Ex Vivo Perfusion of the Rodent Placenta

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.

1. Modify the vessel chamber.
   NOTE: Figure 1A,B shows the modified single vessel chamber.
   1. Move the thermistor sensor from beneath the clip and bend it so that it hangs freely in the perfusion bath (Figure 1B).
   2. 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.
2. Install two 70−100 µm glass micropipettes. For further information on these procedures, please review the following references^10,11.
   NOTE: The tip diameter commonly used in these experiments is in the range of 70−100 µm. Tip diameters larger than this range may add difficulty to the cannulation process while tip diameters smaller than 60 µm may puncture the vascular wall during cannulation.
3. 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.
4. 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 ± 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.
5. Label microcentrifuge tubes for collecting “maternal” and “fetal” effluents at each time point, i.e., “MBaseline, M10, M20, …” up to the 180 min timepoint. Prepare the same for fetal effluents (FBaseline, M10, M20, ….).
6. 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.
7. Prepare a dissection chamber (4” diameter x 1” deep) or a circulating heater dish by filling a layer (< 0.25”) of silicone rubber. This modification must be done in advance and as it will take 12−24 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.

2. Preparation of the surgical station and equilibration of equipment

1. 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.
2. 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.
3. 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.
4. 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 “off” facing the direction of the pipette to secure the fluid within the pipette.
5. 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).

3. Placental harvesting

1. 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%−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.
2. 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.
3. Using 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°C) PSS. Regardless of whether the right or left horn is chosen, maintain that same side for selection in each experiment for consistency.
   NOTE: 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: 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.
4. 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.
5. Using fine forceps and scissors, remove the amniotic membrane from the fetal surface of the placenta, taking care to avoid the umbilical cord.
6. Unravel and ligate the umbilical cord to separate the fetal pup.
7. 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.
8. Gently separate the umbilical artery and vein from each other.
9. The entire placental unit including the uterine vasculature, uterine muscle, placenta, and umbilical cord, may be cut and removed. 

4. Placental perfusion

1. 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.
2. Using a pair of fine forceps in each hand, cannulate the proximal and distal ends of the uterine artery onto the glass micropipettes.
3. 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.
4. Cannulate the umbilical artery (fetal-to-maternal blood flow) onto the 23 G (larger) needle and secure it with black braided silk suture.
5. Cannulate the umbilical vein (maternal-to-fetal blood flow) onto the 25 G (smaller) blunt needle and secure it with black braided silk suture.
6. Move to the placental perfusion station and backfill all stopcocks and tubings to prevent air bubbles. Connect all tubings according to Figure 2.
7. 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.
8. 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.
9. 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.
10. 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.
11. 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. 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.

5. Mock experiment

1. 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.
2. 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.
3. Initiate sample collection at 10-minute intervals by collecting the effluents from the distal uterine artery and umbilical vein. For example, administer a 900 µL 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).
4. 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.

6. Cleaning the equipment

1. 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.
2. 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.