Axonal Regeneration in the Sciatic Nerve of a Mouse Following a Crush Injury

1. Materials and Reagents

1. Animals
   1. Use six-week-old female CF1 mice weighing 30–35 g for the experiments.
      NOTE: The mice were group-housed (5 mice per cage) in individually ventilated sterilized cages with 1/4” corn cob bedding and 2” square nestlets for nesting. The cages were maintained in a controlled 12-h light-dark cycle and the room temperature was between 19.4 °C and 25 °C. The mice were fed with a global rodent diet and automatic water supplying system.
2. Instruments
   1. Use the following surgical instruments to efficiently conduct the surgery: disposable syringes (27 G, 1.0 mL) for intraperitoneal injection, fur clipper, stereo dissection microscope, micro-surgery forceps, micro-surgery scissors, and small bone rongeurs and sterilized non-woven sponges.
      NOTE: All the instruments and materials are autoclaved before surgery or pre-sterilized by the producers. During surgery, the instruments are sprayed with 75% alcohol to maintain sterility.
3. Anesthetic Solutions
   1. Use anesthetic solution #1 to induce the anesthesia and use anesthesia solution #2 before dorsal root ganglia (DRG) injection to maintain the anesthesia.
      1. To make anesthetic solution #1, dilute ketamine and xylazine in sterile saline at a final concentration of 10 mg/mL and 1.2 mg/mL.
      2. To make anesthetic solution #2, dilute avertin in sterile saline at a final concentration of 20 mg/mL.
4. DNA Plasmid and RNA Oligos
   1. Prepare the plasmids using the commercial kit following the manufacturer's protocols accordingly. Dilute the plasmid DNAs in sterile deionized water to the concentration of 2.0 µg/µL.
   2. Add Fast Green dye solution to reach a final concentration of 0.005–0.01% (v/v) for better visualization of the DRG outline during injection.
   3. Dissolve the small interfering RNA (siRNA) or microRNA oligonucleotides (microRNA oligos) in the corresponding buffers provided by manufacturers to reach a final concentration of 50 µM.
5. Micro-Injection Pipette
   1. Pull the capillary glass using the micropipette puller and cut the tip of the pulled capillary glass pipette with microsurgery scissors to generate an opening with an approximate 50 µm outer diameter. Then, sterilize the glass pipette under UV light on a clean bench for approximately 20–30 minutes.
   2. Mount the sterilized glass pipette on the pipette holder connected to an intracellular microinjection dispense system. Set the parameters of the microinjection system at 30 psi of pressure and 8 ms of duration.
6. Electroporation System
   1. Connect the electrodes to the square wave electroporation system and set the following parameters: 15 ms pulses at 35 V with 950 ms intervals for in vivo electroporation.
7. Perfusion and Fixing Solution
   1. Dissolve the paraformaldehyde (PFA) powder in 1x phosphate-buffered saline (PBS) at a concentration of 4% (w/v). Store the PFA solution at 4 °C.

2. Experimental Procedures

1. Surgical Exposure of the L4 and L5 DRGs
   1. Anesthetize the mouse using an intraperitoneal injection of the anesthetic solution #1 prepared previously (with ketamine 100 mg/kg body weight and xylazine 12 mg/kg body weight).
   2. Shave the surgical area with the fur clipper.
      NOTE: Since there will be another surgery for the left sciatic nerve crush, the fur of the left posterior thigh can be shaved simultaneously.
   3. Place the mouse on the heated blanket (35.0 °C) and monitor the rectal temperature with a temperature probe.
   4. To test the anesthesia, pinch the toes and tail of the mouse and observe the behavioral responses.
   5. Tape the four limbs of the mouse on the corkboard.
   6. Wipe the surgical area with an iodophor solution and then use 75% alcohol to remove the iodophor before cutting the skin.
   7. Apply ophthalmic ointment to the eyes to prevent dryness while under anesthesia.
   8. Before incision, mark both sides of the iliac crests with a fine marker pen. Draw a line connecting the two points of the iliac crests to facilitate identifying the positions of L5 DRGs.
   9. Make a 3 cm incision along the midline of the lower back with microscissors. Moreover, detach the paraspinous muscles, such as the musculus multifidus and the musculus longissimus lumborum, from the L3 to S1 spinous processes, and expose the facet joints of L4-5 and L5-6.
   10. Use a micro-rongeur to remove the facet joints of L4-5 and L5-6. Moreover, remove the left neural arch of L4 and L5 to expose the dorsal side of the DRGs.
2. DRG Injection
   1. Inject an anesthetic solution #2 (with avertin 200 mg/kg body weight) intraperitoneally to maintain the anesthesia before DRG injection.
   2. Load the DNA plasmids or RNA oligos (1 µL per DRG) into the glass capillary pipette.
   3. Insert the tip of the capillary glass pipette carefully into the DRG.
   4. Gradually inject 1.0 µL solution of DNA plasmids or RNA oligos into the DRG using the intracellular microinjection dispense systems (30 psi, 8 ms).
      NOTE: The duration of the injection should last no less than 5 minutes.
3. Electroporation
   1. Drop PBS on the tips of the electrodes.
   2. Clean up the bleeding with sterilized square cotton gauze.
   3. Pinch the target DRG with the electrodes gently and apply 5 square electric pulses with the electroporation system.
   4. Close the muscle and skin layers respectively with 5-0 nylon sutures.
   5. Place the mouse on a heated blanket (35.0 °C) under close attention until it has regained sufficient consciousness to maintain sternal recumbency. Return the mouse to the home cage after it has fully recovered from the anesthesia.
      NOTE: It takes approximately 60 minutes for the mouse to completely recover from anesthesia on the 37 °C blanket. Dissolve one pill (200 mg) ibuprofen into 1,000 ml water (0.2 mg/ml) for daily feeding to relieve the post-surgical pain.
4. Sciatic Nerve Crush
   1. Two or three days (depending on the experimental design) after the DRG electroporation, anesthetize the mouse intraperitoneally with anesthetic solution #2 (with avertin 400 mg/kg body weight).
   2. Tape the four limbs of the mouse on the corkboard.
   3. Make a 1 cm incision 0.5 cm to the left side along the midline. Cut the muscles, such as the gluteus maximus muscle and the piriformis muscle, longitudinally. Expose the segment of the sciatic nerve between the greater sciatic foramen and the sciatic notch.
   4. Crush the nerve with microsurgery forceps for 12 s and mark the crush site with a 10-0 nylon epineural suture. Make a knot on the dural membrane to mark the crush site.
   5. Close the muscle and skin layers with 5-0 nylon suture.
   6. Place the mouse on a heated blanket (35.0 °C) with close attention until it has completely regained sufficient consciousness to maintain sternal recumbency. Return the mouse to the home cage after it has fully recovered from the anesthesia.
      NOTE: It takes approximately 60 min for the mouse to completely recover from anesthesia on the 37 °C blanket. Dissolve one pill (200 mg) ibuprofen into 1,000 ml water (0.2 mg/ml) for daily feeding to relieve the post-surgical pain.
5. Mouse Perfusion, DRG, and Sciatic Nerve Harvest
   1. Two or three days after the sciatic nerve crush (depending on the experimental design), anesthetize the mouse intraperitoneally with anesthetic solution #2.
   2. Perfuse the mouse transcardially with PBS (pH 7.4) followed by ice-cold 4% paraformaldehyde (PFA) (pH 7.5) in PBS.
   3. After perfusion, separate the DRGs together with the nerve roots and sciatic nerve before the knee carefully with micro-scissors and micro-forceps under the dissection microscope. Place the sciatic nerve directly in 4% PFA overnight at 4 °C.
6. Imaging and Measuring the Fluorescence-Labeled Sensory Axons
   1. Strip off the attached tissue and membrane on the fixed sciatic nerve with micro-scissors and micro-forceps carefully under the dissection microscope. Then, exchange the 4% PFA with PBS and wash the nerve three times.
   2. Place the sciatic nerve on a slide and keep it straight. Add 80 µL of antifade solution around the nerve, then lay a coverslip on it. Flatten the whole mounted tissue with pressure.
   3. Place the flattened tissues on the inverted epifluorescent microscope equipped with an accessory for mosaic acquisition and image processing.
      NOTE: The excitation and emission lengths are 488 nm and 509 nm. It is also possible to take multiple overlapping images manually and stitch the images together using ImageJ with the Mosaic plug-in.
   4. When measuring the length of regenerated axons, trace all identifiable fluorescently-labeled axons in the sciatic nerve from the crush site (marked with the 10-0 epineural suture) to the distal axon ends.