a Spemann pipette or other type of micropipette.</li>
	<li>Then, remove the tape from the window on the host egg and use the Spemann pipette to place the donor neural fold alongside the host embryo adjacent to the region of the neural tube that will receive the transplant.</li>
</ol>
7. Separate the Host Tissue and Transplant the Donor Tissue
<ol>
	<li>Separate the neural fold from the neural tube of the host embryo, as done with the donor. Take care to remove a graft of equal size as the donor tissue.&#160;</li>
	<li>Gently push this host tissue far away from the embryo.</li>
	<li>Then, with a blunt tungsten needle or the rounded tip of a micropipette, gently move the donor neural fold into the host neural tube, maintaining the proper antero-posterior and dorso-ventral orientations. Make sure the graft is tucked in along the sides but be sure not to poke or damage to the underlying tissues.&#160;To help keep track of the original orientation note any asymmetry in the donor tissue or differential staining from the Neutral Red. We also photo-document the donor embryo with the accompanying excised tissue, as well as the chimera immediately following surgery.</li>
	<li>Sterile saline should be added to the egg if the host embryo shows signs of desiccation.</li>
	<li>Now, carefully re-seal and label the egg, and gently return it to a high humidity incubator (70-80%) where the chimeric embryo can develop to the desired stage for analysis.</li>
</ol>
8. Collection of Chimeras
<ol>
	<li>Be sure to collect chimeric embryos in freshly made&#160;cold Serra&#8217;s fixative and immediately place them on a rocker at 4 &#176;C for an O/N fixation. This will allow for more&#160;sensitive detection of quail cells with the anti-quail antibody.</li>
	<li>For gene expression analyses via RT-qPCR, freeze embryos directly in liquid nitrogen prior to RNA extraction.</li>
</ol>

The authors have nothing to disclose.

The neural crest is a transient embryonic cell population that migrates extensively throughout the embryo and differentiates into diverse cell types, including chondrocytes and osteoblasts, that contribute to the craniofacial skeleton. Transplanting neural crest in the quail-duck chimeric system has contributed greatly to our understanding of the tissue interactions and signaling pathways that regulate development of the craniofacial skeleton. However, given the vast potential of neural crest to also generate smooth muscle cells, adipocytes, melanocytes, schwann cells, and neurons, the quail-duck chimera system has tremendous potential for future applications, particularly in conjunction with the rapid advancement of stem cell biology and regenerative medicine. Since quail and duck are both commercially bred species, a ready supply of relatively inexpensive fertilized eggs is available from a variety of farms. Thus, this technique should be accessible to researchers operating within a wide range of budgets and facility space.
Although this technique is very powerful, there remain several limitations. Like other surgical techniques, the quality and viability of quail-duck chimeras rely on the surgical skills of the researcher, and therefore, there will be more inter- and intra-individual variation between experiments as compared to other models, such as those utilizing mouse genetics. Moreover, there is also variation in the rates of development and stages of individual embryos that contributes to the reproducibility and success of each transplant. Avian embryos are also very susceptible to dehydration and therefore critical steps during surgery include keeping the light levels low, the time under the microscope to a minimum, the eggs sealed with tape as much as possible, and high humidity in the post-operative incubator to avoid desiccation.&#160;
In terms of viability of the chimeras, usually between 50-75% survive, although these percentages can decrease the older the collection stage.&#160;In a typical 4-6 hr session of surgery, an experienced surgeon can generate 10-15 chimeras.&#160;The success of the transplants also depends greatly on the quality of the tools. Good tools lead to more consistent, reproducible results. Using a propane torch to make tungsten needles allows extremely sharp needles to be made.&#160;The type of torch used makes a big difference because it controls the size of the flame. Electrolytic sharpening can also be used, but this approach does not even come close to producing needles as sharp.&#160;Use tungsten rods instead of spooled wire so that the needles can be made straight.
The Spemann micropipette, while time-consuming and difficult to make, is an ideal instrument for tissue&#160;transfer.&#160;The pipette can be customized with different-sized openings, and can be used repeatedly. A critical factor for using a Spemann&#160;micropipette is to have some fluid in the pipette before touching the tip to the surface of the embryo. Some of the fluid will always flow out when contact is made with the meniscus over the embryo.&#160;Pressing&#160;on the diaphragm allows fluid and graft tissue to be ejected very precisely, whereas slightly letting up on the diaphragm gently sucks the donor graft tissue into the pipette.&#160;Maintaining a bit of positive pressure on the diaphragm keeps the donor graft tissue at the tip of pipette during transfer, and a little additional pressure on the diaphragm allows the donor graft tissue to be deliberately placed in the host.&#160;
For protection of the Spemann pipette during storage and sterilization, remove the bulb from the wide end and carefully insert the tapered tip into the bulb.&#160;Place the Spemann pipette in a glass test tube, cover the top with aluminum foil, and autoclave prior to surgery. After multiple pipettes are ready to be sterilized again for surgery, invert the pipettes, heat them nearly to&#160;a boil in the same solution of distilled water and glassware detergent, and then rinse repeatedly with distilled water. Autoclave pipettes in their individual&#160;tubes. The rubber tubing that forms the diaphragm and the rubber bulb should be replaced after several sterilizations or when they become darkened and stiff.
Many of the components of this protocol involve dangerous equipment. For example, the procedure for making a Spemann Micropipette involves three types of flames as well as heating, pulling, blowing, bending, cutting, and polishing glass. Therefore, wearing proper personal protective equipment (PPE) that increases safety such as goggles and a lab coat is critical. Additionally, because many people suffer from, or have the potential to develop, egg allergies, always use gloves when handling eggs. With these precautions in mind, the quail-duck chimeric system is a safe, efficient, and relatively accessible method that has many future applications.

&#160;The facial skeleton develops from the growth and fusion of multiple facial processes that are composed of neural crest and mesodermal mesenchyme surrounded by ectodermal and endodermal epithelial layers1-11. Morphogenetic events within each process are governed by distinct signaling interactions between the mesenchyme and surrounding epithelia12-16. Alterations to these signaling interactions and/or their downstream effectors contribute to disease phenotypes and may also be relevant to evolution of the craniofacial skeleton17,18. Therefore, elucidating the timing and nature of tissue interactions has great potential to increase our understanding of the developmental and evolutionary biology of the facial skeleton.
The use of chimeric embryos to investigate tissue interactions has a long history in developmental biology. This approach was pioneered by Hans Spemann and his lab who discovered embryonic &#8220;organizers&#8221; by transplanting tissues between embryos of different amphibian species. Spemann was a master of micro-surgical techniques whose hand-skills were complemented by his development of specialized tools, notably the Spemann pipette. Viktor Hamburger was a graduate student in the laboratory of Hans Spemann in Freiburg during the 1920s, which is when the original transplant experiments that led to Spemann&#8217;s Nobel Prize were performed.&#160;When Hamburger moved to Washington University in St. Louis in 1935, he detailed the process of making a Spemann micropipette in his Manual of Experimental Embryology19.&#160;Drew Noden was a graduate student in Hamburger&#8217;s lab at Washington University until 1972.&#160;After moving to the University of Massachusetts, Amherst and then to Cornell University, Noden continued fabricating and using Spemann micropipettes for his surgical transplants involving quail-chick chimeras.&#160;While a graduate student, one of the authors (Rich Schneider) trained with Drew Noden at Cornell from 1995 to 1998.&#160;The following protocol for making a Spemann micropipette is based on descriptions written by Hamburger and Noden, and includes subsequent modifications made by Schneider.
The use of quail-chick chimeras for the study of craniofacial development and especially for understanding the contributions of neural crest cells was pioneered by Noden and by Le Douarin in the early 1970s, reviewed in Le Douarin et al20. This approach has been broadly adopted in many studies and by numerous other investigators1,4,5,21-38. The equivalent rates of growth and morphology of quail and chick make transplants within them ideal for the study of cell fate and lineage tracing. However, because of the similarities between quail and chick, morphological changes induced by donor cells are difficult to decipher. In contrast, other avian chimeric systems have included domestic duck as a way to study mechanisms that make embryos anatomically distinct39-50.&#160;More specifically, the quail-duck chimeric system offers multiple benefits for discerning the effects of the donor on the host, and vice versa. First, quail and duck embryos are distinct in body size and shape, which provides a direct way to explore donor- or host-specific mechanisms of pattern formation by assaying for differential domains of gene expression (Figures 1 A&#160;and B). Second, quail and duck embryos have considerably different rates of maturation, with quail hatching in 17 days and duck hatching in 28 days. Transplanted neural crest maintains its intrinsic maturation rate within the host environment, and thus, identification of temporal changes in gene expression, tissue interactions, histogenesis, and morphogenesis is possible51-57. Finally, the anti-quail nuclear antibody (Q&#162;PN) allows donor and host cellular contributions to be permanently distinguished from one another by recognizing a protein that is ubiquitously expressed in quail cells but absent from duck cells.

<table border="0" cellpadding="0" cellspacing="0" width="746">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:169px;">1x PBS</td>
			<td style="width:180px;">TEK</td>
			<td style="width:145px;">TEKZR114</td>
			<td style="width:252px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:169px;">Hank&#8217;s BSS w/o Phenol Red</td>
			<td style="width:180px;">Invitrogen</td>
			<td style="width:145px;">14025-092</td>
			<td style="width:252px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:169px;">Neutral Red</td>
			<td style="width:180px;">Sigma Aldrich</td>
			<td style="width:145px;">N4638-5G</td>
			<td style="width:252px;">.22&#181;m filter-sterilized</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:169px;">18G Needles</td>
			<td style="width:180px;">BD</td>
			<td style="width:145px;">305195</td>
			<td style="width:252px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:169px;">5 ml syringe</td>
			<td style="width:180px;">BD</td>
			<td style="width:145px;">309646</td>
			<td style="width:252px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:169px;">No. 5 Dumont forceps</td>
			<td style="width:180px;">Fine Science Tools</td>
			<td style="width:145px;">11252-20</td>
			<td style="width:252px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:169px;">Straight Scissors</td>
			<td style="width:180px;">Fine Science Tools</td>
			<td style="width:145px;">14028-10</td>
			<td style="width:252px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:169px;">Curved Scissors</td>
			<td style="width:180px;">Fine Science Tools</td>
			<td style="width:145px;">14029-10</td>
			<td style="width:252px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:169px;">Spemann Pipet</td>
			<td style="width:180px;"></td>
			<td style="width:145px;"></td>
			<td style="width:252px;">Hand-made in lab</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:169px;">Egg holder</td>
			<td style="width:180px;"></td>
			<td style="width:145px;"></td>
			<td style="width:252px;">Glass ashtray and modeling clay</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:169px;">Alcohol burner</td>
			<td style="width:180px;">Fisher</td>
			<td style="width:145px;">04-245-1</td>
			<td style="width:252px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:169px;">Transparent tape</td>
			<td style="width:180px;">3M Scotch</td>
			<td style="width:145px;">600</td>
			<td style="width:252px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:169px;">Glass Stirring Rod</td>
			<td style="width:180px;">Fisher</td>
			<td style="width:145px;">11-380C</td>
			<td style="width:252px;">Tip is narrowed and rounded using a flame</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:169px;">Tungston wire (.004 x 3 inches)</td>
			<td style="width:180px;">A-M Systems</td>
			<td style="width:145px;">7190</td>
			<td style="width:252px;">Tip is flame-sharpened in a propane torch</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Bunsun burner</td>
			<td>Fisher Scientific</td>
			<td>&#160;S49117</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pasteur pipette</td>
			<td>Fisherbrand</td>
			<td>&#160;22-183-632</td>
			<td>9-inch (229 mm)&#160;</td>
		</tr>
		<tr height="81">
			<td height="81" style="height:81px;width:169px;">rubber tubing</td>
			<td>Fisher Scientific&#160;</td>
			<td>14-178C</td>
			<td style="width:252px;">amber, thin wall natural rubber; wall thickness: 0.0625 inches/1.6 mm; O.D.: 0.375 inches/9.5 mm; I.D.: 0.25 inches/6.4mm</td>
		</tr>
		<tr height="61">
			<td height="61" style="height:61px;">&#160;Propane fuel cylinder</td>
			<td>BernzOmatic</td>
			<td>UL2317</td>
			<td style="width:252px;">&#160;TX-9 with torch style &#34;A&#34; with a screw-on brass &#34;pencil flame&#34; torch&#160;&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Diamond point pencil</td>
			<td>Fisher Scientific</td>
			<td>&#160;22-268912</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Rubber bulbs</td>
			<td>Fisherbrand</td>
			<td>S32325</td>
			<td></td>
		</tr>
	</tbody>
</table>

assessing species-specific contributions to craniofacial development using quail-duck chimeras

The generation of chimeric embryos is a widespread and powerful approach to study cell fates, tissue interactions, and species-specific contributions to the histological and morphological development of vertebrate embryos. In particular, the use of chimeric embryos has established the importance of neural crest in directing the species-specific morphology of the craniofacial complex. The method described herein utilizes two avian species, duck and quail, with remarkably different craniofacial morphology. This method greatly facilitates the investigation of molecular and cellular regulation of species-specific pattern in the craniofacial complex.&#160;Experiments in quail and duck chimeric embryos have already revealed neural crest-mediated tissue interactions and cell-autonomous behaviors that regulate species-specific pattern in the craniofacial skeleton, musculature, and integument. The great diversity of neural crest derivatives suggests significant potential for future applications of the quail-duck chimeric system to understanding vertebrate development, disease, and evolution.

Prior to further analysis, the efficiency of the transplant needs to be assayed. For histological, morphological, or gene expression analyses on tissue samples, quail cells should be detected by immunohistochemistry using Q&#162;PN antibody as described36. For RNA analyses, species-specific contributions to tissues of interest can be calculated using a PCR-based strategy58. After the efficacy of the transplant has been validated, further morphological or molecular outcome measures can be evaluated in chimeras. Interactions between the donor neural crest cells and surrounding host-derived tissues that underlie proper histogenesis and morphogenesis of the craniofacial complex have previously been extensively studied. In particular, neural crest mesenchyme directs species-specific morphology of the face7,13,51,59, feather pattern52, muscle pattern56, and cartilage53,57 through the regulation of host gene expression.&#160;For example, neural crest mesenchyme dictates when bone forms in the mandible by temporally regulating Bmp4 expression55.
Recently, investigations have centered on tissue interactions occurring very early in craniofacial development. In this regard, experiments utilizing quail-duck chimeras have shown host embryos influence neural crest migration by determining morphological boundaries. That is, in chimeric quck embryos, donor quail neural crest cells migrate into the host duck mandibular arch in a duck-like pattern (Figure 1D). Despite this host contribution to the size of the neural crest population, the donor neural crest continues to generate a mandibular skeleton that is quail-like in size and shape (Figure 1E).
<img alt="Figure 1" fo:content-width="6in" fo:src="/files/ftp_upload/51534/51534fig1highres.jpg" src="/files/ftp_upload/51534/51534fig1.jpg" /> 
Figure 1.&#160;Quail-Duck Chimeric System. (A) Quail and (B) duck skulls display considerable differences in size and shape, and thus, are ideally suited for using a chimeric system to study craniofacial development (modified from Tokia&#160;et al.56).&#160;(C) Experimental design for generating unilateral chimeric quck embryos from stage-matched HH9.5 quail and duck embryos. The neural fold is removed from one side of quail embryos and transplanted into duck embryos after an equivalent piece of the neural fold has been removed.&#160;(D) Quail donor cells (green) can be followed in chimeras using an anti-quail antibody (Q&#162;PN) as shown in ventral view of HH12 chimeric embryo.&#160;(F) In quck mandibles at HH38, the quail donor-derived Meckel&#8217;s cartilage is shorter and straighter than that observed for the contralateral duck host-derived Meckel&#8217;s cartilage, which is larger and curved. Reprinted Tokita et al., Dev.&#160;Bio.&#160;306, 377 (2007) with permission from Elsevier.

Watch this Scientific Journal Video about Assessing Species-specific Contributions To Craniofacial Development Using Quail-duck Chimeras at JoVE.com

Assessing Species-specific Contributions To Craniofacial Development Using Quail-duck Chimeras

This article describes a method to generate chimeric embryos that are&#160;designed to test the species-specific contributions of neural crest and/or other tissues to craniofacial development.

The generation of chimeric embryos is a widespread and powerful approach to study cell fates, tissue interactions, and species-specific contributions to ...

assessing-species-specific-contributions-to-craniofacial-development

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JoVE Journal

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10.9K vues.  University of California at San Francisco.  Cet article décrit une méthode pour générer des embryons chimériques qui sont conçus pour tester les contributions spécifiques à l'espèce de crête neurale et / ou d'autres tissus de développement cranio-facial. 

Vidéo: Assessing Species-specific Contributions To Craniofacial Development Using Quail-duck Chimeras

Development of New Therapeutic Applications Using Microfluidics

In vivo Bioluminescent Imaging of Mammary Tumors Using IVIS Spectrum

4T1 mouse mammary tumor cells can be implanted sub-cutaneously in nu/nu mice to form palpable tumors in 15 to 20 days. This xenograft tumor model system is valuable for the pre-clinical in vivo evaluation of putative antitumor compounds.
The 4T1 cell line has been engineered to constitutively express the firefly luciferase gene (luc2). When mice carrying 4T1-luc2 tumors are injected with Luciferin the tumors emit a visual light signal that can be monitored using a sensitive optical imaging system like the IVIS Spectrum. The photon flux from the tumor is proportional to the number of light emitting cells and the signal can be measured to monitor tumor growth and development. IVIS is calibrated to enable absolute quantitation of the bioluminescent signal and longitudinal studies can be performed over many months and over several orders of signal magnitude without compromising the quantitative result.
Tumor growth can be monitored for several days by bioluminescence before the tumor size becomes palpable or measurable by traditional physical means. This rapid monitoring can provide insight into early events in tumor development or lead to shorter experimental procedures.
Tumor cell death and necrosis due to hypoxia or drug treatment is indicated early by a reduction in the bioluminescent signal. This cell death might not be accompanied by a reduction in tumor size as measured by physical means. The ability to see early events in tumor necrosis has significant impact on the selection and development of therapeutic agents.
Quantitative imaging of tumor growth using IVIS provides precise quantitation and accelerates the experimental process to generate results.

In vivo Bioluminescent Imaging of Mammary Tumors Using IVIS Spectrum

Mammary tumor cells expressing luciferase are implanted subcutaneously in mice and visualized using optical imaging to monitor tumor growth and development non-invasively in a longitudinal study.

4T1 mouse mammary tumor cells can be implanted sub-cutaneously in nu/nu mice to form palpable tumors in 15 to 20 days. This xenograft tumor model system ...

Tissue Targeted Embryonic Chimeras: Zebrafish Gastrula Cell Transplantation

Certain fundamental questions in the field of developmental biology can only be answered when cells are placed in novel environments or when small groups of cells in a larger context are altered.  Watching how one cell interacts with and behaves in a unique environment is essential to characterizing cell functions.  Determining how the localized misexpression of a specific protein influences surrounding cells provides insightful information on the roles that protein plays in a variety of developmental processes.  Our lab uses the zebrafish model system to uniquely combine genetic approaches with classical transplantation techniques to generate genotypic or phenotypic chimeras.  We study neuron-glial cell interactions during the formation of forebrain commissures in zebrafish. This video describes a method that allows our lab to investigate the role of astroglial populations in the diencephalon and the roles of specific guidance cues that influence projecting axons as they cross the midline. Due to their transparency zebrafish embryos are ideal models for this type of ectopic cell placement or localized gene misexpression. Tracking transplanted cells can be accomplished using a vital dye or a transgenic fish line expressing a fluorescent protein. We demonstrate here how to prepare donor embryos with a vital dye tracer for transplantation, as well as how to extract and transplant cells from one gastrula staged embryo to another.  We present data showing ectopic GFP+ transgenic cells within the forebrain of zebrafish embryos and characterize the location of these cells with respect to forebrain commissures.  In addition, we show laser scanning confocal timelapse microscopy of Alexa 594 labeled cells transplanted into a GFP+ transgenic host embryo.  These data provide evidence that gastrula staged transplantation enables the targeted positioning of ectopic cells to address a variety of questions in Developmental Biology.

Zebrafish cell transplantation enables the combination of genetics and embryology to generate tissue specific chimeras. This video demonstrates gastrula staged cell transplantations that have allowed our lab to investigate the roles of astroglial populations and specific guidance cues during commissure formation in the forebrain.

Certain fundamental questions in the field of developmental biology can only be answered when cells are placed in novel environments or when small groups ...

Dechorionation of Medaka Embryos and Cell Transplantation for the Generation of Chimeras

Medaka is a small egg-laying freshwater fish that allows both genetic and embryological analyses and is one of the three vertebrate model organisms in which genome-wide phenotype-driven mutant screens were carried out 1. 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 the vertebrate genome functions. Manipulation of medaka embryos, such as dechorionation, mounting embryos for imaging and cell transplantation, are key procedures to work on both medaka and zebrafish in a laboratory. Cell transplantation examines cell autonomy of medaka mutations. Chimeras are generated by transplanting labeled cells from donor embryos into unlabeled recipient embryos. Donor cells can be transplanted to specific areas of the recipient embryos based on the fate maps 3 so that clones from transplanted cells can be integrated in the tissue of interest during development. Due to the hard chorion and soft embryos, manipulation of medaka embryos is more involved than in zebrafish. In this video, we show detailed procedures to manipulate medaka embryos.

Due to the hard chorion and soft embryos, manipulation of medaka embryos is more involved than in zebrafish. This video shows step-by-step procedures for how to manipulate medaka embryos, including dechorionation, mounting in agarose for imaging and cell transplantation for the production of chimeras. These procedures are essential to use medaka and zebrafish in a laboratory to take full advantage of their complementary features for the genetic dissection of vertebrate genome functions.

Medaka is a small egg-laying freshwater fish that allows both genetic and embryological analyses and is one of the three vertebrate model organisms in ...

Assessing Signaling Properties of Ectodermal Epithelia During Craniofacial Development

The accessibility of avian embryos has helped experimental embryologists understand the fates of cells during development and the role of tissue interactions that regulate patterning and morphogenesis of vertebrates (e.g., 1, 2, 3, 4). Here, we illustrate a method that exploits this accessibility to test the signaling and patterning properties of ectodermal tissues during facial development. In these experiments, we create quail-chick 5 or mouse-chick 6 chimeras by transplanting the surface cephalic ectoderm that covers the upper jaw from quail or mouse onto either the same region or an ectopic region of chick embryos. The use of quail as donor tissue for transplantation into chicks was developed to take advantage of a nucleolar marker present in quail but not chick cells, thus allowing investigators to distinguish host and donor tissues 7. Similarly, a repetitive element is present in the mouse genome and is expressed ubiquitously, which allows us to distinguish host and donor tissues in mouse-chick chimeras 8. The use of mouse ectoderm as donor tissue will greatly extend our understanding of these tissue interactions, because this will allow us to test the signaling properties of ectoderm derived from various mutant embryos.

This article describes a tissue transplantation technique that was designed to test the signaling and patterning properties of surface cephalic ectoderm during craniofacial development.

The accessibility of avian embryos has helped experimental embryologists understand the fates of cells during development and the role of tissue ...

Electroporation of Craniofacial Mesenchyme

Electroporation is an efficient method of delivering DNA and other charged macromolecules into tissues at precise time points and in precise locations. For example, electroporation has been used with great success to study neural and retinal development in Xenopus, chicken and mouse 1-10. However, it is important to note that in all of these studies, investigators were not targeting soft tissues. Because we are interested in craniofacial development, we adapted a method to target facial mesenchyme.
When we searched the literature, we found, to our surprise, very few reports of successful gene transfer into cartilaginous tissue. The majority of these studies were gene therapy studies, such as siRNA or protein delivery into chondrogenic cell lines, or, animal models of arthritis 11-13. In other systems, such as chicken or mouse, electroporation of facial mesenchyme has been challenging (personal communications, Dept of Craniofacial Development, KCL). We hypothesized that electroporation into procartilaginous and cartilaginous tissues in Xenopus might work better. In our studies, we show that gene transfer into the facial cartilages occurs efficiently at early stages (28), when the facial primordium is still comprised of soft tissue prior to cartilage differentiation.
Xenopus is a very accessible vertebrate system for analysis of craniofacial development. Craniofacial structures are more readily visible in Xenopus than in any other vertebrate model, primarily because Xenopus embryos are fertilized externally, allowing analyses of the earliest stages, and facilitating live imaging at single cell resolution, as well as reuse of the mothers 14. Among vertebrate models developing externally, Xenopus is more useful for craniofacial analysis than zebrafish, as Xenopus larvae are larger and easier to dissect, and the developing facial region is more accessible to imaging than the equivalent region in fish. In addition, Xenopus is evolutionarily closer to humans than zebrafish (&tilde;100 million years closer) 15. Finally, at these stages, Xenopus tadpoles are transparent, and concurrent expression of fluorescent proteins or molecules will allow easy visualization of the developing cartilages. We anticipate that this approach will allow us to rapidly and efficiently test candidate molecules in an in vivo model system.

Craniofacial cartilages develop in close contact with other tissues and are difficult to manipulate in live animals. We are using electroporation to deliver molecular tools during growth of the craniofacial skeleton while bypassing early embryonic effects. This approach will allow us to efficiently test candidate molecules in vivo.

Electroporation is an efficient method of delivering DNA and other charged macromolecules into tissues at precise time points and in precise locations. ...

A Semi-quantitative Approach to Assess Biofilm Formation Using Wrinkled Colony Development

Biofilms, or surface-attached communities of cells encapsulated in an extracellular matrix, represent a common lifestyle for many bacteria. Within a biofilm, bacterial cells often exhibit altered physiology, including enhanced resistance to antibiotics and other environmental stresses 1. Additionally, biofilms can play important roles in host-microbe interactions. Biofilms develop when bacteria transition from individual, planktonic cells to form complex, multi-cellular communities 2. In the laboratory, biofilms are studied by assessing the development of specific biofilm phenotypes. A common biofilm phenotype involves the formation of wrinkled or rugose bacterial colonies on solid agar media 3. Wrinkled colony formation provides a particularly simple and useful means to identify and characterize bacterial strains exhibiting altered biofilm phenotypes, and to investigate environmental conditions that impact biofilm formation. Wrinkled colony formation serves as an indicator of biofilm formation in a variety of bacteria, including both Gram-positive bacteria, such as Bacillus subtilis 4, and Gram-negative bacteria, such as Vibrio cholerae 5, Vibrio parahaemolyticus 6, Pseudomonas aeruginosa 7, and Vibrio fischeri 8.
The marine bacterium V. fischeri has become a model for biofilm formation due to the critical role of biofilms during host colonization: biofilms produced by V. fischeri promote its colonization of the Hawaiian bobtail squid Euprymna scolopes 8-10. Importantly, biofilm phenotypes observed in vitro correlate with the ability of V. fischeri cells to effectively colonize host animals: strains impaired for biofilm formation in vitro possess a colonization defect 9,11, while strains exhibiting increased biofilm phenotypes are enhanced for colonization 8,12. V. fischeri therefore provides a simple model system to assess the mechanisms by which bacteria regulate biofilm formation and how biofilms impact host colonization.
In this report, we describe a semi-quantitative method to assess biofilm formation using V. fischeri as a model system. This method involves the careful spotting of bacterial cultures at defined concentrations and volumes onto solid agar media; a spotted culture is synonymous to a single bacterial colony. This 'spotted culture' technique can be utilized to compare gross biofilm phenotypes at single, specified time-points (end-point assays), or to identify and characterize subtle biofilm phenotypes through time-course assays of biofilm development and measurements of the colony diameter, which is influenced by biofilm formation. Thus, this technique provides a semi-quantitative analysis of biofilm formation, permitting evaluation of the timing and patterning of wrinkled colony development and the relative size of the developing structure, characteristics that extend beyond the simple overall morphology.

We provide a simple, semi-quantitative method to investigate biofilm formation in vitro. This method takes advantage of the Zeiss stemi 2000-C Dissecting Microscope (with camera attachment) to monitor both the timing and pattern of biofilm formation, as assessed by the development of wrinkled colonies.

Biofilms, or surface-attached communities of cells encapsulated in an extracellular matrix, represent a common lifestyle for many bacteria. Within a ...

Conversion of a Capture ELISA to a Luminex xMAP Assay using a Multiplex Antibody Screening Method

The enzyme-linked immunosorbent assay (ELISA) has long been the primary tool for detection of analytes of interest in biological samples for both life science research and clinical diagnostics. However, ELISA has limitations. It is typically performed in a 96-well microplate, and the wells are coated with capture antibody, requiring a relatively large amount of sample to capture an antigen of interest . The large surface area of the wells and the hydrophobic binding of capture antibody can also lead to non-specific binding and increased background. Additionally, most ELISAs rely upon enzyme-mediated amplification of signal in order to achieve reasonable sensitivity. Such amplification is not always linear and can thus skew results.
In the past 15 years, a new technology has emerged that offers the benefits of the ELISA, but also enables higher throughput, increased flexibility, reduced sample volume, and lower cost, with a similar workflow 1, 2. Luminex xMAP Technology is a microsphere (bead) array platform enabling both monoplex and multiplex assays that can be applied to both protein and nucleic acid applications 3-5. The beads have the capture antibody covalently immobilized on a smaller surface area, requiring less capture antibody and smaller sample volumes, compared to ELISA, and non-specific binding is significantly reduced. Smaller sample volumes are important when working with limiting samples such as cerebrospinal fluid, synovial fluid, etc. 6. Multiplexing the assay further reduces sample volume requirements, enabling multiple results from a single sample.
Recent improvements by Luminex include: the new MAGPIX system, a smaller, less expensive, easier-to-use analyzer; Low-Concentration Magnetic MagPlex Microspheres which eliminate the need for expensive filter plates and come in a working concentration better suited for assay development and low-throughput applications; and the xMAP Antibody Coupling (AbC) Kit, which includes a protocol, reagents, and consumables necessary for coupling beads to the capture antibody of interest. (See Materials section for a detailed list of kit contents.)
In this experiment, we convert a pre-optimized ELISA assay for TNF-alpha cytokine to the xMAP platform and compare the performance of the two methods 7-11. TNF-alpha is a biomarker used in the measurement of inflammatory responses in patients with autoimmune disorders.
We begin by coupling four candidate capture antibodies to four different microsphere sets or regions. When mixed together, these four sets allow for the simultaneous testing of all four candidates with four separate detection antibodies to determine the best antibody pair, saving reagents, sample and time. Two xMAP assays are then constructed with the two most optimal antibody pairs and their performance is compared to that of the original ELISA assay in regards to signal strength, dynamic range, and sensitivity.

An ELISA can be easily converted to a Luminex xMAP assay and, through the benefits of multiplexing, several antibodies can be screened simultaneously to identify an optimum antibody pair, resulting in increased sensitivity and dynamic range, while reducing assay cost.

The enzyme-linked immunosorbent assay (ELISA) has long been the primary tool for detection of analytes of interest in biological samples for both life ...

Cell Tracking Using Photoconvertible Proteins During Zebrafish Development

Embryogenesis is a dynamic process that is best studied by using techniques that allow the documentation of developmental changes in vivo. The use of genetically-encoded fluorescent proteins has proven a valuable strategy for elucidating dynamic morphogenetic processes as they occur in the intact organism. During the past decade, the development of photoactivatable and photoconvertible fluorescent proteins has opened the possibility to investigate the fate of discrete subpopulations of tagged proteins1. Unlike photoactivatable proteins, photoconvertible fluorescent proteins (PCFPs) are readily tracked and imaged in their native emission state prior to photoconversion, making it easier to identify and select regions by optical inspection. PCFPs, such as Kaede2, KikGR3, Dendra4 and EosFP5, can be shifted from green to red upon exposure to UV or blue light due to a His-Tyr-Gly tripeptide sequence which forms a green chromophore that can be photoconverted to a red one by a light-catalyzed &beta;-elimination and subsequent extension of a &pi;-conjugated system3. PCFPs and their monomeric variants are useful tools for tracking cells6-10 and studying protein dynamics11-14, respectively. During recent years, PCFPs have been expressed in different animal model, such as zebrafish6, chicken7,8 and mouse9,10 for cell fate tracking. Here we report a protocol for cell-specific photoconversion of PCFPs in the living zebrafish embryo and further tracking of photoconverted proteins at later developmental stages. This methodology allows studying, in a tissue-specific manner, cell biological events underlying morphogenesis in the zebrafish animal model.

Here, we present a method for the photoactivated switch of photoconvertible fluorescent proteins (PCFPs) in the living zebrafish embryo and further tracking of photoconverted protein at specific time points during development. This methodology allows monitoring of cell biological events underlying different developmental processes in a live vertebrate organism.

Embryogenesis is a dynamic process that is best studied by using techniques that allow the documentation of developmental changes in vivo. The use of ...

Assessing Murine Resistance Artery Function Using Pressure Myography

Pressure myograph systems are exquisitely useful in the functional assessment of small arteries, pressurized to a suitable transmural pressure. The near physiological condition achieved in pressure myography permits in-depth characterization of intrinsic responses to pharmacological and physiological stimuli, which can be extrapolated to the in vivo behavior of the vascular bed. Pressure myograph has several advantages over conventional wire myographs. For example, smaller resistance vessels can be studied at tightly controlled and physiologically relevant intraluminal pressures. Here, we study the ability of 3rd order mesenteric arteries (3-4 mm long), preconstricted with phenylephrine, to vaso-relax in response to acetylcholine. Mesenteric arteries are mounted on two cannulas connected to a pressurized and sealed system that is maintained at constant pressure of 60 mmHg. The lumen and outer diameter of the vessel are continuously recorded using a video camera, allowing real time quantification of the vasoconstriction and vasorelaxation in response to phenylephrine and acetylcholine, respectively.
	To demonstrate the applicability of pressure myography to study the etiology of cardiovascular disease, we assessed endothelium-dependent vascular function in a murine model of systemic hypertension. Mice deficient in the &alpha;1 subunit of soluble guanylate cyclase (sGC&alpha;1-/-) are hypertensive when on a 129S6 (S6) background (sGC&alpha;1-/-S6) but not when on a C57BL/6 (B6) background (sGC&alpha;1-/-B6). Using pressure myography, we demonstrate that sGC&alpha;1-deficiency results in impaired endothelium-dependent vasorelaxation. The vascular dysfunction is more pronounced in sGC&alpha;1-/-S6 than in sGC&alpha;1-/-B6 mice, likely contributing to the higher blood pressure in sGC&alpha;1-/-S6 than in sGC&alpha;1-/-B6 mice. 
	Pressure myography is a relatively simple, but sensitive and mechanistically useful technique that can be used to assess the effect of various stimuli on vascular contraction and relaxation, thereby augmenting our insight into the mechanisms underlying cardiovascular disease.

In pressure myography, an intact small segment of a vessel is mounted onto two small cannulas and pressurized to a suitable luminal pressure. Here, we describe the method to measure vasorelaxation response of the mouse 3rd order mesenteric arteries in c57 and sGC&alpha;1-/- mice using pressure myography.

Pressure myograph systems are  exquisitely useful in the functional assessment of small arteries, pressurized  to a suitable transmural pressure. The  ...