This work was supported by the National Institute of Environmental Health Sciences (R00-ES024783), Rutgers Center for Environmental Exposures and Disease (P30-ES005022), and Rutgers Joint Graduate Program in Toxicology (T32-ES007148). We would also like to thank Drs. Michael Goedken, Marianne Polunas, and Pedro Louro for their technical expertise and Dr. Adam Goodwill for his assistance in designing our perfusion schematic (Figure 5).

All experimental procedures were approved by the Institutional Animal Care and Use Committee of Rutgers University.
1. Preparation before the experiment
NOTE: These steps may be performed within days/weeks prior to the experiment.
<ol>
	<li>Modify the vessel chamber. 
	NOTE: Figure 1A,B shows the modified single vessel chamber.
	<ol>
		<li>Move the thermistor sensor from beneath the clip and bend it so that it hangs freely in the perfusion bath (Figure 1B).</li>
		<li>Install two 4-inch blunt-tip stainless steel needles with standard luer connection hubs, one 25 G and one 23 G secured beneath the thermistor clip and add a 3-way stopcock to allow for cannulation of the umbilical vasculature (Figure 1B). 
		NOTE: The luer connections of different gauge sizes are color coded. This facilitates easy identification of vessels during and after the experiment.</li>
	</ol>
	</li>
	<li>Install two 70&#8722;100 &#181;m glass micropipettes. For further information on these procedures, please review the following references10,11. 
	NOTE: The tip diameter commonly used in these experiments is in the range of 70&#8722;100 &#181;m. Tip diameters larger than this range may add difficulty to the cannulation process while tip diameters smaller than 60 &#181;m may puncture the vascular wall during cannulation.</li>
	<li>Prepare ties from sterile nylon suture (for proximal and distal ends of the uterine artery) and from black braided silk non-sterile suture (for the umbilical vessels). Make single ties by looping suture as to initiate a square knot or tie a shoe, as described previously11. 
	NOTE: Single ties are commonly made and kept in a Petri dish filled with a small layer of silicone rubber to help work on a sticky background.</li>
	<li>Prepare 2 L of physiological salt solution (PSS) containing, in mmol/L, 129.8 NaCl, 5.4 KCl, 0.5 NaH2PO4, 0.83 MgSO4, 19 NaHCO3, 1.8 CaCl2, and 5.5 glucose. Adjust the pH to 7.40 &#177; 0.02 by slowly adding a few drops of acid (1 M hydrochloric acid) or base (1 N sodium hydroxide) to the solution and wait at least 20 s before reading the adjusted pH measurement. Store PSS until ready for use in the refrigerator to reduce contamination but do not store for more than 2 weeks.</li>
	<li>Label microcentrifuge tubes for collecting &#8220;maternal&#8221; and &#8220;fetal&#8221; effluents at each time point, i.e., &#8220;MBaseline, M10, M20, &#8230;&#8221; up to the 180 min timepoint. Prepare the same for fetal effluents (FBaseline, M10, M20, &#8230;.).</li>
	<li>Calibrate all equipment used within the system, including the circulating bath temperature and blood pressure monitors associated with maintaining uterine (80 mmHg) and umbilical (50 mmHg) perfusate pressures.</li>
	<li>Prepare a dissection chamber (4&#8221; diameter x 1&#8221; deep) or a circulating heater dish by filling a layer (&#60; 0.25&#8221;) of silicone rubber. This modification must be done in advance and as it will take 12&#8722;24 h for the rubber to dry. 
	NOTE: Use of a recirculating heater/chilling dish is recommended for novice surgeons, as the dish will not move and the tissue will maintain a consistent temperature.</li>
</ol>
2. Preparation of the surgical station and equilibration of equipment
<ol>
	<li>Turn on all equipment that supports the perfusion system to check for proper function. Remove the PSS from refrigeration and warm to room temperature. The PSS will be used as both a perfusate and a superfusate.</li>
	<li>Place a small bubbling stone to deliver a gas mixture into the superfusate reservoir. Commonly used mixtures include 21% O2, 8% O2, 3% O2, and 0% O2. Turn on the gas to provide small bubbles to the superfusate solution. Adjust the delivery of gas to avoid splashing.</li>
	<li>Arrange the animal dissecting station by checking anesthetic, arranging surgical equipment, and preparing suture ties. Prepare the dissecting chamber by collecting dissecting pins, turning on the chiller (or retrieving ice), and filling the dissecting chamber (lined with rubber silicone) with cold PSS.</li>
	<li>Gently fill all chambers, needles, glass micropipettes, tubing, and reservoirs with warmed PSS, carefully watching for and eliminating air bubbles by suctioning them out with a fine tip transfer pipette. Turn all 3-way stopcocks with &#8220;off&#8221; facing the direction of the pipette to secure the fluid within the pipette.</li>
	<li>Place a single tie on each of the two glass micropipettes prepared for uterine cannulation and on the two blunt tip needles designated for umbilical cannulation. Secure ties to the pipettes and blunt tip needles in order to prevent loss during chamber movements (Figure 1C).</li>
</ol>
3. Placental harvesting
<ol>
	<li>Anesthetize a pregnant female rat on gestational day 20 with 5% isoflurane for about 4 min or until the animal exhibits labored breathing. Move the animal to the nose cone and administer 2.5%&#8722;3% isoflurane to maintain anesthesia. Confirm unconsciousness by a lack of toe pinch reflex. 
	NOTE: The use of a rat at gestational day 20 is presented in this protocol. However, the protocol remains the same if experimental conditions require placental evaluation to take place earlier in gestation.</li>
	<li>Identify and isolate the uterine horn (right or left) of choice by lifting out and spreading out long-ways outside of the rat carcass. Using a braided silk suture, tie off the uterine artery at the ovary end and vaginal end of the horn. Include the ovary inside of the suture with the uterine horn.</li>
	<li>Using&#160;surgical scissors, excise away the uterine horn by making cuts on the proximal side of the ovary tie and distal side of the vaginal tie, leaving the uterine horn tied off with the sutures on both ends. Transfer the uterine horn into the dissecting dish lined with silicone rubber and filled with cold (4&#176;C) PSS. Regardless of whether the right or left horn is chosen, maintain that same side for selection in each experiment for consistency. 
	NOTE:&#160;For fetal anesthesia, pups are kept in ice-cold PSS. Therefore, PSS temperature within the dissecting chamber is maintained by chilled circulating bath or dissecting chamber should be kept on ice. 
	NOTE:&#160;At this point anesthetized dam may be euthanized per laboratory IACUC protocol approval. In this case, euthanasia occurs via pneumothorax (by cutting the diaphragm) and removal of the maternal heart.</li>
	<li>Gently push a dissecting pin through the uterine horn into the silicone rubber, with the ovary side to the left and vaginal side to the right to visualize the uterine vasculature. This will stabilize the uterus and prevent movement of the tissue during dissection of the fetal compartment. Select a maternal-placenta-fetal unit central to the horn and ligate the uterine artery and vein with surgical scissors. Cut the uterine muscle proximal and distal to the selected placenta and fetus. The uterine muscle can be retracted by pulling it to the side; it is important to leave it intact but pull it away from covering the fetal pup. 
	NOTE: Selection of the maternal-placenta-fetal unit should be based on the length of the uterine artery segment on either side of the arcuate artery. Longer segments will permit more successful cannulation.</li>
	<li>Using fine forceps and scissors, remove the amniotic membrane from the fetal surface of the placenta, taking care to avoid the umbilical cord.</li>
	<li>Unravel and ligate the umbilical cord to separate the fetal pup.</li>
	<li>Identify the umbilical artery (thicker vessel) and vein (thinner vessel). Mark the umbilical vein for easy identification by cutting it slightly shorter than the umbilical artery.</li>
	<li>Gently separate the umbilical artery and vein from each other.</li>
	<li>The entire placental unit including the uterine vasculature, uterine muscle, placenta, and umbilical cord, may be cut and removed.&#160;</li>
</ol>
4. Placental perfusion
<ol>
	<li>Maintain anatomical blood flow retaining correct distal-to-proximal orientation of uterine artery, place the placental unit (composed of the uterine vasculature, uterine muscle, placenta, and umbilical cord) into the modified isolated vessel chamber filled with warm, oxygenated PSS.</li>
	<li>Using a pair of fine forceps in each hand, cannulate the proximal and distal ends of the uterine artery onto the glass micropipettes.</li>
	<li>Tightly secure the uterine artery using the sterile nylon suture tie previously secured onto the micropipette. 
	NOTE: Two-tie loops (knot) may be necessary; however, a single tie loop allows for greater adjustment.</li>
	<li>Cannulate the umbilical artery (fetal-to-maternal blood flow) onto the 23 G (larger) needle and secure it with black braided silk suture.</li>
	<li>Cannulate the umbilical vein (maternal-to-fetal blood flow) onto the 25 G (smaller) blunt needle and secure it with black braided silk suture.</li>
	<li>Move to the placental perfusion station and backfill all stopcocks and tubings to prevent air bubbles. Connect all tubings according to Figure 2.</li>
	<li>Place small weighing boats under the distal cannulation of maternal uterine artery and the needle cannulation of fetal umbilical vein to catch the effluent that will emerge during the procedure.</li>
	<li>Turn on the peristaltic pump, pressure controller, and pressure monitor and open the stopcock to permit fluid flow through the tubing toward the pressure transducer, but not yet to the placenta. Slowly increase the pressure to 80 mm Hg.</li>
	<li>Slowly turn the stopcock to the placenta to open and watch for leaks inside the chamber and around the ties. 
	NOTE: If the peristaltic pump is running at a high speed, re-evaluate the proximal uterine cannulation and ties for fluid leaks. If identified, leaks must be remedied. Also note, while the mean uterine artery pressure will remain constant (set to 80 mmHg), the fluid flow rate may be variable depending on the vascular and placental physiological responses. Quantification of the fluid flow may be identified as an experimental variable in response to xenobiotic exposure.</li>
	<li>Turn the stopcock to permit fluid flow into the umbilical artery. Set the pressure to approximately 50 mmHg, which can be implemented with a peristaltic pump (Figure 3) or a hydrostatic column.</li>
	<li>Refill all reservoirs (uterine, umbilical, and superfusate) to maintain fluid volume throughout the experiment. 
	NOTE: Once perfusion has been initiated and experiment is underway, anesthetized pups remaining in the dissecting dish may be assessed for viability by touching the pups with forceps to elicit a neurological response.&#160;Exsanguination of the pups will occur through hysterectomy of uterine horn; further euthanasia can be completed through approved IACUC protocols. In this case, opening the amniotic sac in the cold PSS and creating a pneumothorax by cutting the ribs.</li>
</ol>
5. Mock experiment
<ol>
	<li>After cannulation and initiation of the PSS perfusion, let tissues equilibrate for 30 min to allow the vasculature to adjust to the new fluid flow. Suction up the effluent by a pipette and save it in a correspondingly labeled microcentrifuge tube.</li>
	<li>After equilibration, establish the baseline perfusion by collecting effluents for 10 min from both weighing boats in microcentrifuge tubes and measuring the volume of fluid that emerges through the uterine artery and umbilical vein.</li>
	<li>Initiate sample collection at 10-minute intervals by collecting the effluents from the distal uterine artery and umbilical vein. For example, administer a 900 &#181;L bolus dose of 20 nm rhodamine-labeled polystyrene nanoparticles (8 x 1014 particles/mL) suspended in 0.01% surfactant to the line perfusing the uterine artery to identify the time course passage of xenobiotic nanoparticles across the placental barrier. 
	NOTE: These effluent samples will allow for the measurement of contaminants within the fluid (either xenobiotic, pharmacologic, or metabolite) and provide the rate of fluid flow through the uterine artery or across the placenta and into the fetal compartment (Figure 4).</li>
	<li>After bolus infusion, collect samples from the distal uterine artery and fetal umbilical vein effluent every 10 min for a total of 180 min after infusion.</li>
</ol>
6. Cleaning the equipment
<ol>
	<li>After each experiment, remove the placental unit from the pipettes. All sutures may be saved to be reused in future experiments. The placental tissue may be saved for additional histological or mechanistic studies.</li>
	<li>Clean all tubings and cannulas of the perfusion system with 70% ethanol followed by distilled water and vacuum dry. 
	NOTE: If tubings or pipettes begin to discolor or appear damaged, replace prior to the next experiment. Further, after an experimental cohort is complete (e.g., all studies pertaining to a single contaminant), replace all tubings within the chamber to prevent cross contamination.</li>
</ol>

<ol>
	<li>Troen, P., Gordon, E. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Perfusion+studies+of+the+human+placenta.+I.+Effect+of+estradiol+and+human+chorionic+gonadotropin+on+citric+acid+metabolism.">Perfusion studies of the human placenta. I. Effect of estradiol and human chorionic gonadotropin on citric acid metabolism.</a> Journal of Clinical Investigation. 37, 1516-1523 (1958).</li><li>Alexander, D. P., Huggett, A. S., Nixon, D. A., Widdas, W. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+placental+transfer+of+sugars+in+the+sheep:+the+influence+of+concentration+gradient+upon+the+rates+of+hexose+formation+as+shown+in+umbilical+perfusion+of+the+placenta.">The placental transfer of sugars in the sheep: the influence of concentration gradient upon the rates of hexose formation as shown in umbilical perfusion of the placenta.</a> Journal of Physiology. 129, 367-383 (1955).</li><li>Alexander, D. P., Andrews, R. D., Huggett, A. S., Nixon, D. A., Widdas, W. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+placental+transfer+of+sugars+in+the+sheep:+studies+with+radioactive+sugar.">The placental transfer of sugars in the sheep: studies with radioactive sugar.</a> Journal of Physiology. 129, 352-366 (1955).</li><li>Dancis, J., Money, W. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transfer+of+sodium+and+iodo-antipyrine+across+guinea+pig+placenta+with+an+in+situ+perfusion+technique.">Transfer of sodium and iodo-antipyrine across guinea pig placenta with an in situ perfusion technique.</a> American Journal of Obstetrics and Gynecology. 80, 215-220 (1960).</li><li>London, W. T., Money, W. L., Rawson, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Placental+Transport+of+I-131-Labeled+Thyroxine+and+Triiodothyronine+in+the+Guinea+Pig.">Placental Transport of I-131-Labeled Thyroxine and Triiodothyronine in the Guinea Pig.</a> Endocrinology. 73, 205-209 (1963).</li><li>Stulc, J., Stulcova, B., Svihovec, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transport+of+calcium+across+the+dually+perfused+placenta+of+the+rat.">Transport of calcium across the dually perfused placenta of the rat.</a> Journal of Physiology. 420, 295-311 (1990).</li><li>Goeden, N., Bonnin, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ex+vivo+perfusion+of+mid-to-late-gestation+mouse+placenta+for+maternal-fetal+interaction+studies+during+pregnancy.">Ex vivo perfusion of mid-to-late-gestation mouse placenta for maternal-fetal interaction studies during pregnancy.</a> Nature Protocols. 8, 66-74 (2013).</li><li>Bond, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Artificial+perfusion+of+the+fetal+circulation+of+the+in+situ+mouse+placenta:+methodology+and+validation.">Artificial perfusion of the fetal circulation of the in situ mouse placenta: methodology and validation.</a> Placenta. 27, (2006).</li><li>Illsley, N. P., Fox, H., Van der Veen, F., Chawner, L., Penfold, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+placental+ultrastructure+after+in+vitro+dual+perfusion.">Human placental ultrastructure after in vitro dual perfusion.</a> Placenta. 6, 23-32 (1985).</li><li>Davis, M. J., Kuo, L., Chilian, W. M., Muller, J. M., Barker, J. H., Anderson, G. L., Menger, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolated,+Perfused+Microvessels.">Isolated, Perfused Microvessels.</a> Clinically Applied Microcirculation Research. , 435-456 (1995).</li><li>Butcher, J. T., Goodwill, A. G., Frisbee, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+ex+vivo+isolated+skeletal+microvessel+preparation+for+investigation+of+vascular+reactivity.">The ex vivo isolated skeletal microvessel preparation for investigation of vascular reactivity.</a> Journal of Visualized Experiments. (62), 3674 (2012).</li></ol>

The authors have nothing to disclose.

This perfusion method allows for rapid assessment of placental barrier and physiological function of the uterine vasculature and trophoblast layer. Cannulation and perfusion of the proximal to distal ends of the maternal uterine artery simulates the physiology of maternal blood flow through this major vessel responsible for sending blood to the developing fetus. This methodology allows for physiological evaluation of the isolated maternal, placental and umbilical vasculature, and therefore changes in physiology may be identified as vascular pathology; the immune and neural innervations are removed in the ex-vivo procedure. To ensure proper evaluation, it is therefore critical to cannulate these vessels carefully as to not create any tears or punctures in the vessel walls, and to remove air bubbles. Gas emboli may cause damage to the endothelial layer of the vasculature or obstruct blood vessels. By maintaining the vascular connections between the uterus, placenta and fetus during dissection, the evaluation of fluid and translocation to the fetus can be observed. With the administration of a xenobiotic, in this case 20 nm polystyrene, kinetics to the distal end of the uterine artery and through the placenta to the fetal compartment can be evaluated by analysis of effluents over a time course of 180 minutes.
While a dual-perfusion model was described and the transfer of particles and fluid from the maternal to the fetal compartment was monitored in this article, assessments can also be made in reverse from the fetal to the maternal compartment. One limitation of the method described here is that the distal uterine vein was not cannulated or sampled. In future studies, especially those focused on the fetal-to-maternal transfer, it will be important to cannulate and sample the distal uterine vessel. The effluents taken from this mock experiment were used to assess xenobiotic transfer; however, a wide array of assessments pertaining to endocrine and molecular placental functions or fetal nutrition may be performed.
The strengths of this protocol far outweigh its minor limitations. The preparation maintains the physiological structure and integrity of a whole organ to assess experimental conditions. Ex vivo placental perfusion is a scientific progression from cellular-based in vitro to whole animal exposure to properly determine reproductive risk assessment. This may be considered to be a valuable technique for studies evaluating placental pharmacologic drug disposition, pharmacokinetics, toxicology, physiology, and maternal-fetal medicine.

By maintaining morphological structure and physiological responsiveness, organ perfusion has been an accepted system- or tissue-based approach for analyzing metabolic function. These perfusion techniques allow for the ex vivo examination of intact tissue responses to a variety of pharmacological and mechanical stimuli. Perfusion of human placenta was initially described in 1958 to identify the hormonal effects on the metabolic activity of the citric acid cycle; having previously been identified in tissue homogenates, Troen and Gordon recognized the need to clarify endocrine activity using a novel physiological approach1. In the same era, single-perfusion (maternal-to-fetal or fetal-to-maternal) strategies were described in large2,3 and small4 animal models to understand the placental transfer of sugars, salts, and antipyrine drugs. In vivo and ex vivo dual-perfusion (coordinated maternal and fetal perfusion) techniques were described to further characterize placental transfer using in vivo5 and ex vivo6,7,8 methodologies. Technological advancements in transmission and scanning electron microscopy allowed researchers to verify the structural and functional integrity of human placental tissues after perfusion9.
While perfusion of human placental tissues and individual cotyledon is most relevant, the rapid development of pharmacological agents and environmental contaminants necessitates the use of an animal perfusion model for the early screening of xenobiotic transfer across the placental barrier. This placental perfusion method allows for the evaluation of transfer across the placental barrier using more easily attainable and physiologically relevant rat placenta. In addition, fluid flow across the placental barrier over a period of time after an exposure can be evaluated by measuring the volume of perfusate coming from the umbilical artery. By virtue of allowing placental perfusion from both the maternal and fetal circulations, this dual-flow whole organ approach can be advantageous compared to current in vitro and in vivo approaches. This method allows the administration of a xenobiotic through the maternal aspect to be measured from perfusate that emerges across the placenta through the umbilical vein, or vice versa. The protocol presented here will describe the transfer of 20 nm polystyrene (a common nanoplastic used in food and medical products) from the maternal uterine artery to the fetal compartment and an associated decrease in fluid flow across the placenta to illustrate the use of this method in multiple physiological, pharmacological, and toxicological settings to assess placental transfer, metabolism, and physiological alterations affecting maternal and/or fetal flow.

<table>
	<tbody>
		<tr>
			<td>Black braided silk non-absorbable surgical suture non-sterile</td>
			<td>Surgical Specialties Look</td>
			<td>AACO805</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine forceps</td>
			<td>FST by Dumont Switzerland</td>
			<td>11252-20</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine scissors</td>
			<td>FST by Dumont Switzerland</td>
			<td>14060-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass cannula pack</td>
			<td>Living Systems Instrumentation (LSI)</td>
			<td>GCP-75-100</td>
			<td></td>
		</tr>
		<tr>
			<td>Microcentrifuge Tubes 2.0mL polypropylene graduated tube with locking lid MIXED</td>
			<td>Fisherbrand</td>
			<td>02-681-299</td>
			<td></td>
		</tr>
		<tr>
			<td>Non-serrated fine curved micro serrefine clamps</td>
			<td>InterFocus</td>
			<td>18052-03</td>
			<td></td>
		</tr>
		<tr>
			<td>Perfusate pump</td>
			<td>ISMATEC</td>
			<td>ISM795C</td>
			<td></td>
		</tr>
		<tr>
			<td>Pressure monitor</td>
			<td>Living Systems Instrumentation (LSI)</td>
			<td>Mode PM-4</td>
			<td></td>
		</tr>
		<tr>
			<td>Self-heating single vessel chamber</td>
			<td>Living Systems Instrumentation (LSI)</td>
			<td>CH-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Servo Pump</td>
			<td>Living Systems Instrumentation (LSI)</td>
			<td>ModelPS-200-P</td>
			<td></td>
		</tr>
		<tr>
			<td>Stainless steel blunt needle 23 gauge</td>
			<td>Component Supply Co.</td>
			<td>04651-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Stainless steel blunt needle 25 gauge</td>
			<td>Component Supply Co.</td>
			<td>07116-01</td>
			<td></td>
		</tr>
		<tr>
			<td>STERILE Nylon Suture</td>
			<td>AROSurgical Instruments Corporation</td>
			<td>T04A00N07-13</td>
			<td></td>
		</tr>
		<tr>
			<td>Stopcock</td>
			<td>Sedation Resource</td>
			<td>6-205-04</td>
			<td></td>
		</tr>
		<tr>
			<td>Temperature Controller</td>
			<td>Living Systems Instrumentation (LSI)</td>
			<td>Model TC-09S</td>
			<td></td>
		</tr>
	</tbody>
</table>

ex vivo perfusion of the rodent placenta

The placenta is a key organ during pregnancy that serves as a barrier to fetal xenobiotic exposure and mediates the exchange of nutrients for waste. An assay is described here to perfuse an isolated rat placenta and evaluate the maternal-to-fetal translocation of xenobiotics ex vivo. In addition, the evaluation of physiological processes such as fluid flow to the fetus and placental metabolism may be conducted with this methodology. This technique is suitable for evaluating maternal-to-fetal kinetics of pharmaceutical candidates or environmental contaminants. In contrast to current alternative approaches, this methodology allows the evaluation of the isolated maternal-fetal vasculature, with the systemic neural or immune involvement removed, allowing any observed changes in physiological function to be attributable to local factors within the isolated tissue.

Figure 5 shows the proof-of-principle experiments using Evan&#8217;s blue dye, allowing us to test the system and visualize appropriate fluid and placental barrier function and to prevent containment transfer to the fetal compartment. The Evan&#8217;s blue dye reached and perfused the tissue of the placenta within this system (Figure 5A). Upon further investigation, it is clear that the Evan&#8217;s blue dye did not enter the fetal umbilical vein (Figure 5B), which is expected as Evan&#8217;s blue dye is bound to albumin.
Figure 6 shows data for the mock experiment described in this protocol. Effluent samples from the distal end of the uterine artery and fetal umbilical vein were measured at each 10-minute segment to evaluate the fluid flow over time after the bolus dose was administered to the maternal uterine artery (Figure 6). Reduced fluid transfer to the fetal compartment within 10 minutes after polystyrene infusion was identified. To quantify the transfer of polystyrene to the fetal compartment during the time course when it occurs, 25 &#181;L of the perfused fluid from each time point was placed in a 96 well plate in duplicate to measure the sample fluorescence. Fluorescence was determined by spectroscopic reading at 546/575 nm (ex/em) using a fluorescent microplate reader. Polystyrene transfer to the fetal compartment occurred within 10 minutes and peaked at 20 minutes and continued for 90 minutes (Figure 6B).
A subset of perfused placental tissues were saved for histopathology and morphological assessments. The tissues were formalin-fixed and hematoxylin and eosin stained and reviewed by a board certified veterinary pathologist. These experts identified no structural abnormalities in placentas perfused by only PSS, or PSS with the bolus dose of rhodamine-labeled polystyrene.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59412/59412fig1.jpg" /> 
Figure 1: The modified single vessel chamber. (A) An overview of the modified chamber. (B) A close-up image of the blunt-tip needles secured within the vessel chamber. The red arrow indicates the thermistor clip that has been altered to hold the needles in place for umbilical cannulation. (C) A representative image of the four cannulas prepared for tissue cannulation. Red arrows point to each of the four cannulas. <a href="https://www.jove.com/files/ftp_upload/59412/59412fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59412/59412fig2.jpg" /> 
Figure 2: A closer view of the placental perfusion chamber. (A) This represents the tubing attached to the pressure transducer and cannulated the proximal maternal uterine artery, or the &#8220;inflow&#8221;. Pressure is set to a constant 80 mmHg as defined by the literature. (B) This represents the chamber drain port of the superfusate surrounding the placental tissue during perfusion. (C) This represents the chamber inflow of the superfusate to bathe the placenta with warmed PSS during perfusion. (D) This represents the distal maternal uterine port where effluent from the uterine perfusion may be collected. (E) This represents the temperature port, where the vessel chamber can be attached to a thermometer and heater to maintain a consistent temperature throughout the experiment. (F) This represents the umbilical artery cannulation. The umbilical artery is pressurized to 50 mmHg to allow for countercurrent flow at the level of the placenta. (G) This represents the umbilical vein effluent collection. Fluid that flows toward the fetal compartment during perfusion will be collected here. (H) This is the center of the perfusion system, where the placenta is cannulated and maintained throughout perfusion. <a href="https://www.jove.com/files/ftp_upload/59412/59412fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59412/59412fig3.jpg" /> 
Figure 3: A view of the placental perfusion system. (A and B) The pressure control system used to monitor and maintain 80 mmHg of perfusate through uterine artery. (C) This represents the thermo-regulation of the perfusion chamber. (D) Microscope. (E) Perfusion chamber. (F) Gravity-fed umbilical artery perfusion set at 50 mmHg. (G) A peristaltic pump used to fill and drain placental superfusate PSS. <a href="https://www.jove.com/files/ftp_upload/59412/59412fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/59412/59412fig4.jpg" /> 
Figure 4: Schematic of the placental perfusion system. <a href="https://www.jove.com/files/ftp_upload/59412/59412fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/59412/59412fig5.jpg" /> 
Figure 5: Representative images of proof-of-principle experiments using Evan&#8217;s blue dye. (A and B) Proof-of-principle that Evan&#8217;s blue will perfuse the uterine vasculature, uterine muscle, and placenta but will not cross the placental barrier due to albumin binding. The green arrow indicates the blue venous drainage from the placenta back to the maternal circulation. The Red arrow indicates the umbilical vein effluent toward the fetal compartment. Note the lack of blue dye. (C) A representative image of collecting effluent draining from the umbilical vein. The red arrow indicates drop formation prior to collection. <a href="https://www.jove.com/files/ftp_upload/59412/59412fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/59412/59412fig6.jpg" /> 
Figure 6: Data derived from the mock experiment. Fluorescence measurements of rhodamine-labeled polystyrene nanomaterials, normalized to baseline fluorescence, through the collection of (A) uterine artery and (B) fetal umbilical vein effluents. Mean normalized to baseline fluorescence &#177; standard error (SE). *: p &#60; 0.05 and T: p &#60; 0.1 via analysis of variance (ANOVA). <a href="https://www.jove.com/files/ftp_upload/59412/59412fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Ex Vivo Perfusion of the Rodent Placenta at JoVE.com

Ex Vivo Perfusion of the Rodent Placenta

Here is presented a protocol of ex vivo maternal-fetal vascular perfusion to enable the administration of a test article into maternal vasculature and to evaluate placental transfer of xenobiotic particles or pharmacological agents in addition to alterations in placental physiology.

The placenta is a key organ during pregnancy that serves as a barrier to fetal xenobiotic exposure and mediates the exchange of nutrients for waste. An ...

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Developmental Biology

9.6K Views.  Rutgers University. Here is presented a protocol of ex vivo maternal-fetal vascular perfusion to enable the administration of a test article into maternal vasculature and to evaluate placental transfer of xenobiotic particles or pharmacological agents in addition to alterations in placental physiology.

Video: Ex Vivo Perfusion of the Rodent Placenta

Efficient Gene Transfer in Chick Retinas for Primary Cell Culture Studies: An Ex-ovo Electroporation Approach

The cone photoreceptor-enriched cultures derived from embryonic chick retinas have become an indispensable tool for researchers around the world studying the biology of retinal neurons, particularly photoreceptors. The applications of this system go beyond basic research, as they can easily be adapted to high throughput technologies for drug development. However, genetic manipulation of retinal photoreceptors in these cultures has proven to be very challenging, posing an important limitation to the usefulness of the system. We have recently developed and validated an ex&#160;ovo plasmid electroporation technique that increases the rate of transfection of retinal cells in these cultures by five-fold compared to other currently available protocols1. In this method embryonic chick eyes are enucleated at stage 27, the RPE is removed, and the retinal cup is placed in a plasmid-containing solution and electroporated using easily constructed custom-made electrodes. The retinas are then dissociated and cultured using standard procedures. This technique can be applied to overexpression studies as well as to the downregulation of gene expression, for example via the use of plasmid-driven RNAi technology, commonly achieving transgene expression in 25% of the photoreceptor population. The video format of the present publication will make this technology easily accessible to researchers in the field, enabling the study of gene function in primary retinal cultures. We have also included detailed explanations of the critical steps of this procedure for a successful outcome and reproducibility.

Efficient Gene Transfer in Chick Retinas for Primary Cell Culture Studies: An Ex-ovo Electroporation Approach

Embryonic chick retinal cell cultures constitute valuable tools for the study of photoreceptor biology. We have developed an efficient gene transfer technique based on ex&#160;ovo plasmid electroporation of the retinas prior to culture. This technique considerably increases transfection efficiencies over currently available protocols, making genetic manipulation feasible for this system.

The cone photoreceptor-enriched cultures derived from embryonic chick retinas have become an indispensable tool for researchers around the world studying ...

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 ...

Establishment of a Clinically Relevant Ex Vivo Mock Cataract Surgery Model for Investigating Epithelial Wound Repair in a Native Microenvironment

The major impediment to understanding how an epithelial tissue executes wound repair is the limited availability of models in which it is possible to follow and manipulate the wound response ex vivo in an environment that closely mimics that of epithelial tissue injury in vivo. This issue was addressed by creating a clinically relevant epithelial ex vivo injury-repair model based on cataract surgery. In this culture model, the response of the lens epithelium to wounding can be followed live in the cells&#8217; native microenvironment, and the molecular mediators of wound repair easily manipulated during the repair process. To prepare the cultures, lenses are removed from the eye and a small incision is made in the anterior of the lens from which the inner mass of lens fiber cells is removed. This procedure creates a circular wound on the posterior lens capsule, the thick basement membrane that surrounds the lens. This wound area where the fiber cells were attached is located just adjacent to a continuous monolayer of lens epithelial cells that remains linked to the lens capsule during the surgical procedure. The wounded epithelium, the cell type from which fiber cells are derived during development, responds to the injury of fiber cell removal by moving collectively across the wound area, led by a population of vimentin-rich repair cells whose mesenchymal progenitors are endogenous to the lens1. These properties are typical of a normal epithelial wound healing response. In this model, as in vivo, wound repair is dependent on signals supplied by the endogenous environment that is uniquely maintained in this ex vivo culture system, providing an ideal opportunity for discovery of the mechanisms that regulate repair of an epithelium following wounding.

Establishment of a Clinically Relevant Ex Vivo Mock Cataract Surgery Model for Investigating Epithelial Wound Repair in a Native Microenvironment

Described here is the establishment of a clinically relevant ex vivo mock cataract surgery model that can be used to investigate mechanisms of the injury response of epithelial tissues within their native microenvironment.

The major impediment to understanding how an epithelial tissue executes wound repair is the limited availability of models in which it is possible to ...

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.

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.

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

Ex Vivo Culture of Chick Cerebellar Slices and Spatially Targeted Electroporation of Granule Cell Precursors

The cerebellar external granule layer (EGL) is the site of the largest transit amplification in the developing brain, and an excellent model for studying neuronal proliferation and differentiation. In addition, evolutionary modifications of its proliferative capability have been responsible for the dramatic expansion of cerebellar size in the amniotes, making the cerebellum an excellent model for evo-devo studies of the vertebrate brain. The constituent cells of the EGL, cerebellar granule progenitors, also represent a significant cell of origin for medulloblastoma, the most prevalent paediatric neuronal tumour. Following transit amplification, granule precursors migrate radially into the internal granular layer of the cerebellum where they represent the largest neuronal population in the mature mammalian brain. In chick, the peak of EGL proliferation occurs towards the end of the second week of gestation. In order to target genetic modification to this layer at the peak of proliferation, we have developed a method for genetic manipulation through ex vivo electroporation of cerebellum slices from embryonic Day 14 chick embryos. This method recapitulates several important aspects of in vivo granule neuron development and will be useful in generating a thorough understanding of cerebellar granule cell proliferation and differentiation, and thus of cerebellum development, evolution and disease.

Ex Vivo Culture of Chick Cerebellar Slices and Spatially Targeted Electroporation of Granule Cell Precursors

The cerebellar external granule layer is the site of the largest transit amplification in the developing brain. Here, we present a protocol to target genetic modification to this layer at the peak of proliferation using ex vivo electroporation and culture of cerebellar slices from embryonic Day 14 chick embryos.

The cerebellar external granule layer (EGL) is the site of the largest transit amplification in the developing brain, and an excellent model for studying ...

Epicardial Outgrowth Culture Assay and Ex Vivo Assessment of Epicardial-derived Cell Migration

A single layer of epicardial cells lines the heart, providing paracrine factors that stimulate cardiomyocyte proliferation and directly contributing cardiovascular progenitors during development and disease. While a number of factors have been implicated in epicardium-derived cell (EPDC) mobilization, the mechanisms governing their subsequent migration and differentiation are poorly understood. Here, we present in vitro and ex vivo strategies to study EPDC motility and differentiation. First, we describe a method of obtaining primary epicardial cells by outgrowth culture from the embryonic mouse heart. We also introduce a detailed protocol to assess three-dimensional migration of labeled EPDC in an organ culture system. We provide evidence using these techniques that genetic deletion of myocardin-related transcription factors in the epicardium attenuates EPDC migration. This approach serves as a platform to evaluate candidate modifiers of EPDC biology and could be used to develop genetic or chemical screens to identify novel regulators of EPDC mobilization that might be useful for cardiac repair.

Epicardial Outgrowth Culture Assay and Ex Vivo Assessment of Epicardial-derived Cell Migration

Here, we describe methods for isolating primary mouse epicardial cells by an outgrowth culture assay and assessing the functional migration of epicardial-derived cells (EPDC) using an ex vivo heart culture system. These protocols are suitable for identifying genetic and chemical modulators of epicardial epithelial-to-mesenchymal transition (EMT) and motility.

A single layer of epicardial cells lines the heart, providing paracrine factors that stimulate cardiomyocyte proliferation and directly contributing ...

Isolation and Expansion of Mesenchymal Stem/Stromal Cells Derived from Human Placenta Tissue

Mesenchymal stem/stromal cells (MSC) are promising candidates for use in cell-based therapies. In most cases, therapeutic response appears to be cell-dose dependent. Human term placenta is rich in MSC and is a physically large tissue that is generally discarded following birth. Placenta is an ideal starting material for the large-scale manufacture of multiple cell doses of allogeneic MSC. The placenta is a fetomaternal organ from which either fetal or maternal tissue can be isolated. This article describes the placental anatomy and procedure to dissect apart the decidua (maternal), chorionic villi (fetal), and chorionic plate (fetal) tissue. The protocol then outlines how to isolate MSC from each dissected tissue region, and provides representative analysis of expanded MSC derived from the respective tissue types. These methods are intended for pre-clinical MSC isolation, but have also been adapted for clinical manufacture of placental MSC for human therapeutic use.

Herein we describe methods for the dissection of fetal and maternal tissues from human term placenta, followed by isolation and expansion of mesenchymal stem/stromal cells (MSC) from these tissues.

Mesenchymal stem/stromal cells (MSC) are promising candidates for use in cell-based therapies. In most cases, therapeutic response appears to be cell-dose ...

An Ex Vivo Method for Time-Lapse Imaging of Cultured Rat Mesenteric Microvascular Networks

Angiogenesis, defined as the growth of new blood vessels from pre-existing vessels, involves endothelial cells, pericytes, smooth muscle cells, immune cells, and the coordination with lymphatic vessels and nerves. The multi-cell, multi-system interactions necessitate the investigation of angiogenesis in a physiologically relevant environment. Thus, while the use of in vitro cell-culture models have provided mechanistic insights, a common critique is that they do not recapitulate the complexity associated with a microvascular network. The objective of this protocol is to demonstrate the ability to make time-lapse comparisons of intact microvascular networks before and after angiogenesis stimulation in cultured rat mesentery tissues. Cultured tissues contain microvascular networks that maintain their hierarchy. Immunohistochemical labeling confirms the presence of endothelial cells, smooth muscle cells, pericytes, blood vessels and lymphatic vessels. In addition, labeling tissues with BSI-lectin enables time-lapse comparison of local network regions before and after serum or growth factor stimulation characterized by increased capillary sprouting and vessel density. In comparison to common cell culture models, this method provides a tool for endothelial cell lineage studies and tissue specific angiogenic drug evaluation in physiologically relevant microvascular networks.

An Ex Vivo Method for Time-Lapse Imaging of Cultured Rat Mesenteric Microvascular Networks

Angiogenesis involves multi-cell, multi-system interactions that need to be investigated in a physiologically relevant environment. The objective of this study is to demonstrate the ability of the rat mesentery culture model to make time-lapse comparisons of intact microvascular networks during angiogenesis.

Angiogenesis, defined as the growth of new blood vessels from pre-existing vessels, involves endothelial cells, pericytes, smooth muscle cells, immune ...

Isolation and Characterization of Mesenchymal Stromal Cells from Human Umbilical Cord and Fetal Placenta

The human umbilical cord (UC) and placenta are non-invasive, primitive and abundant sources of mesenchymal stromal cells (MSCs) that have increasingly gained attention because they do not pose any ethical or moral concerns. Current methods to isolate MSCs from UC yield low amounts of cells with variable proliferation potentials. Since UC is an anatomically-complex organ, differences in MSC properties may be due to the differences in the anatomical regions of their isolation. In this study, we first dissected the cord/placenta samples into three discrete anatomical regions: UC, cord-placenta junction (CPJ), and fetal placenta (FP). Second, two distinct zones, cord lining (CL) and Wharton&#39;s jelly (WJ), were separated. The explant culture technique was then used to isolate cells from the four sources. The time required for the primary culture of cells from the explants varied depending on the source of the tissue. Outgrowth of the cells occurred within 3 - 4 days of the CPJ explants, whereas growth was observed after 7 - 10 days and 11 - 14 days from CL/WJ and FP explants, respectively. The isolated cells were adherent to plastic and displayed fibroblastoid morphology and surface markers, such as CD29, CD44, CD73, CD90, and CD105, similarly to bone marrow (BM)-derived MSCs. However, the colony-forming efficiency of the cells varied, with CPJ-MSCs and WJ-MSCs showing higher efficiency than BM-MSCs. MSCs from all four sources differentiated into adipogenic, chondrogenic, and osteogenic lineages, indicating that they were multipotent. CPJ-MSCs differentiated more efficiently in comparison to other MSC sources. These results suggest that the CPJ is the most potent anatomical region and yields a higher number of cells, with greater proliferation and self-renewal capacities in vitro. In conclusion, the comparative analysis of the MSCs from the four sources indicated that CPJ is a more promising source of MSCs for cell therapy, regenerative medicine, and tissue engineering.

Here, we present a protocol for the dissection of human umbilical cord (UC) and fetal placenta sample into cord lining (CL), Wharton&#39;s jelly (WJ), cord-placenta junction (CPJ), and fetal placenta (FP) for the isolation and characterization of mesenchymal stromal cells (MSCs) using the explant culture technique.

The human umbilical cord (UC) and placenta are non-invasive, primitive and abundant sources of mesenchymal stromal cells (MSCs) that have increasingly ...

Immuno-fluorescent Labeling of Microtubules and Centrosomal Proteins in Ex Vivo Intestinal Tissue and 3D In Vitro Intestinal Organoids

The advent of 3D in vitro organoids that mimic the in vivo tissue architecture and morphogenesis has greatly advanced the ability to study key biological questions in cell and developmental biology. In addition, organoids together with recent technical advances in gene editing and viral gene delivery promises to advance medical research and development of new drugs for treatment of diseases. Organoids grown in vitro in basement matrix provide powerful model systems for studying the behavior and function of various proteins and are well suited for live-imaging of fluorescent-tagged proteins. However, establishing the expression and localization of the endogenous proteins in ex vivo tissue and in in vitro organoids is important to verify the behavior of the tagged proteins. To this end we have developed and modified tissue isolation, fixation, and immuno-labeling protocols for localization of microtubules, centrosomal, and associated proteins in ex vivo intestinal tissue and in in vitro intestinal organoids. The aim was for the fixative to preserve the 3D architecture of the organoids/tissue while also preserving antibody antigenicity and enabling good penetration and clearance of fixative and antibodies. Exposure to cold depolymerizes all but stable microtubules and this was a key factor when modifying the various protocols. We found that increasing the ethylenediaminetetraacetic acid (EDTA) concentration from 3 mM to 30 mM gave efficient detachment of villi and crypts in the small intestine while 3 mM EDTA was sufficient for colonic crypts. The developed formaldehyde/methanol fixation protocol gave very good structural preservation while also preserving antigenicity for effective labeling of microtubules, actin, and the end-binding (EB) proteins. It also worked for the centrosomal protein ninein although the methanol protocol worked more consistently. We further established that fixation and immuno-labeling of microtubules and associated proteins could be achieved with organoids isolated from or remaining within the basement matrix.

Immuno-fluorescent Labeling of Microtubules and Centrosomal Proteins in Ex Vivo Intestinal Tissue and 3D In Vitro Intestinal Organoids

We present protocols for isolation of intestinal 3D structures from in vivo tissue and in vitro basement matrix embedded organoids, and detail different fixation and staining protocols optimized for immuno-labeling of microtubule, centrosomal, and junctional proteins as well as cell markers including the stem cell protein Lgr5.

The advent of 3D in vitro organoids that mimic the in vivo tissue architecture and morphogenesis has greatly advanced the ability to study key biological ...