Combined In Vivo Electroporation and Short-Term Reinnervation of the Cranial Levator Auris Longus Skeletal Muscle

Summary

Here, we present a protocol that combines in vivo electroporation and denervation of the cranial levator auris longus (LAL) muscle. This procedure enables the study of the potential role of muscle-derived proteins in the regeneration of the neuromuscular synapse.

Abstract

The neuromuscular junction (NMJ) is the peripheral synapse controlling the contraction of skeletal muscle fibers to allow the coordinated movement of many organisms. At the NMJ, a presynaptic motor axon terminal contacts a muscle postsynaptic domain and is covered by terminal Schwann cells. The integrity and function of the NMJ is compromised under several conditions, including aging, neuromuscular diseases, and traumatic injuries. To analyze the potential contribution of muscle-derived proteins to NMJ maintenance and regeneration, an in vivo gene transfer strategy has been combined with the denervation of the cranial levator auris longus (LAL) muscle after mechanical nerve injury. Previous findings showed that the forced expression of control fluorescent proteins does not alter NMJ organization or neurotransmission. This procedure aims to describe a detailed method of in vivo electroporation of the LAL muscle followed by transection or crushing of the specific branch of the facial nerve innervating cranial muscles, leading to NMJ denervation and reinnervation, respectively. The combination of these experimental strategies in the convenient LAL muscle constitutes an efficient method to study the potential contribution of muscle protein overexpression or silencing in the context of short-term NMJ reinnervation.

Introduction

The neuromuscular junction (NMJ) is the peripheral synapse that controls muscle contraction and, indirectly, the coordinated movement of organisms¹. It is formed by a presynaptic motor axon terminal, a muscle postsynaptic domain enriched in acetylcholine receptors (AChRs), and non-myelinic terminal Schwann cells covering the axon terminal^2,3,4. Upon NMJ denervation due to traumatic peripheral nerve injury, disease, or pharmacologic intervention, contractile muscle activity is lost^5,6. As certain permissive microenvironments allow timely and efficient functional recovery, the search for proteins that could help the regeneration process is a permanent necessity. Here, two procedures have been combined to specifically evaluate the potential role of manipulating muscle protein expression in the short-term regeneration of the NMJ after nerve injury.

The levator auris longus (LAL) muscle, located on the dorsal surface of the skull, is a thin and flat muscle that controls the movement of the pinna. The LAL muscle consists of rostral and caudal bands, each containing two or three layers of muscle fibers^7,8. The posterior auricular branch of the facial nerve innervates the LAL muscle, generating a well-described innervation pattern consisting of five rostral (R1-R5) and two caudal (C1-C2) innervation regions^8,9,10. The LAL is a superficially exposed and easily accessible muscle, so its use requires minimally invasive procedures. This allows visualization of the NMJ using real-time microscopy, drug delivery, as well as the intervention of complete muscle preparations both in vivo and ex vivo^11,12. Altogether, these features make the LAL muscle an excellent model to study NMJ behavior and function^10,13.

In vivo electroporation is a gene-transfer technique that allows the incorporation of exogenous DNA into the tissue through the transient permeabilization of cell membranes and the mobility of DNA inside the cells, both generated after inducing an electric field^14,15. This procedure is local and can be performed at any stage of the animal's development. Considering the unique features of the LAL muscle, its in vivo electroporation represents a minimally invasive, highly efficient, and quick procedure¹⁶, making it possible to observe robust protein expression two or three days after electroporation.

The facial nerve crush procedure has been widely employed in research as it offers several advantages^5,17,18. This protocol has been previously modified to specifically target the posterior auricular branch of the facial nerve that innervates the cranial muscles, including the LAL muscle, to avoid facial paralysis; consequently, mice do not require special care after surgery (e.g., lubricating eye ointment for the loss of blink reflex). As NMJ reinnervation at the LAL muscle begins between 7-9 days after injury⁶, it is a good short-term reinnervation model compared to the commonly used nerve crush injury model of the sciatic nerve to denervate hindlimb muscles, where reinnervation occurs around three weeks post nerve injury¹⁸.

The combination of in vivo electroporation and facial nerve injury at the LAL muscle will enable researchers to evaluate the behavior of synaptic components against muscle-derived protein modulation at different times of NMJ regeneration, including short-term and long-term morphological effects^6,16, through a wide variety of techniques including immunohistochemical, histological, and functional assays.

Protocol

All experimental procedures are approved by the Bioethics Committee at Universidad Austral de Chile, Chile (protocol Nr. 503/2023), and followed the norms imposed by the Bioethics Committee of the National Research and Development Agency, Chile (ANID), as well as the guidelines of the European Council Directive for the Care of Laboratory Animals. The present study used adult (3-6 months old) CF-1 mice of both sexes (30-45 g). Euthanasia was accomplished by inhalant anesthetic overdose followed by exsanguination. The reagents and equipment used in the study are listed in the Table of Materials.

1. In vivo LAL muscle electroporation

1. Preparation
   1. Gently sedate the mouse in an acrylic induction chamber under inhalation of 2.5% isofluorane with 0.8 L/min oxygen (following institutionally approved protocols). After that, maintain mouse sedation using a nose tubing mask connected to an inhalation anesthesia machine under a dissecting stereomicroscope.
      ​NOTE: Ensure that the mouse is asleep before starting the procedure; test the touch reflex of the mouse by gently squeezing one foot with a forceps (do this step every time after inducing anesthesia).
   2. Disinfect the cervical area at the level of the sagittal suture of the skull with the antiseptic chlorhexidine gluconate 2% and inject subcutaneously 20 µL of 2 mg/mL hyaluronidase in PBS using a Hamilton syringe to facilitate the DNA plasmid access to the muscle surface.
   3. Turn off the isofluorane flow and allow the mouse to breathe pure oxygen for 1 min.
   4. Weigh the mouse to calculate the appropriate Meloxicam dose (see step 1.3.2.).
   5. Place the animal in an empty cage (without any bedding) to recover from anesthesia.
   6. Return the animal to normal housing for 30 min¹⁶.
2. Surgery procedure
   1. Re-anesthetize the mouse by the inhalation of 2.5% isofluorane with a 0.8 L/min oxygen mixture (following institutionally approved protocols) and keep it sedated throughout the surgical procedure using an inhalation anesthesia machine, under a dissecting stereomicroscope.
   2. Shave the cervical area using a double-edged razor blade and disinfect with chlorhexidine and 70% ethanol 3 times.
   3. Make an 8 mm incision over the sagittal suture of the skull and expose the LAL muscle by gently removing the fat and connective tissue located just under the skin using scissors/forceps.
      NOTE: The LAL muscle is the most superficial muscle in the cranial region after skin incision. It is easily recognized by the orientation of muscle fibers that run diagonally from the middle skull towards each pinna^8,16.
   4. Using a Hamilton syringe, inject 10 µL of the commercially obtained DNA (0.8-1 µg/µL) in PBS into the fascia that is distributed throughout the muscle, forming a bubble.
   5. Place two gold needle electrodes 5 mm apart, parallel, and over muscle fibers, occupying the entire length of the LAL muscle (Figure 1A).
   6. Using an electroporator, generate five pulses of 100 V/cm of 20 ms duration each, at 1 Hz frequency.
   7. Repeat the procedure in the contralateral LAL muscle (repeating steps 1.2.4-1.2.6).
3. Suture and recovery
   1. Suture (5/0 strain size, HR15 needle) the skin with absorbable polyglycolic acid thread.
   2. Inject subcutaneously the analgesic Meloxicam in doses of 1-2 mg/kg of weight to reduce post-surgery pain.
   3. Turn off the isofluorane flow and allow the mouse to breathe pure oxygen for 1 min.
   4. Place the animal in an empty cage (without any bedding) to recover from anesthesia.
   5. Return the animal to its regular housing and monitor it for 2 days after surgery or until its recovery. Monitor pain or stress signs, allocating scores for each of the different parameters considering the humane endpoint criteria previously described¹⁹.
      NOTE: Position the mouse in a heated pad covered with a surgical pad to prevent its body temperature from dropping during surgery. Try not to take more than 30 min for the surgical procedure, thus preventing the mouse from dying due to excess anesthesia. If the expressed protein yields low fluorescence levels, it is recommended to co-electroporate with a tracer (e.g., tdTomato or GFP) in a 1:5 ratio (tracer: DNA of interest).

2. Crush nerve injury-induced denervation

NOTE: This protocol is a modification of a previously described protocol¹⁷. Perform this procedure with a difference of at least 3 days from the electroporation. Depending on the research questions, this procedure can be performed before or after electroporation.

1. Anesthesia and animal positioning
   1. Perform the animal sedation in an acrylic induction chamber under inhalation of 2.5% isofluorane with 0.8 L/min oxygen (following institutionally approved protocols).
   2. Weight the mouse to calculate the Meloxicam dose (see step 2.3.2.).
   3. Maintain mouse sedation with a nose tubing mask connected to an inhalation anesthesia machine under a dissecting stereomicroscope.
   4. Position the mouse on its left side (to denervate the right LAL muscle). Tape the right ear towards the nose to expose the posterior area of the pinna.
2. Surgery procedure
   1. Shave the posterior area of the pinna using a double-edged razor blade and disinfect with chlorhexidine and 70% ethanol 3 times.
   2. Locate the posterior auricular vein and make an incision in the skin, 2 mm away caudally from it, of approximately 5 mm with spring scissors. Avoid cutting blood vessels and/or muscle tissue.
   3. Cut the fat and connective tissues with forceps and scissors. Find the spinal accessory nerve; continue until finding the belly of the digastric muscle and the cartilaginous auditory canal (semi-transparent whitish tissue) (Figure 2A).
   4. Locate the beginning of the facial nerve branching just below the digastric muscle.
   5. Follow the posterior auricular branch (the most dorsal branch of the facial nerve), and gently remove the connective tissue surrounding it, carefully avoiding direct nerve manipulation.
   6. Crush the posterior auricular branch of the facial nerve for 30 s, applying constant pressure on the nerve with a forceps with a 45° angled tip.
3. Suture and recovery
   1. Suture (5/0 strain size, HR15 needle) the skin with absorbable polyglycolic acid thread.
   2. Inject subcutaneously the analgesic Meloxicam in doses of 1-2 mg/kg of weight to reduce post-surgery pain.
   3. Remove the tape from the ear, turn off the isofluorane flow, and allow the mouse to breathe pure oxygen for 1 min.
   4. Place the animal in an empty cage (without any bedding) to recover from anesthesia.
   5. Return the mouse to its regular housing and monitor it for 2 days after surgery or until its recovery.
   6. After recovery, examine the mouse for failures in the movement of the affected ear as a sign of efficient denervation.
      NOTE: In Sham control animals, make an incision in the skin, expose the facial nerve without touching it directly, and subsequently suture.

Results

The high fluorescence quantum yield of the tdTomato protein²⁰ makes it an appropriate tracer for electroporation efficiency. Therefore, the expression of tdTomato protein in LAL muscles by in vivo electroporation was evaluated (Figure 1). LAL muscles were dissected 21 days after electroporation. Whole-mount preparations show the thin caudal band (cLAL, cyan polygon) of the LAL muscle with its two innervation regions, and the thick rostral band (rLAL, yellow p...

Discussion

The combination of in vivo muscle electroporation and denervation is a valuable experimental approach to investigating a regenerative niche at the NMJ⁴. Since the LAL muscle is superficially exposed, these combined procedures can be readily complemented with the delivery of probes or drugs that could affect protein expression or activity¹². In addition, electroporation of reporter genes into the LAL muscle could provide a valuable tool to follow the activation or i...

Materials

Name	Company	Catalog Number	Comments
#5/45 Dumont forceps	Fine Science Tools	11251-35	Sterilize before use
anti 2H3	DSHB		Dilute 1 : 300
anti S100 Ready to Use	Dako	IR504	Dilute 1 : 5
anti SV2	DSHB		Dilute 1 : 50
Chlorhexidine gluconate 2%	DifenPharma
Cold light dissecting stereomicroscope	Motic	Model SMZ-171
DAPI	Invitrogen	D1306	Dilute 1 : 100
Dumont #5SF Forceps	Fine Science Tools	11252-00	Sterilize before use
Dumont Mini Forceps –Style 5	Fine Science Tools	11200-14	Sterilize before use
ECM 830 electroporator	BTX Harvard Apparatus
Gold needle-type electrodes	Genetrodes, BTX	45-0114	10 mm straight
Hamilton syringe	Hamilton	80400
Hyaluronidase	Sigma-Aldrich	H3884
Inhalation anesthesia machine	SciVerma Scientific	M3000 Table Top
Isoflurane	Baxter	218-082
Mice	Instituto de Salud Publica - Chile		Adult (3-6 months old) CF-1 mice, of both sexes (30-45 g)
Razor blades	Schick
Suture Glicosorb 5/0	TAGUM	GS0812.J	Absorbable polyglycolic acid thread
tdTomato plasmid	Addgene	54642	tdTomato plasmid under the control of the CMV promoter
Tramadol hydrochloride 5% (Triamcol)	Drag Pharma
Vannas Spring Scissors	Fine Science Tools	91500-09	Sterilize before use
WGA Alexa Fluor 488	Invitrogen	W7024	Dilute
α-BTX Alexa Fluor 488	Invitrogen	B13422	Dilute 1 : 500