The authors were supported by: a CancerFree KIDS Research Grant, a March of Dimes Basil O&#8217;Connor Starter Scholar Award (#5-FY14-32), a Cincinnati Children&#8217;s Hospital Medical Center (CCHMC) Trustee Grant Award; a CCHMC Research Innovation and Pilot Funding Award; and CCHMC developmental funds. Authors also acknowledge the Capel laboratory for the initial optimization of this technique.

All mice used in these studies were CD-1 mice obtained from Charles River Laboratories. Previous culture experiments have also been performed on other strains, such as C57BL/6J (data not shown), but any strain can be used. Pregnant adult females were approximately 2-3 months old and were euthanized via CO2 inhalation followed by cervical dislocation and bilateral thoracotomy prior to embryo removal. Mice were housed in accordance with NIH guidelines, and experimental protocols were approved by the Institutional Animal Care and Use Committee of Cincinnati Children&#8217;s Hospital Medical Center.
1. Preparation of Instruments, Culture Media, and Dishes
<ol>
	<li>Prepare a 70% ethanol solution. Autoclave deionized water (for making humidified chambers and for making/diluting solutions). Sterilize dissection tools (forceps, scissors, and 27 G needle syringes) by spraying down with 70% ethanol.</li>
	<li>Prepare 10X phosphate-buffered saline (PBS) containing calcium and magnesium (80 g NaCl, 2 g KCl, 14.4 g Na2HPO4, 2.4 g KH2PO4, 6.75 ml&#160;of 1 M CaCl2, 3.75 ml&#160;of 1 M MgCl2). Stir to dissolve in 1 L sterile autoclaved water and then filter with filter paper. Dilute 10-fold with water to make 1X PBS.</li>
	<li>Heat inactivate fetal bovine serum (FBS) at 55 &#176;C for 30 min and filter sterilize with a 0.22 &#181;m syringe filter.</li>
	<li>Prepare 38 ml&#160;1X complete culture medium (called complete DMEM, cDMEM), consisting of Dulbecco&#8217;s Modified Eagle Medium (DMEM), 10% filtered heat-inactivated FBS and 1/100 dilution of penicillin-streptomycin stock. This usually should be used fresh, but can be stored at 4 &#176;C for 1 month.</li>
	<li>Reconstitute the VEGFR Tyrosine Kinase Inhibitor II (TKI II) in DMSO for a stock concentration of 5 mg/ml, and dilute stock with cDMEM to make a working solution. The final concentration for this study is 1.875 &#181;g/ml&#160;in cDMEM. According to the company&#8217;s recommendations, stock solutions are stable for up to 6 months at -20 &#176;C. As for working solutions, make fresh and use on the same day. 
	Note: The concentration of this drug in the working solution should be two-fold higher than the desired final concentration (since it will be diluted two-fold in the droplet). At the same time, prepare the same volume of cDMEM containing the same volume of DMSO for the &#8220;control&#8221; treatment.</li>
	<li>Prepare the tail digestion reagents (50 mM NaOH and 1 M Tris-HCl [pH 8.0]) separately in distilled water.</li>
	<li>Make a 25 &#956;M stock of the XY PCR primers (XY Forward 5&#39;-TGA AGC TTT TGG CTT TGA G-3&#39; and XY Reverse 5&#39;-CCG CTG CCA AAT TCT TTG G-3&#39;) together in nuclease-free water.</li>
	<li>Prepare 50X TAE buffer (242 g Trizma Base, 57.1 ml&#160;Glacial Acetic Acid, 100 ml&#160;0.5 M EDTA, pH 8.5, and add water to 1 L final volume). Prepare 1X TAE Running buffer (20 ml&#160;50X TAE diluted with water to 1 L total volume).</li>
	<li>To create a 2.5% agarose solution (0.625 g agarose, 0.5 ml&#160;50X TAE Buffer, 24.5 ml&#160;water), heat agarose solution in microwave until boiling, then let cool until about 55-65 &#176;C (able to touch). Once cool, add 2 &#181;l&#160;of 1% ethidium bromide solution, and pour gel. 
	CAUTION! Ethidium bromide is a mutagen and teratogen; please use nitrile gloves and proper safety protocol when handling. Alternatively, other potentially less toxic gel resolving agents or dyes can be used.</li>
	<li>Prepare Blocking Solution (PBS with 0.1% Triton X-100, 10% fetal bovine serum [FBS], and 3% bovine serum albumin [BSA]) for immunofluorescence.</li>
	<li>Prepare Washing Solution (PBS with 0.1% Triton X-100, 1% FBS, and 3% BSA) for immunofluorescence.</li>
	<li>Prepare PBTx (PBS with 0.1% Triton X-100) for immunofluorescence.</li>
	<li>Prepare the incubation chambers for culture. Fill large (100 mm diameter) petri dishes with 35 ml&#160;autoclaved water. Place near where dissection and cultures will be performed.</li>
</ol>
2. Isolation of Fetal Testes from Mus musculus
<ol>
	<li>Check the female mouse for vaginal plugs each morning. Noon on the day at which the vaginal plug is detected is considered E0.5. Euthanize the pregnant female mouse at E11.5, in a manner consistent with approved animal protocols.</li>
	<li>Spray down mouse abdomen with 70% ethanol.</li>
	<li>Open the belly skin and peritoneum with a V-shaped incision using fine scissors and pull back flap of tissue to expose interior organs.</li>
	<li>Locate the ovaries at both ends of the uterine horn. Using scissors, cut at the ovary and the connective tissue to separate the uterus containing the embryos from the mother&#8217;s body.</li>
	<li>Open the uterine wall by gently cutting the side opposite the placentae, exposing the yolk sacs. Be careful not to puncture yolk sac to avoid damaging or losing embryos.</li>
	<li>Cut near the placentae to remove the embryo-containing yolk sacs and place them into a dish containing PBS with calcium and magnesium. The presence of calcium and magnesium ions will help support cell adhesion molecules to maintain tissue architecture and prevent the tissue from sticking to the dissection tools.</li>
	<li>With forceps, remove the yolk sac and amnion from the embryo by carefully puncturing the yolk sac to create a hole in which the embryo can slide out. Once the embryo is outside of the yolk sac, cut the umbilical cord.</li>
	<li>From this point forward, use a microscope located in a sterile tissue culture hood. Remove the head of embryo and discard by pinching with forceps on either side of neck.</li>
	<li>Remove the tail and place into new microcentrifuge tube for XY PCR analysis to determine sex of embryo. 
	Note: While it is optimal to culture gonads as soon as possible after harvest, it is also possible to remove tail DNA and perform XX/XY genotyping prior to culture, so that the sex of each embryo is known and only the desired sex needs to be cultured. While the genotyping is being done, embryos can be kept separated (so that they can be tracked) at 4 &#176;C or on ice until ready for culture. One caveat is that this period outside in utero or culture conditions may potentially affect viability for culture or subsequent development during culture.</li>
	<li>Secure the embryo in a supine position against the bottom of the culture dish by pinning the armpits of embryo with one pair of forceps (usually the forceps held in the weak hand). Maintain these forceps in place to hold the embryo still during the entire dissection process.</li>
	<li>With other pair of forceps, remove skin covering abdomen and gently remove liver, intestines, and other organs to expose back body wall.</li>
	<li>As the gonads will be located on the back body wall on either side of the dorsal aorta, a large blood vessel running along the midline of the body, scoop underneath the urogenital ridge with closed forceps and remove the urogenital ridge by opening the forceps and lifting up. As the gonads will be loosely attached to the wall, be careful not to pull up too fast without removing these connections or the gonads may stretch, causing damage to the organs.</li>
	<li>Move the urogenital ridge into cDMEM in a dish to acclimate tissue to media.</li>
	<li>Separate the gonad-mesonephros complex from the rest of the urogenital ridge using 27 G needles. Use one needle to cut by pressing down, and use the other to guide the tissue correctly to allow optimal separation. Avoid using a sawing motion against the dish bottom, as sharp needles will create shards of plastic that can stick to the tissue. Make sure to keep the mesonephros completely attached to the gonad.</li>
</ol>
3. Culturing of Gonad with a Small-molecule Inhibitor
<ol>
	<li>Set two 20&#160;&#181;l&#160;pipettes to 15 &#181;l&#160;each. Cut about 1-2 mm off one of the barrier pipette tips using a clean, sterilized razor blade for transfer of the gonads and addition of control cDMEM, while the other pipette will be used solely for drug-treated cDMEM.</li>
	<li>Line up gonads with their long axis parallel to the pipette tip so they can easily be pipetted using the cut-off tip. Be careful of gonads sticking to the inside of the pipette tip.</li>
	<li>Label one side as the control droplet and the other as the drug-containing droplet. Only two droplets can fit on a 35 mm dish lid. Make sure that labels on lids match the labels on tail-tissue-containing microcentrifuge tubes.</li>
	<li>Pipette 15 &#181;l&#160;of cDMEM containing a single gonad into a droplet in the lid of a small (35 mm) culture dish (use only the lids, as the bottoms are too tall to fit within a humidified petri dish chamber). Place two separate gonad-containing droplets on either side of the dish, well-separated from one another. Check under the microscope to ensure that the gonads have been transferred to the droplets.</li>
	<li>To the droplet designated as the control, add an additional 15 &#181;l&#160;of cDMEM containing DMSO (made in step 1.5) to make a droplet with a 30 &#181;l&#160;total volume. Use only the &#8220;control&#8221; pipette that will not contact the drug.</li>
	<li>To the other droplet (drug-treated sample), add 15 &#181;l&#160;of TKI-II-containing cDMEM to make a droplet with a 30 &#181;l&#160;total volume. Use only the pipette designated for drug-treated samples.</li>
	<li>Using the pipette, spread out the droplets in an expanding circular pattern until they are about 15-18 mm in diameter and the gonad is located roughly in the middle. Orient the gonad so that it lies on its side and the gonad and mesonephros are easily distinguishable. Orient the lung so that the two lobes lay flat.
	<ol>
		<li>Although droplet diameter may vary slightly, ensure that gonads are not floating and are held in place by surface tension. Make sure that droplets do not touch each other or the side of the lid. Without the surface tension, the organs will grow onto the lid during the culture and flatten, thus distorting organ morphology.</li>
	</ol>
	</li>
	<li>Carefully place the culture dish lid containing the droplets upright onto the surface of the water in the large (100 mm) humidifier dish. Ensure there are no bubbles trapped underneath the bottom of the lid and that it is lying flat on the surface of the water. Do not invert the lid and be careful not to let any water touch the top of the lid where the droplets are located.</li>
	<li>To create a small, humidified chamber, immediately cover with the lid of the larger humidifier dish to enclose the gonad cultures. Act as quickly as possible to minimize any evaporation of media. Ensure that the smaller dish has room between it and the lid of the larger dish to move around freely; otherwise, condensation may make a seal and block air exchange to the tissue.</li>
	<li>Once two small lids are placed into the chamber, immediately place the chamber into the incubator. Incubate the droplet cultures for 48 hr in a humidified CO2 incubator at 37 &#176;C.</li>
</ol>
4. Polymerase Chain Reaction for Determining the Sex of Embryos
<ol>
	<li>Add the embryonic tails to bottom of a 1.5 ml&#160;microcentrifuge tube.</li>
	<li>Add to each tube 200 &#181;l&#160;of 50 mM NaOH.</li>
	<li>Place at 95 &#176;C for 15 min or until tissue is completely dissolved.</li>
	<li>Add 50 &#181;l&#160;of 1M Tris-HCl and vortex lightly and briefly.</li>
	<li>In a 1.5 ml&#160;microcentrifuge tube, make a fresh PCR master mix by multiplying each volume by the total number of samples: 0.5 &#181;l&#160;25 &#181;M Primer solution, 2.5 &#181;l&#160;10X Taq Buffer, 0.5 &#181;l&#160;10 mM dNTPs (nucleotides), 19.3 &#181;l&#160;nuclease-free water, 0.2 &#181;l&#160;DNA Taq polymerase. Mix well.</li>
	<li>Add 23 &#181;l&#160;of PCR master mix to PCR tubes containing 3 &#956;l&#160;lysate of digested tails.</li>
	<li>Flick tubes to mix them, then perform a short spin to bring the samples to the bottom of the tube.</li>
	<li>Run XY (Short) PCR program (Table 1).</li>
	<li>Load the samples (with 5x dye) and ladder in a 2.5% agarose gel in 1X TAE buffer.</li>
	<li>Run the agarose gel electrophoresis until XX and XY bands can be resolved.</li>
	<li>Image the gel with UV light and capture image.</li>
	<li>Analyze the X-chromosome-specific vs Y chromosome-specific bands (X chromosome=331 base pairs [bp] and XY=302 bp)13; XY (male) samples will have two bands of 331 and 302 bp, whereas XX (female) samples will appear as a single 331 bp band.</li>
</ol>
5. Whole Mount Organ Immunofluorescence
Day 1:
<ol>
	<li>Remove the gonads from the culture using a 1,000 &#181;l&#160;pipette with a cut-off tip. If desired, they may be placed in a dish of PBS to wash off media and reagents. Place into PBS in a 0.5 ml&#160;microcentrifuge tube. The smaller tube size will conserve reagents and make it easier to keep track of samples in the tube.
	<ol>
		<li>Be sure to keep control and treated samples, as well as XX and XY samples, in separate tubes and remove them from culture with separate sets of pipettes to avoid potential drug contamination.</li>
	</ol>
	</li>
	<li>Wash the gonads twice with PBS. From this point on, this protocol can use 1X PBS lacking calcium and magnesium. To wash them, allow the gonads to sink to the bottom of the tube (by gravity) and then remove the liquid above. Be careful not lose gonads by pipetting too close to them.</li>
	<li>Remove as much PBS as possible from the tube. Add 250 &#181;l&#160;of 4% paraformaldehyde in PBS containing 0.1% Triton X-100, and let incubate O/N at 4 &#176;C on a rocker. Alternatively, this fixation can be done for 2 hr at 4 &#176;C on a rocker. CAUTION! Paraformaldehyde is a toxic substance, so follow safety protocol when handling.</li>
</ol>
Day 2:
<ol start="4">
	<li>Rinse the gonads twice in 250 &#181;l&#160;PBTx at RT.</li>
	<li>Wash the gonads 3 times with 250 &#181;l&#160;PBTx for 10 min each at RT on a rocker.</li>
	<li>Block the gonads at least 1 hr in 250 &#181;l&#160;Blocking solution at RT on a rocker.</li>
	<li>Stain the gonads O/N at 4 &#176;C on a rocker in 250 &#181;l&#160;primary antibody diluted in Blocking Solution.</li>
</ol>
Day 3:
<ol start="8">
	<li>Rinse the gonads twice in 250 &#181;l&#160;Washing Solution.</li>
	<li>Wash the gonads 3 times with 250 &#181;l&#160;Washing Solution for 10 min each at RT on a rocker.</li>
	<li>Block the gonads 1 hr in 250 &#181;l&#160;Blocking Solution at RT on a rocker.</li>
	<li>Cover the microcentrifuge tube with aluminum foil to protect fluorescent secondary antibodies from light.</li>
	<li>Incubate the gonads 2-4 hr at RT on a rocker in 250 &#181;l&#160;secondary antibody diluted in Blocking Solution containing Hoechst 33342. Alternatively, perform secondary antibody and Hoechst 33342 incubation O/N at 4 &#176;C on rocker.</li>
	<li>Rinse the gonads twice in 250 &#181;l&#160;PBTx.</li>
	<li>Wash the gonads 3 times in 250 &#181;l&#160;PBTx for 10 min&#160;each at RT on a rocker.</li>
	<li>Using a cut-off pipette tip, transfer gonads onto a slide with minimal liquid. Remove excess liquid with pipette, but make sure that tissue does not dry out.</li>
	<li>Orient the gonads in desired fashion with forceps, and quickly add a drop of mounting media to mount the gonads on the slide. Make sure the mounting media coats all samples completely; it is critical to avoid letting samples dry out.</li>
	<li>Use coverslips (number 1.5 style) of appropriate size to gently cover samples. Let mounting media spread to contact the entire surface of coverslip. If necessary, very lightly press on the corners of the coverslip so that there are no large air bubbles.</li>
	<li>Seal the slides with clear nail polish around the edges to reduce evaporation of the mounting media. Store mounted slides in the dark at 4 &#176;C.</li>
</ol>

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The authors declare that they have no competing financial interests.

This study demonstrates an ex vivo whole organ droplet method that has many potential applications for studying fetal development. This technique can be used for multiple organs, and allows the researcher to address biological questions that are difficult to examine using in vivo approaches due to inaccessibility of embryos and potential embryonic lethality. This culture method has additional benefits over other in vitro approaches such as mammalian cell lines: whole organs can be used, therefo...

Organ regeneration in vivo in humans is very limited; therefore, tissue engineering, the development of tissues and organs from individual cells donated by a host, is becoming an attractive potential therapy for organ replacement. However, for this therapeutic strategy to be successful, factors and cellular interactions involved in morphogenesis of the organ must be thoroughly studied and well-understood. Due to the inability to study development of specific organs with traditional approaches, researchers have turned to alternative whole embryo or whole organ cultures. Kalaskar et al.1 have shown that ex vivo whole embryogenesis culture yields comparable results (in 58% of cultured embryos) to in utero development, suggesting that ex vivo culture methods are a feasible alternative for organogenesis studies.
An individualized organ culture system, such as this ex vivo droplet culture system, allows for whole organ analysis independent of systemic effects, while permitting manipulation of a specific signaling pathway or cellular interactions via addition of pharmacological reagents or antibodies. Traditionally, the study of fetal organ development has been limited to transgenic and knockout mouse technologies, in addition to pharmacological reagents delivered maternally. However, there are technical issues involving these techniques and treatments in vivo; most concerns revolve around the effects of influencing various organs simultaneously which often results in embryonic lethality. An additional concern of studies manipulating fetal development pharmacologically is the maternal effect of drugs on embryonic development in utero (e.g., maternal metabolism of the drug before it reaches the embryo) and if such reagents can pass through the placental barrier.
The whole organ culture technique described here was adapted from a protocol first described by Maatouk et al.2, in which whole fetal gonads are incubated in ex vivo upright droplet cultures. One significant advantage of culturing fetal gonads is that small-molecule inhibitors can readily access the whole organ by simple diffusion. DeFalco et al. have shown that utilizing this ex vivo droplet culture method in conjunction with small-molecule inhibitors can be used to study signaling processes and interactions occurring during gonad development3; these processes would be difficult to examine in vivo due to technical challenges (e.g., passage of drugs through the placenta or lethality of affecting multiple organs using genetic or pharmacological approaches).
The droplet culture is not only an improvement in certain aspects over in utero experimentation, but also it is an improvement over in vitro and ex vivo systems as well. The use of cell lines to study morphogenesis is extremely difficult because they lack the diverse cell types, lack critical extracellular matrix (ECM) components that permit the formation of organ architecture, and can exhibit artifacts in signaling cascades. Although tissue engineering has made significant improvements in creating scaffolds simulating ECM, the lack of knowledge with regard to which signals are required by each cell type during organogenesis makes it challenging to build an organ system in vitro. Other ex vivo systems have been previously established to study organogenesis, or more specifically morphogenesis, and have been very successful for live imaging of fetal organs in agar4, transwells5, filters6, and other scaffold matrices7,8. The advantage of the droplet culture system is that it allows the study of morphogenesis by providing the ability to utilize less reagents, which are often expensive, but also giving the organ surface tension, which is important for growth and signaling capabilities9.
In the mouse, initial testis morphogenesis takes place between embryonic (E) stages E11.5 and E13.5; these stages comprise the optimal time window for examining factors that influence sex-specific differentiation. Among the critical processes that occur during testis formation are the generation of testis cord architecture and the formation of a testis-specific vascular network. Utilizing this ex vivo whole organ droplet culture system, one is able to alter male-specific vascularization and inhibit testis morphogenesis through the use of a small-molecule inhibitor that blocks the activity of the receptors for vascular endothelial growth factor (VEGF); VEGF-mediated vascular remodeling is critical for testis development10-12. This technique can successfully be applied to other organs and can target specific time windows of development. Whole-mount organ imaging allows the visualization of vital structures as well as structural and cellular changes resulting from the administration of various inhibitors. Importantly, this system is advantageous in that the researcher can bypass potential confounding effects from maternal drug administration or systemic disruption during in vivo targeted gene strategies. Thus, this whole organ ex vivo droplet culture system can significantly improve the ability to understand the interactions and signaling which occur specifically within particular organs during fetal development.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Superfrost Plus Microscope Slides</td>
			<td>Fisherbrand</td>
			<td>12-550-15</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover Glasses: Squares (22 mm x 22 mm, No. 1.5)</td>
			<td>Fisherbrand</td>
			<td>12-541B</td>
			<td></td>
		</tr>
		<tr>
			<td>Sally Hansen Xtreme Wear Nail Polish, Invisible</td>
			<td>Sally Hansen</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>8-Strip 0.2 mL PCR Tubes &#38; Detached Flat Caps</td>
			<td>GeneMate</td>
			<td>T3218-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipetman L P1000L, P200L, P20L, P10L, P2L</td>
			<td>Gilson</td>
			<td>FA10006M, FA10005M, FA10003M, FA10002M, FA10001M</td>
			<td></td>
		</tr>
		<tr>
			<td>Dumont #5 Forceps</td>
			<td>FST</td>
			<td>91150-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine Scissors</td>
			<td>FST</td>
			<td>91460-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Posi-Click 1.7 ml microcentrifuge tubes</td>
			<td>Denville</td>
			<td>C2170</td>
			<td></td>
		</tr>
		<tr>
			<td>Posi-Click 0.6 ml microcentrifuge tubes</td>
			<td>Denville</td>
			<td>C2176</td>
			<td></td>
		</tr>
		<tr>
			<td>10 &#956;l SHARP Precision Barrier Tips</td>
			<td>Denville</td>
			<td>P1096FR</td>
			<td></td>
		</tr>
		<tr>
			<td>20 &#956;l SHARP Precision Barrier Tips</td>
			<td>Denville</td>
			<td>P1121</td>
			<td></td>
		</tr>
		<tr>
			<td>200 &#956;l SHARP Precision Barrier Tips</td>
			<td>Denville</td>
			<td>P1122</td>
			<td></td>
		</tr>
		<tr>
			<td>1000 &#956;l SHARP Precision Barrier Tips</td>
			<td>Denville</td>
			<td>P1126</td>
			<td></td>
		</tr>
		<tr>
			<td>1 ml syringe with 27gauge needles</td>
			<td>BD PrecisionGlide</td>
			<td>309623</td>
			<td></td>
		</tr>
		<tr>
			<td>10 ml syringe</td>
			<td>BD</td>
			<td>305559</td>
			<td></td>
		</tr>
		<tr>
			<td>0.2 &#956;M PES syringe filter</td>
			<td>VWR</td>
			<td>28145-501</td>
			<td></td>
		</tr>
		<tr>
			<td>Grade 3 Qualitative Filter Paper Standard Grade, circle, 185 mm</td>
			<td>Whatman</td>
			<td>1003-185</td>
			<td></td>
		</tr>
		<tr>
			<td>Primaria 35mm Easy Grip Style Cell Culture Dish</td>
			<td>Falcon/Corning</td>
			<td>353801</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri Dishes, Sterile (100 mm x 15 mm)</td>
			<td>VWR</td>
			<td>25384-088</td>
			<td></td>
		</tr>
		<tr>
			<td>New Brunswick Galaxy 14 S CO2 Incubator</td>
			<td>Eppendorf</td>
			<td>CO14S-120-0000</td>
			<td></td>
		</tr>
		<tr>
			<td>Biosafety Cabinet</td>
			<td>Nuare</td>
			<td>NU-425-400</td>
			<td></td>
		</tr>
		<tr>
			<td>Mini-centrifuge&#160;</td>
			<td>Fisher Scientific</td>
			<td>05-090-100</td>
			<td></td>
		</tr>
		<tr>
			<td>BioExpress GyroMixer Variable XL</td>
			<td>GeneMate</td>
			<td>R-3200-1XL</td>
			<td></td>
		</tr>
		<tr>
			<td>Mastercycler Pro Thermal Cycler with control panel</td>
			<td>Eppendorf</td>
			<td>950040015</td>
			<td></td>
		</tr>
		<tr>
			<td>SMZ445 stereomicroscope</td>
			<td>Nikon</td>
			<td>SMZ445</td>
			<td></td>
		</tr>
		<tr>
			<td>MultiImage Light Cabinet with AlphaEase Software</td>
			<td>Alpha Innotech Corporation</td>
			<td>Discontinued</td>
			<td></td>
		</tr>
		<tr>
			<td>Absolute 200 proof Ethanol</td>
			<td>Fisher</td>
			<td>BP2818-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Fisher</td>
			<td>BP151-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Phosphate (Dibasic MW 142) Na2HPO4</td>
			<td>Fisher</td>
			<td>S374-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Phosphate (Monobasic MW 136) KH2PO4</td>
			<td>Sigma-Aldrich</td>
			<td>P5379-1KG</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Chloride (NaCl)</td>
			<td>Fisher</td>
			<td>S671-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium Chloride (KCl)</td>
			<td>Sigma-Aldrich</td>
			<td>P3911-1KG</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium Chloride (MgCl2)</td>
			<td>Sigma</td>
			<td>M2393-100g</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium Chloride (CaCl2)</td>
			<td>Sigma</td>
			<td>C5670-100g</td>
			<td></td>
		</tr>
		<tr>
			<td>Ambion Nuclease-Free Water</td>
			<td>Life Technologies</td>
			<td>AM9938&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>XY PCR Primer&#160;</td>
			<td>IDT</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Glacial Acetic Acid</td>
			<td>Fisher</td>
			<td>A38-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Ethylenediamine Tetraacetic Acid (EDTA)</td>
			<td>Fisher</td>
			<td>BP2482-1</td>
			<td></td>
		</tr>
		<tr>
			<td>1% Ethidium bromide solution</td>
			<td>Fisher</td>
			<td>BP1302-10</td>
			<td>Toxic</td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>GeneMate</td>
			<td>E-3120-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Hydroxide (NaOH)</td>
			<td>Sigma-Aldrich</td>
			<td>367176-2.5KG</td>
			<td></td>
		</tr>
		<tr>
			<td>Trizma Base</td>
			<td>Sigma</td>
			<td>T1503-1KG</td>
			<td></td>
		</tr>
		<tr>
			<td>dNTP Set, 100 mM Solutions</td>
			<td>Thermo Scientific</td>
			<td>R0182</td>
			<td></td>
		</tr>
		<tr>
			<td>DNA Choice Taq polymerase with 10x Buffer</td>
			<td>Denville</td>
			<td>CB-4050-3</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde</td>
			<td>Fisher</td>
			<td>O4042-500</td>
			<td>Toxic</td>
		</tr>
		<tr>
			<td>FluorMount-G</td>
			<td>Southern Biotech</td>
			<td>0100-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrogen Chloride (HCl)</td>
			<td>Fisher</td>
			<td>A144212</td>
			<td></td>
		</tr>
		<tr>
			<td>Bovine Serum Albumin (BSA), powder, Fraction V, Heat shock isolation</td>
			<td>Bioexpress</td>
			<td>0332-100g</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagle Medium (DMEM)&#160;</td>
			<td>Life Technologies</td>
			<td>11965-092</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bovine Serum (FBS), triple 100-nm filtered</td>
			<td>Fisher</td>
			<td>03-600-511</td>
			<td>Heat-inactivate before using</td>
		</tr>
		<tr>
			<td>Penicillin-Streptomycin (10,000 U/mL)</td>
			<td>Life Technologies</td>
			<td>15140-122</td>
			<td>Use at 1:100</td>
		</tr>
		<tr>
			<td>Dimethyl sulfoxide (DMSO), Hybri-max, sterile-filtered</td>
			<td>Sigma</td>
			<td>D2650</td>
			<td></td>
		</tr>
		<tr>
			<td>VEGFR Tyrosine Kinase Inhibitor II - CAS 269390-69-4 - Calbiochem</td>
			<td>EMD Millipore</td>
			<td>676481</td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit Anti-Sox9 Antibody</td>
			<td>Millipore</td>
			<td>AB5535</td>
			<td>Use at dilution: 1:4,000</td>
		</tr>
		<tr>
			<td>Rat Anti-Mouse PECAM1 (CD31) Antibody</td>
			<td>BD Pharmingen</td>
			<td>553370</td>
			<td>Use at dilution: 1:250</td>
		</tr>
		<tr>
			<td>Rabbit Cleaved Caspase-3 (Asp175) Antibody</td>
			<td>Cell Signaling</td>
			<td>9661S</td>
			<td>Use at dilution: 1:250</td>
		</tr>
		<tr>
			<td>Rat E-cadherin / CDH1 Antibody (ECCD-2)</td>
			<td>Life Technologies</td>
			<td>13-1900</td>
			<td>Use at dilution: 1:500</td>
		</tr>
		<tr>
			<td>Hoechst 3342, trihydrochloride, trihydrate</td>
			<td>Invitrogen (Molecular Probes)</td>
			<td>H1399</td>
			<td>Use at 2ug/ml</td>
		</tr>
		<tr>
			<td>Cy3 AffiniPure Donkey Anti-Rat IgG (H+L)</td>
			<td>Jackson Immunoresearch</td>
			<td>712-165-153</td>
			<td>Use at dilution: 1:500</td>
		</tr>
		<tr>
			<td>Alexa Fluor 647 AffiniPure Donkey Anti-Rat IgG (H+L)</td>
			<td>Jackson Immunoresearch</td>
			<td>712-605-153</td>
			<td>Use at dilution: 1:500</td>
		</tr>
		<tr>
			<td>Donkey anti-Rabbit IgG (H+L) Secondary Antibody, Alexa Fluor 555 conjugate</td>
			<td>Life Technologies</td>
			<td>A31572</td>
			<td>Use at dilution: 1:500</td>
		</tr>
		<tr>
			<td>Donkey anti-Rabbit IgG (H+L) Secondary Antibody, Alexa Fluor 488 conjugate</td>
			<td>Life Technologies</td>
			<td>A21206</td>
			<td>Use at dilution: 1:500</td>
		</tr>
		<tr>
			<td>Donkey anti-Rat IgG (H+L) Secondary Antibody, Alexa Fluor 488 conjugate</td>
			<td>Life Technologies</td>
			<td>A21208</td>
			<td>Use at dilution: 1:500</td>
		</tr>
		<tr>
			<td>Donkey anti-Rabbit IgG (H+L) Secondary Antibody, Alexa Fluor 647 conjugate</td>
			<td>Life Technologies</td>
			<td>A31573</td>
			<td>Use at dilution: 1:500</td>
		</tr>
	</tbody>
</table>

using ex vivo upright droplet cultures of whole fetal organs to study developmental processes during mouse organogenesis

Investigating organogenesis in utero is a technically challenging process in placental mammals due to inaccessibility of reagents to embryos that develop within the uterus. A newly developed ex vivo upright droplet culture method provides an attractive alternative to studies performed in utero. The ex vivo droplet culture provides the ability to examine and manipulate cellular interactions and diverse signaling pathways through use of various blocking and activating compounds; additionally, the effects of various pharmacological reagents on the development of specific organs can be studied without unwanted side effects of systemic drug delivery in utero. As compared to other in vitro systems, the droplet culture not only allows for the ability to study three-dimensional morphogenesis and cell-cell interactions, which cannot be reproduced in mammalian cell lines, but also requires significantly less reagents than other ex vivo and in vitro protocols. This paper demonstrates proper mouse fetal organ dissection and upright droplet culture techniques, followed by whole organ immunofluorescence to demonstrate the effectiveness of the method. The ex vivo droplet culture method allows the formation of organ architecture comparable to what is observed in vivo and can be utilized to study otherwise difficult-to-study processes due to embryonic lethality in in vivo models. As a model application system, a small-molecule inhibitor will be utilized to probe the role of vascularization in testicular morphogenesis. This ex vivo droplet culture method is expandable to other fetal organ systems, such as lung and potentially others, although each organ must be extensively studied to determine any organ-specific modifications to the protocol. This organ culture system provides flexibility in experimentation with fetal organs, and results obtained using this technique will help researchers gain insights into fetal development.

The ex vivo droplet culture allows one to manipulate whole organs, such as the gonad, to study cellular interactions and dynamics. Figure 1 demonstrates in a step-wise fashion how to prepare an E11.5 gonadal droplet culture. The first steps in the culture protocol include the initial removal of the embryo-containing uterus from the mother mouse (Figure 1A and 1B). After removal of the uterus from the mother, the uterine wall is cut and the embryos are liberated from the yolk sac...

Watch this Scientific Journal Video about Using Ex Vivo Upright Droplet Cultures of Whole Fetal Organs to Study Developmental Processes during Mouse Organogenesis at JoVE.com

Using Ex Vivo Upright Droplet Cultures of Whole Fetal Organs to Study Developmental Processes during Mouse Organogenesis

The&#160;ex vivo&#160;upright droplet culture is an alternative to current&#160;in vitro&#160;and&#160;in vivo&#160;experimental techniques. This protocol is easy to perform and requires smaller amounts of reagent, while permitting the ability to manipulate and study fetal vascularization, morphogenesis, and organogenesis.

Using Ex Vivo Upright Droplet Cultures of Whole Fetal Organs to Study Developmental Processes during Mouse Organogenesis

Investigating organogenesis in utero is a technically challenging process in placental mammals due to inaccessibility of reagents to embryos that develop ...

using-ex-vivo-upright-droplet-cultures-whole-fetal-organs-to-study

Research

JoVE Journal

Developmental Biology

12.4K Views.  Cincinnati Children's Hospital Medical Center. The ex vivo upright droplet culture is an alternative to current in vitro and in vivo experimental techniques. This protocol is easy to perform and requires smaller amounts of reagent, while permitting the ability to manipulate and study fetal vascularization, morphogenesis, and organogenesis.

Video: Using Ex Vivo Upright Droplet Cultures of Whole Fetal Organs to Study Developmental Processes during Mouse Organogenesis

Ex Vivo Culture of Pharyngeal Arches to Study Heart and Muscle Progenitors and Their Niche

The pharyngeal mesoderm of developing embryos contributes to broad regions of head and heart musculature. We have developed a novel method to study head and heart progenitor cell development with pharyngeal arches (also known as branchial arches) ex vivo. Using this method, we have recently described that the second pharyngeal arch contains self-renewing heart progenitors and serves as a microenvironment for expansion of the progenitors during mouse heart development. The progenitor cells remain undifferentiated and expansive inside the arch, but quickly become functional cardiomyocytes as they migrate out of the arch. We also reported that first pharyngeal arch contains muscle progenitors giving rise to myotubes after leaving the arch. Here, we demonstrate the procedure for the dissection and ex vivo culture of first and second pharyngeal arches from developing mouse embryos. The method enables one to study head and heart progenitor/muscle development, including cardiomyocyte and myotube formation in detail ex vivo.

Ex Vivo Culture of Pharyngeal Arches to Study Heart and Muscle Progenitors and Their Niche

Here, we present a protocol to culture pharyngeal arches to study the biology of heart and muscle progenitor cells and their microenvironment.

The pharyngeal mesoderm of developing embryos contributes to broad regions of head and heart musculature. We have developed a novel method to study head ...

Using Primary Neurosphere Cultures to Study Primary Cilia

The primary cilium is fundamentally important for the proliferation of neural stem/progenitor cells and for neuronal differentiation during embryonic, postnatal, and adult life. In addition, most differentiated neurons possess primary cilia that house signaling receptors, such as G-protein-coupled receptors, and signaling molecules, such as adenylyl cyclases. The primary cilium determines the activity of multiple developmental pathways, including the sonic hedgehog pathway during embryonic neuronal development, and also functions in promoting compartmentalized subcellular signaling during adult neuronal function. Unsurprisingly, defects in primary cilium biogenesis and function have been linked to developmental anomalies of the brain, central obesity, and learning and memory deficits. Thus, it is imperative to study primary cilium biogenesis and ciliary trafficking in the context of neural stem/progenitor cells and differentiated neurons. However, culturing methods for primary neurons require considerable expertise and are not amenable to freeze-thaw cycles. In this protocol, we discuss culturing methods for mixed populations of neural stem/progenitor cells using primary neurospheres. The neurosphere-based culturing methods provide the combined benefits of studying primary neural stem/progenitor cells: amenability to multiple passages and freeze-thaw cycles, differentiation potential into neurons/glia, and transfectability. Importantly, we determined that neurosphere-derived neural stem/progenitor cells and differentiated neurons are ciliated in culture and localize signaling molecules relevant to ciliary function in these compartments. Utilizing these cultures, we further describe methods to study ciliogenesis and ciliary trafficking in neural stem/progenitor cells and differentiated neurons. These neurosphere-based methods allow us to study cilia-regulated cellular pathways, including G-protein-coupled receptor and sonic hedgehog signaling, in the context of neural stem/progenitor cells and differentiated neurons.

The primary cilium is fundamentally important in neural progenitor cell proliferation, neuronal differentiation, and adult neuronal function. Here, we describe a method to study ciliogenesis and the trafficking of signaling proteins to cilia in neural stem/progenitor cells and differentiated neurons using primary neurosphere cultures.

The primary cilium is fundamentally important for the proliferation of neural stem/progenitor cells and for neuronal differentiation during embryonic, ...

Whole-mount Clearing and Staining of Arabidopsis Flower Organs and Siliques

Due to its formidable tools for molecular genetic studies, Arabidopsis thaliana is one of the most prominent model species in plant biology and, especially, in plant reproductive biology. However, plant morphological, anatomical, and ultrastructural analyses traditionally involve time-consuming embedding and sectioning procedures for bright field, scanning, and electron microscopy. Recent progress in confocal fluorescence microscopy, state-of-the-art 3-D computer-aided microscopic analyses, and the continuous refinement of molecular techniques to be used on minimally processed whole-mount specimens, has led to an increased demand for developing efficient and minimal sample processing techniques. In this protocol, we describe techniques for properly dissecting Arabidopsis flowers and siliques, basic clearing techniques, and some staining procedures for whole-mount observations of reproductive structures.

Whole-mount Clearing and Staining of Arabidopsis Flower Organs and Siliques

In this protocol, we describe techniques for the proper dissection of Arabidopsis flowers and siliques, some basic clearing techniques, and selected staining procedures for whole-mount observations of reproductive structures.

Due to its formidable tools for molecular genetic studies, Arabidopsis thaliana is one of the most prominent model species in plant biology and, ...

Fetal Mouse Cardiovascular Imaging Using a High-frequency Ultrasound (30/45MHZ) System

Congenital heart defects (CHDs) are the most common cause of childhood morbidity and early mortality. Prenatal detection&#160;of the underlying molecular mechanisms of CHDs is&#160;crucial for inventing new preventive and therapeutic strategies. Mutant mouse models are powerful tools to discover new&#160;mechanisms&#160;and environmental stress modifiers that drive cardiac development and their potential&#160;alteration in CHDs. However, efforts to establish the causality of these putative contributors have been limited to histological&#160;and molecular studies&#160;in non-survival animal experiments, in which&#160;monitoring the key physiological and hemodynamic parameters is&#160;often absent. Live imaging technology has become an essential tool to establish the&#160;etiology of CHDs. In particular,&#160;ultrasound imaging can be used prenatally without surgically exposing the fetuses,&#160;allowing maintaining&#160;their baseline physiology while monitoring the impact of environmental stress&#160;on the hemodynamic and structural aspects of cardiac chamber development. Herein, we use the High-Frequency Ultrasound (30/45)&#160;system to examine the cardiovascular system in fetal mice at E18.5 in utero at the baseline and in response to prenatal hypoxia exposure. We demonstrate the feasibility of the system to measure cardiac chamber size, morphology, ventricular function, fetal heart rate, and umbilical artery flow indices, and their alterations in fetal mice exposed to systemic chronic hypoxia in utero in real time.

Fetal Mouse Cardiovascular Imaging Using a High-frequency Ultrasound 30/45MHZ System

High-frequency ultrasound imaging of the fetal mouse has improved imaging resolution and can provide precise non-invasive characterization of cardiac development and structural defects. The protocol outlined herein is designed to perform real-time fetal mice echocardiography in vivo.

Congenital heart defects (CHDs) are the most common cause of childhood morbidity and early mortality. Prenatal detection&#160;of the underlying molecular ...

Three-dimensional Organotypic Cultures of Vestibular and Auditory Sensory Organs

The sensory organs of the inner ear are challenging to study in mammals due to their inaccessibility to experimental manipulation and optical observation. Moreover, although existing culture techniques allow biochemical perturbations, these methods do not provide a means to study the effects of mechanical force and tissue stiffness during development of the inner ear sensory organs. Here we describe a method for three-dimensional organotypic culture of the intact murine utricle and cochlea that overcomes these limitations. The technique for adjustment of a three-dimensional matrix stiffness described here permits manipulation of the elastic force opposing tissue growth. This method can therefore be used to study the role of mechanical forces during inner ear development. Additionally, the cultures permit virus-mediated gene delivery, which can be used for gain- and loss-of-function experiments. This culture method preserves innate hair cells and supporting cells and serves as a potentially superior alternative to the traditional two-dimensional culture of vestibular and auditory sensory organs.

Three-dimensional organotypic cultures of the murine utricle and cochlea in optically clear collagen I gels preserve innate tissue morphology, allow for mechanical stimulation through adjustment of matrix stiffness, and permit virus-mediated gene delivery.

The sensory organs of the inner ear are challenging to study in mammals due to their inaccessibility to experimental manipulation and optical observation. ...

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

Spermatogenesis is the development of spermatogonia into mature spermatozoa in the seminiferous tubules of the testis. This process is supported by Sertoli cell junctions at the blood-testis barrier (BTB), which is the tightest tissue barrier in the mammalian body and segregates the seminiferous epithelium into two compartments, a basal and an adluminal. The BTB creates a unique microenvironment for germ cells in meiosis I/II and for the development of postmeiotic spermatids into spermatozoa via spermiogenesis. Here, we describe a reliable assay to monitor BTB integrity of mouse testis in vivo. An intact BTB blocks the diffusion of FITC-conjugated inulin from the basal to the apical compartment of the seminiferous tubules. This technique is suitable for studying gene candidates, viruses, or environmental toxicants that may affect BTB function or integrity, with an easy procedure and a minimal requirement of surgical skills compared to alternative methods.

An In Vivo Method to Study Mouse Blood-Testis Barrier Integrity

Here, we present a protocol to assess the blood-testis barrier integrity by injecting inulin-FITC into testes. This is an efficient in vivo method to study blood-testis barrier integrity that can be compromised by genetic and environmental elements.

Spermatogenesis is the development of spermatogonia into mature spermatozoa in the seminiferous tubules of the testis. This process is supported by ...

Analysis of Cardiac Chamber Development During Mouse Embryogenesis Using Whole Mount Epifluorescence

The goal of this protocol is to describe a method for the dissection of mouse embryos and visualization of embryonic mouse ventricular chambers during heart development using ventricular specific fluorescent reporter knock-in mice (MLC-2v-tdTomato mice). Heart development involves a linear heart tube formation, the heart tube looping, and four chamber septation. These complex processes are highly conserved in all vertebrates. The mouse embryonic heart has been widely used for heart developmental studies. However, due to their extremely small size, dissecting mouse embryonic hearts is technically challenging. In addition, visualization of cardiac chamber formation often needs in situ hybridization, beta-galactosidase staining using LacZ reporter mice, or immunostaining of sectioned embryonic hearts. Here, we describe how to dissect mouse embryonic hearts and directly visualize ventricular chamber formation of MLC-2v-tdTomato mice using whole mount epifluorescent microscopy. With this method, it is possible to directly examine heart tube formation and looping, and four chamber formation without further experimental manipulation of mouse embryos. Although the MLC-2v-tdTomato reporter knock-in mouse line is used in this protocol as an example, this protocol can be applied to other heart-specific fluorescent reporter transgenic mouse lines.

We present the protocols to examine mouse heart development using whole mount epifluorescent microscopy on mouse embryos dissected from ventricular specific MLC-2v-tdTomato reporter knock-in mice. This method allows us to directly visualize each stage of the ventricular formation during mouse heart development without labor-intensive histochemical methods.

The goal of this protocol is to describe a method for the dissection of mouse embryos and visualization of embryonic mouse ventricular chambers during ...

Human Ex vivo Wound Model and Whole-Mount Staining Approach to Accurately Evaluate Skin Repair

Chronic non-healing wounds, which primarily affect the elderly and diabetic, are a significant area of clinical unmet need. Unfortunately, current chronic wound treatments are inadequate, while available pre-clinical models poorly predict the clinical efficacy of new therapies. Here we describe a high throughput, pre-clinical model to assess multiple aspects of the human skin repair response. Partial thickness wounds were created in human ex vivo skin and cultured across a healing time course. Skin wound biopsies were collected in fixative for the whole-mount staining procedure. Fixed samples were blocked and incubated in primary antibody, with detection achieved via fluorescently conjugated secondary antibody. Wounds were counterstained and imaged via confocal microscopy before calculating percentage wound closure (re-epithelialization) in each biopsy. Applying this protocol, we reveal that 2 mm excisional wounds created in healthy donor skin are fully re-epithelialized by day 4-5 post-wounding. On the contrary, closure rates of diabetic skin wounds are significantly reduced, accompanied by perturbed barrier reformation. Combining human skin wounding with a novel whole-mount staining approach allows a rapid and reproducible method to quantify ex vivo wound repair. Collectively, this protocol provides a valuable human platform to evaluate the effectiveness of potential wound therapies, transforming pre-clinical testing and validation.

Here we demonstrate an optimized technique for assessing wound repair using ex vivo human skin combined with a whole-mount staining approach. This methodology provides a pre-clinical platform for the evaluation of potential wound therapies.

Chronic non-healing wounds, which primarily affect the elderly and diabetic, are a significant area of clinical unmet need. Unfortunately, current chronic ...

Ex Utero Culture of Mouse Embryos from Pregastrulation to Advanced Organogenesis

Postimplantation mammalian embryo culture methods have been generally inefficient and limited to brief periods after dissection out of the uterus. Platforms have been recently developed for highly robust and prolonged ex utero culture of mouse embryos from egg-cylinder stages until advanced organogenesis. These platforms enable appropriate and faithful development of pregastrulating embryos (E5.5) until the hind limb formation stage (E11). Late gastrulating embryos (E7.5) are grown in rotating bottles in these settings, while extended culture from pregastrulation stages (E5.5 or E6.5) requires a combination of static and rotating bottle cultures. In addition, sensitive regulation of O2 and CO2 concentration, gas pressure, glucose levels, and the use of a specific ex utero culture medium are critical for proper embryo development. Here, a detailed step-by-step protocol for extended ex utero mouse embryo culture is provided. The ability to grow normal mouse embryos ex utero from gastrulation to organogenesis represents a valuable tool for characterizing the effect of different experimental perturbations during embryonic development.

Ex Utero Culture of Mouse Embryos from Pregastrulation to Advanced Organogenesis

An enhanced platform for whole-embryo culture allows continuous and robust ex utero development of postimplantation mouse embryos for up to six days, from pregastrulation stages until advanced organogenesis. In this protocol, we detail the standard procedure for successful embryo culture using static plates and rotating bottle systems.

Postimplantation mammalian embryo culture methods have been generally inefficient and limited to brief periods after dissection out of the uterus. ...

Isolation of Whole Cell Protein Lysates from Mouse Facial Processes and Cultured Palatal Mesenchyme Cells for Phosphoprotein Analysis

Mammalian craniofacial development is a complex morphological process during which multiple cell populations coordinate to generate the frontonasal skeleton. These morphological changes are initiated and sustained through diverse signaling interactions, which often include protein phosphorylation by kinases. Here, two examples of physiologically-relevant contexts in which to study phosphorylation of proteins during mammalian craniofacial development are provided: mouse facial processes, in particular E11.5 maxillary processes, and cultured mouse embryonic palatal mesenchyme cells derived from E13.5 secondary palatal shelves. To overcome the common barrier of dephosphorylation during protein isolation, adaptations and modifications to standard laboratory methods that allow for isolation of phosphoproteins are discussed. Additionally, best practices are provided for proper analysis and quantification of phosphoproteins following western blotting of whole cell protein lysates. These techniques, particularly in combination with pharmacological inhibitors and/or murine genetic models, can be used to gain greater insight into the dynamics and roles of various phosphoproteins active during craniofacial development.

The protocol presents a method for isolating whole cell protein lysates from dissected mouse embryo facial processes or cultured mouse embryonic palatal mesenchyme cells and performing subsequent western blotting to assess phosphorylated protein levels.

Mammalian craniofacial development is a complex morphological process during which multiple cell populations coordinate to generate the frontonasal ...