Investigating Mammalian Axon Regeneration: In Vivo Electroporation of Adult Mouse Dorsal Root Ganglion

Summary

Electroporation is an effective approach to deliver genes of interests into cells. By applying this approach in vivo on the neurons of adult mouse dorsal root ganglion (DRG), we describe a model to study axon regeneration in vivo.

Abstract

Electroporation is an essential non-viral gene transfection approach to introduce DNA plasmids or small RNA molecules into cells. A sensory neuron in the dorsal root ganglion (DRGs) extends a single axon with two branches. One branch goes to the peripheral nerve (peripheral branch), and the other branch enters the spinal cord through the dorsal root (central branch). After the neural injury, the peripheral branch regenerates robustly whereas the central branch does not regenerate. Due to the high regenerative capacity, sensory axon regeneration has been widely used as a model system to study mammalian axon regeneration in both the peripheral nervous system (PNS) and the central nervous system (CNS). Here, we describe a previously established approach protocol to manipulate gene expression in mature sensory neurons in vivo via electroporation. Based on transfection with plasmids or small RNA oligos (siRNAs or microRNAs), the approach allows for both loss- and gain-of-function experiments to study the roles of genes-of-interests or microRNAs in regulation of axon regeneration in vivo. In addition, the manipulation of gene expression in vivo can be controlled both spatially and temporally within a relatively short time course. This model system provides a unique tool to investigate the molecular mechanisms by which mammalian axon regeneration is regulated in vivo.

Introduction

Injuries in the nervous system caused by neural trauma or various neurodegenerative diseases usually result in defects in motor, sensory and cognitive functions. Recently, much effort has been devoted to regenerative potency re-establishment in adult neurons to restore the physiological functionsof injured neurons^1,2,3. Sensory neurons in the DRG are a cluster of nerve cells that convey different sensory stimuli, such as pain, temperature, touch, or body posture, to the brain. Each of these neurons is pseudo-unipolar and contains a single axon that bifurcates with one branch extending toward the periphery and the other branch heading toward the spinal cord⁴. The adult sensory neurons in DRGs are among a few mature mammalian neurons known to regenerate their axons actively after injury. Hence, injuries of sensory axons have been extensively employed as a crucial model to study the mechanisms of axonal regeneration in vivo.

In vivo gene transfection techniques, which are usually less time-consuming to set up and more flexible than using transgenic animals, have been playing essential roles in studying the functions of genes and signaling pathways in the nervous system. The main techniques can be categorized into two approaches: instrument-based and virus-based⁵. Viral-based in vivo gene delivery in adult neurons can provide precise spatiotemporal manipulation of gene expression⁶. However, labor-intensive processes are involved in viral-based methods, such as the production and purification of viral particles containing the desired gene. In addition, many viral vectors could activate the immune system of the host, which may interfere with the data acquisition, data analysis and possibly mislead the interpretation of experimental results. Electroporation, a typical instrument-based transfection approach, uses an electrical pulse to increase the permeability of cell and nuclear membranes transiently, which favors the influx of gene vectors or small RNA oligos from space outside of the cells⁷. In vitro electroporation is widely recognized as a transient but highly efficient strategy for manipulating targeted gene expression in many cell types. Although in vivo electroporation only leads to transient gene expression with low transfecting efficiency compared to viral vectors, it has various advantages over viral approaches. For instance, it can be applied to almost all tissues and cells^7,8,9. In addition, either plasmids encoding genes-of-interest or small RNA oligos (e.g., siRNAs, microRNAs) against certain transcripts can be injected into the target tissue directly and then electrically pulsed, which make the procedure less labor- and time- consuming. Moreover, transfecting multiple plasmids and RNA oligos simultaneously with single electroporation is possible.

We have established an in vivo electroporation approach to manipulate gene expression in adult mouse sensory neurons and successfully applied and validated such approach in numerous pioneer studies^1,2,3,8,10. Here, we present a detailed protocol to facilitate the usage of this approach for future studies of mammalian axon regeneration.

Protocol

All animal experiments were performed in accordance with the animal protocol approved by the Johns Hopkins Institutional Animal Care and Use Committee.

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 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 the sterilization.
3. Anesthetic Solutions
   1. Use anesthetic solution #1 to induce the anesthesia and use anesthesia solution #2 before 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 siRNA or microRNA oligos in corresponding buffers provided by manufacturers to reach a final concentration at 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 20–30 min approximately.
   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 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 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 iodophor solution and then with 75% alcohol to remove the iodophor before cutting the skin.
   7. Apply ophthalmic ointment on 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 two points of iliac crests to facilitate identifying the positions of L5 DRGs.
   9. Make a 3 cm incision along the midline of low back with micro-scissors. 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 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 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 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.
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 gluteus maximus muscle and 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.

Results

To quantify the cytotoxicity of the current protocol and to validate that transfection rate of in vivo DRG electroporation is high enough, we injected and electroporated fluorescently-tagged microRNA or siRNA into L4 and L5 DRGs. The detached DRGs were processed through cryo-sectioning and immunohistochemistry (Figure 1A-B). When estimating the cell survival rate after injection and electroporation, the intact DRGs from L4 and L5 wer...

Discussion

Several surgical steps require particular attention. The L4 and L5 DRGs (location of somas), which dominate the sciatic nerve, need to be correctly identified and injected with gene constructs. Otherwise, the GFP-labeling will be absent in sciatic nerve axons. The iliac crests can be viewed as useful anatomical landmarks to pinpoint L4 and L5 DRGs. In most mice, the facet joint between L5 and L6 vertebrae is proximate to iliac crests¹². Alternatively, L3 DRG can be chosen instead of L5, especially...

Materials

Name	Company	Catalog Number	Comments
ECM 830 Square wave electroporation system	BTX Harvard Apparatus	45-0052	For in vivo electroporation
Tweezertrodes electrodes	BTX Harvard Apparatus	45-0524	For in vivo electroporation, 1 mm flat
Picospritzer III	Parker Instrumentation	1096	Intracellular Microinjection Dispense Systems
Glass Capillary Puller	NARISHIGE	PC-10
Borosilicate Glass Capillaries	World Precision Instruments, Inc.	1902328
Stereo Dissection Microscope	Leica	M80
Microsurgery Rongeur	F.S.T	16221-14
Microsurgery Forceps	FST by DUMONT, Switzerland	11255-20	Only for sciatic nerve crush
Glass Capillary	World Precision Instruments, Inc.	TW100-4	10 cm, standard wall
Tape	Fisherbrand	15-901-30	For fixing the mouse on the corkboard
2, 2, 2-Tribromoethanol (Avertin)	Sigma-Aldrich	T48402	Avertin stock solution
2-methyl-2-butanol	Sigma-Aldrich	152463	Avertin stock solution
siRNA Fluorescent Universal Negative Control #1	Sigma-Aldrich	SIC003	Non-target siRNA with fluorescence
microRNA Mimic Transfection Control with Dy547	Dharmacon	CP-004500-01-05	Non-target microRNA with fluorescence
Plasmids preparation kit	Invitrogen Purelink	K210016	GFP-coding plasmid preparation
Fast Green Dye	Millipore-Sigma	F7252	For better visualization of the DRG outline during injection
Ketamine	Putney, Inc	NDC 26637-731-51	Anesthesia induction
Xylazine	AnaSed	NDC 59399-110-20	Anesthesia induction
Acetaminophen	McNeil Consumer Healthcare	NDC 50580-449-36	Post-surgical pain relief