Hydrodynamic Renal Pelvis Injection: A Technique to Directly Deliver DNA into Kidney Cells of Murine Model

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Perform the Hydrodynamic Renal Pelvis Injection Surgery

1. Select mice carefully for surgery.
   NOTE: Strain-specific differences have not yet been observed but may be possible. Most injections have been on the C57BL/6 or FVB backgrounds. Renal pelvis injections work best in mice that are 6–12 weeks old. In mice greater than 16 weeks, up to a 50% failure rate by luciferase imaging is possible for unclear reasons. The same age-related failure rate has been observed for hydrodynamic tail vein injections to the liver so this may be a general limitation relating to the principle of hydrodynamic injection. Along the same lines, others have shown that hydrodynamic tail vein injection into fibrotic rat liver is not as effective as healthy liver, so it may be possible that in settings of renal fibrosis the renal pelvis hydrodynamic injection will not be as effective, but this has not been tested directly.

2. Prepare Mice and DNA Syringes for Surgery

1. Anesthetize the mice with ketamine and xylazine.
   1. Put on the correct personal protective equipment required by the animal facility, such as disposable lab apron, surgical face mask, and nitrile gloves.
   2. Working with 2–4 mice at one time, weigh each mouse in a 500 mL plastic beaker on a scale that is accurate to 0.1 g. Calculate the correct amount of 24 mg/mL ketamine and 2 mg/mL xylazine diluted in normal 0.9% saline to administer to each mouse by intraperitoneal injection (see referenced video for more on intraperitoneal injection). Use the formula (50 μL + ((5 μL) x (weight(g))), or alternatively calculate according to another formula after consultation with the local veterinarian team and IACUC.
   3. Inject the mouse by intraperitoneal injection by standard techniques. Place the mouse in a paper bucket until the mouse is immobile.
      NOTE: Mice are ready for surgery when they no longer respond to the toe-pinch test. Give mice that continue to respond to the toe-pinch test 20–30 min after the initial injection 20–60 μL more anesthetic, depending on the strength of the response.
   4. Place the mouse on a water-circulated heat pad covered with a blue pad and place vet ointment in both eyes to prevent corneal desiccation.
2. Shave the left side of the back of the mouse from shoulder to rump and flank to spine with a shaver designed for pet grooming. Remove loose hair and debris.
3. Prepare a separate syringe containing 100 μL of diluted DNA for each anesthetized mouse, making sure there are no air bubbles.
   1. Use a sterile 29G x ½" 0.5 mL U-100 insulin syringe with a permanently attached needle without a safety.
      NOTE: The syringe type is of critical importance. This gauge allows for a fast injection. The permanently attached needle prevents the solution from leaking out. The presence of safety will impede access to the renal pelvis. The syringes specified in the Materials Table glide more evenly during injection than other brands tested.
   2. Load the syringe to ~120 μL and pull the plunger down to create a space at the top. Tap with a pen until all air bubbles rise to the top. Holding the needle up, carefully depress the plunger to remove all air until a droplet forms at the tip of the needle. There should not be any visible bubbles present, as these may cause an air embolism that will kill the animal.
   3. Finish depressing the plunger until there is 100 μL in the syringe by emptying excess DNA solution into the original microfuge tube. Carefully label if necessary and place the loaded syringes on a sterile drape to maintain sterility if the facility where work is being performed does not allow recapping of needles.

3. Perform Injection Surgery

1. Prepare the site for surgery. Place a sterile drape over the heat pad and empty sterile surgical tools onto the sterile drape without touching them. Pick up the mouse and place it in the field of view. Adjust lighting to illuminate the area.
2. Remove three 3.15% chlorhexidine gluconate and 70% isopropyl alcohol skin antiseptic swabs from the packages and place near the animal.
3. Change into sterile surgical gloves. Working in a circular motion beginning at the incision site, swab the animal with a new chlorhexidine/alcohol swab three times.
4. Locate the incision site as shown in Figure 1A. Pinching the skin with tweezers, use scissors to make a cut to the skin layer approximately 1 cm from the spine and below the ribcage. Once the cut site is approximately 1 cm long, make a similar cut site below, in the muscle layer.
   NOTE: Make the incision the right length to allow the kidney to just barely come through the incision and then be kept in place by the incision itself. Too small an incision and the kidney cannot be exposed; too large and the kidney will continually slip back into the abdominal cavity.
5. Locate the kidney.
   NOTE: It may be visible amongst white adipose tissue. The spleen is also located on the left side of the animal. The color of these organs can be visually differentiated, as the spleen is a dark maroon while the kidney is a dark red-orange. It is not desirable to manipulate the spleen as it can be easily ruptured.
6. Without touching the kidney, gently expose it from the abdominal cavity by putting steady, gentle pressure on the abdomen with the fingers (Figure 2C). Use closed tweezers to gently push undesired organs back into the abdomen if necessary. Do not use open forceps as this may damage the kidney or other organs.
7. Once the kidney is out of the abdomen, gently separate it from the surrounding fat just enough to visualize the renal pelvis, a small white dot (Figure 1B). Push excess fat to one side or remove if necessary. Be sure not to remove the adrenal gland, located near the top pole of the kidney, or the kidney capsule.
8. Using closed forceps to push down on the right side of the kidney so that the renal pelvis remains in view, grasp the loaded insulin syringe with the right hand and hold it parallel to the working surface with the needle pointed at the renal pelvis (Figure 2E). Insert the needle carefully into the renal pelvis of the immobilized kidney as shown in Figure 1B.
9. Inject the solution quickly within three seconds. About one-third of the kidney may clear and change color to white.
   NOTE: It is common to observe fluid build-up in the kidney capsule following injection as well as the formation of a hematoma. Some damage is necessary to achieve a sufficient level of DNA transfection for detection.
10. Keep the needle in place for approximately 10 s to prevent backflow of the solution. Then carefully and slowly remove the needle. Return the organ to the abdominal cavity by gently stretching the incision site and using closed forceps.
11. Suture the muscle layer of the animal with 5-0 absorbable sutures, placing 2–4 independent knots.
12. Suture the skin layer of the animal with 5-0 or 6-0 non-absorbable nylon sutures, placing 2–4 independent knots.
    NOTE: Surgical tools may be reused after placing in a sterile bead bath and cooled down.