Intravenous and Intra-amniotic In Utero Transplantation in the Murine Model

The experimental protocols were approved by the Institutional Animal Care and Use Committee at The Children's Hospital of Philadelphia.

1. Creation of Injection Pipettes

1. Using a vertical micropipette puller, pull a 100 µL microcapillary pipette (Figure 1A – 1C). Calibrate the micropipette puller so that the tapered end is > 1 cm long.
   NOTE: Initially, the settings of the puller should be adjusted for an optimum length. A higher heat setting will make the tip longer, and a higher pull setting will make the diameter of the tip narrower.
2. Cut the tapered end so that it is ≥ 1 cm long. Ensure that the internal diameter at the tip of the needle is between 70 µm and 100 µm and that it is inversely proportional to the length of the tapered end.
   NOTE: The internal diameter also depends on the calibration of the micropipette puller. See the instructions from the manufacturer of the vertical micropipette puller or any preferred micropipette puller type.
3. After ensuring that the needle has the correct internal diameter, create the bevel of 15 – 20 degrees by sharpening the tip using a micropipette beveller with a diamond sharpening wheel (Figure 2A – 2C). Make sure to rest the tip gently on the wheel without too much pressure, to decrease chances of breaking or cracking the tip.
   NOTE: A paintbrush can be used to wipe away any debris that builds up on the needle tip.
4. Evaluate the tip under a microscope and ensure that the tip is round without any chips or cracks. Reevaluate the internal diameter to make sure it is between 70 µm and 100 µm (Figure 2D – 2H).
5. Draw lines on the rest of the needle to designate 5 µL of volume between them (e.g., needles with an internal diameter of 1.3 mm should have lines drawn at 3.77 mm increments).
6. Autoclave the needles prior to use in surgery and handle them with sterile gloves. 

2. In utero Injections

1. Prepare the necessary instruments by autoclaving them ahead of time. Include essential instruments such as microinjector needle holder, a surgical needle driver, a pair of Adson forceps, a pair of curved regular tissue scissors, a 1 mL insulin syringe, a couple of cotton-tip applicators, a transfer pipette, a 50 mL conical tube, and a pack of 4-0 polyglactin 910 sutures.
2. Using a sterile technique, attach the needle to the needle holder and plug it into the microinjector.
   NOTE: The settings of the compressed nitrogen used are as followed: inject 4 – 6 psi, balance 0 psi. Depending on what is being injected, specifically, the viscosity of the injectate, as well as the size of the micropipette, the injection times vary between 0.3 – 1.5 s.
3. Clean out the needle tip of any possible debris by drawing up 5 – 10 µL of sterile 1x phosphate-buffered saline (PBS) and then clearing it out. Repeat this 2 – 3x.
4. Prepare pregnant 2- to 6-month-old female mice for surgery by shaving their abdomens with a clipper. Be careful not to damage the nipples. Administer oral pain medication (e.g., 100 µL of 1.5 mg/mL meloxicam oral suspension per mouse).
5. Start filling the needle with the desired material (cells/vector/drug) at the desired volume. Be careful not to break the needle tip while filling the needle.
   NOTE: The volume of injection per fetus varies depending on the specific experimental design. 20 µL works well for injecting a large number of cells (i.e., up to 10⁷ cells into the vitelline vein. For instance, we delivered 1 x 10⁷ whole bone marrow cells isolated from C57BL/6TgN(act-EGFP)OsbY01 ["B6 Green Fluorescent Protein (GFP)"] mice via the vitelline vein into gestational day-14 Balb/c fetuses. For viral vector injections, a single injection of 10 µL of a 1:1 diluted vector with PBS works well.
6. To calibrate the injection time, take the following steps.
   1. Push the Mode button 3x on the microinjector to get to the injection calibration screen. Adjust the injection time by adding intervals of 10 or 100 ms and push the Mode button 2x.
   2. Push the Balance button and push the foot pedal once. Now push the pedal again and assess how much volume is emptied out of the needle per push. If not calibrated to the desired volume of 5 – 20 µL per pedal push, repeat steps 2.6.1 and 2.6.2.
      NOTE: Generally, it is good to calibrate each push to deliver half the total target volume at a time. While it is possible to inject more than 30 µL or even 40 µL of the total volume, we do not generally go over 20 µL per fetus, intravenous or intra-amniotic.
7. Fill the needle up to the desired level.
8. Start delivering anesthesia to the mouse by adjusting the oxygen flowmeter to 1 L/min and the isoflurane vaporizer to 3%.
9. Confirm whether the mouse is anesthetized by checking for the absence of the pedal reflex. Transfer the mouse to a heating pad in a supine position.
10. Apply lubricant eye gel to avoid corneal desiccation. Secure the mouse in place by taping the upper and lower limbs to the pad.
11. Prep the abdomen with chlorhexidine scrub followed by alcohol and inject a local anesthetic (e.g., 100 µL of 0.25% bupivacaine) subcutaneously (Figure 3A).
12. With scissors, make a 1 – 2 cm skin incision so that the lower border is no closer than 1 cm to the introitus; the fascia beneath is very thin and translucent.
13. Identify the midline of the fascia which is more transparent than the surrounding area. Be careful not to injure the epigastric vessels which lie on either side of the midline. Should the epigastric vessels get injured, hold pressure with cotton-tip applicators to stop the bleeding.
14. Using Adson forceps, pinch the fascia without grabbing any of the underlying organs such as the intestines, bladder, or fetuses. Open the fascia with scissors, being careful not to damage any of the organs. Once safely in the abdomen, extend the fascial incision. Make it no longer than the skin incision.
15. Use cotton-tip applicators to move the intestines into the upper part of the abdomen, thus exposing the gravid uterus. Deliver the uterus out of the incision, carefully identifying the right and left ovaries to ensure all fetuses are counted (Figure 3B).
16. Place the left uterus back into the abdomen so that only the right uterus is exposed; this prevents the desiccation of the uterus and keeps the non-injected fetuses warm.
17. Hold the most lateral fetus/amniotic sac between the thumb and index fingers of the operator's non-dominant hand (Figure 3C). Always be gentle as to not cause any damage to the fetuses.
18. Position the dissection microscope (a 10X magnification is ideal) and adjust the focus so that the fetus is in view. Adjust the lighting for a better visualization.
19. Identify the part that will be injected (vitelline vein, amnion).For intravenous injections, visualize both the vitelline veins and their anastomosis first. For intra-amniotic injections, orient the fetus with the right side in view.
20. Reach the target space with the needle as described below.
    1. For an intravenous injection, do as follows.
       1. Rotate the uterus so that the vitelline vein that is being injected is parallel to the tip of the needle; keep in mind that the injections must be made towards the anastomosis of the two veins.
       2. Lay the needle on the uterus at a 5° angle and pierce the uterine wall. Now that the tip is between the uterine wall and the amniotic sac, place the tip directly atop the vitelline vein.
       3. At a nearly tangential angle, glide the needle over the vein until the bevel pierces and advances into the vessel; this is evident by a flash of blood seen in the needle tip (Figure 3D).
          NOTE: Accessing the vein may take a few tries as the needle may not pierce the vein with the first glide over the vein.
    2. For an intra-amniotic injection, do as follows.
       1. Rotate the amniotic sac and find a location devoid of vessels to pierce.
       2. Point the needle perpendicular to the uterine wall and pierce through the uterus, the yolk sac, and then the amniotic sac. Be careful not to pierce through any fetal tissue. Make sure the needle has passed between the limbs as this confirms that the needle is in the amniotic sac. Then proceed with the injection.
21. Inject the appropriate volume of material desired (usually 10 – 20 µL) by pushing the foot pedal.
    NOTE: Because the injector must maintain the visualization of the needle tip through the microscope at all times, a second person must read the markings on the needle to quantify the injected amount and inform the injector how much volume is left to inject. A discussion prior to the injection between the injector and assistant is important to avoid any confusion and delay. This is especially important for intravenous injections because a delayed removal of the needle will allow a backflow of venous blood into the needle and result in an inaccurate dosing.
22. Withdraw the needle from the injection site once the desired volume is delivered. As there may be some bleeding from the vessel puncture site with intravenous injections, hold pressure with the side of the needle for 10 – 15 s to stop the bleeding.
23. Proceed to the next fetus. Continue until all the fetuses of the right uterine horn have been injected.
24. Remove the left uterine horn from the abdomen and replace the right uterine horn back inside the peritoneal cavity.
    NOTE: Occasionally, the needle needs to be refilled with the injectant.
25. Once all fetuses have been injected, replace the uterus into the abdomen (Figure 3E). Make sure to avoid a uterine or intestinal volvulus.
26. With a disposable transfer pipette, place roughly 2 mL of 1x PBS into the abdomen to replace any insensible losses.
27. Close the fascia and abdomen in one continuous layer using 4-0 polyglactin 910 sutures to avoid injuring the underlying organs during the closure (Figure 3F – 3G).
28. Remove the tape and transfer the mouse to a cage underneath a heat lamp. Be careful not to place the heat lamp too close to the mouse. Make sure the cage has bedding, food, and water.
    NOTE: A thermostatically controlled warm chamber may also be used. The mouse is awake when it is upright and walking.
29. Observe the mouse daily and give pain medication as needed.
    NOTE: We routinely give meloxicam on postoperative days 1 and 2, and sometimes on day 3 if the mouse shows signs of pain.
30. If doing a batch surgery with the same injection material, clean out the injection needle with sterile PBS. If injecting with a different material, dispose of the needle in a sharps container and use a new needle.
    NOTE: We recommend fostering the pups with surrogate dams immediately after birth in case the dam mounts an immune response to the injectant and transfers antibodies to the pups via breast milk.