The Fibular Nerve Injury Method: A Reliable Assay to Identify and Test Factors That Repair Neuromuscular Junctions

Podsumowanie

We have developed a nerve injury method to reliably examine muscle reinnervation, and thus regeneration of neuromuscular junctions in mice. This technique involves injuring the common fibular nerve via a simple and highly reproducible surgery. Muscle reinnervation in then assessed by whole-mounting the extensor digitorum longus muscle.

Streszczenie

The neuromuscular junction (NMJ) undergoes deleterious structural and functional changes as a result of aging, injury and disease. Thus, it is imperative to understand the cellular and molecular changes involved in maintaining and repairing NMJs. For this purpose, we have developed a method to reliably and consistently examine regenerating NMJs in mice. This nerve injury method involves crushing the common fibular nerve as it passes over the lateral head of the gastrocnemius muscle tendon near the knee. Using 70 day old female mice, we demonstrate that motor axons begin to reinnervate previous postsynaptic targets within 7 days post-crush. They completely reoccupy their previous synaptic areas by 12 days. To determine the reliability of this injury method, we compared reinnervation rates between individual 70 day old female mice. We found that the number of reinnervated postsynaptic sites was similar between mice at 7, 9, and 12 days post-crush. To determine if this injury assay can also be used to compare molecular changes in muscles, we examined levels of the gamma-subunit of the muscle nicotinic receptor (gamma-AChR) and the muscle-specific kinase (MuSK). The gamma-AChR subunit and MuSK to are highly upregulated following denervation and return to normal levels following reinnervation of NMJs. We found a close relationship between transcript levels for these genes and innervation status of muscles. We believe that this method will accelerate our understanding of the cellular and molecular changes involved in repairing the NMJ and other synapses.

Wprowadzenie

In young adult and healthy animals, the neuromuscular junction (NMJ) is a highly stable connection between the presynapse, the nerve ending of an α-motor axon, and the postsynapse, the specialized region of an extrafusal muscle fiber where nicotinic acetylcholine receptors (AChRs) selectively aggregate¹. The nearly perfect apposition of the pre- and post-synaptic apparatuses is necessary for proper neurotransmission, survival of α-motor neurons and muscle fibers and motor function. Unfortunately, the function of the NMJ is adversely affected by aging, diseases such as amyotrophic lateral sclerosis (ALS), autoimmune diseases and injury to muscles and peripheral nerves^2-5. These insults often result in degeneration of presynaptic nerve endings, leaving muscles denervated and significantly altering motor skills. For this reason, the identification of molecules that function to maintain and repair the NMJ has become a priority. Because peripheral nerves regenerate and reinnervate targets, peripheral nerve injury models have been used to identify molecular changes associated with regenerating NMJs.

Peripheral nerve injury models often involve either completely cutting or crushing specific nerve branches⁶. Following a cut, the endoneurial tube has to be reformed, delaying axonal regeneration and reinnervation of target cells and tissues. The severity of this type of injury also causes axons to meander away from their original path, resulting in their failure to reach original targets. This is in contrast to nerves injured via crush where the endoneurium remains contiguous, providing a path for efficient and proper regrowth of regenerating axons. It also allows axons to find and reinnervate their original muscle fiber partners. Irrespective of injury model, there are a number of cellular and molecular changes that must occur for axons to regenerate and reinnervate targets. After an injury, the nerve segment proximal to the target is broken down and removed via a process termed Wallerian Degeneration⁷. This process involves reprogramming and de-differentiation of Schwann cells into non-myelinating cells that secrete regenerative factors, clear myelin, and recruit macrophages to the site of injury⁸. Macrophages in turn complete the clearance of myelin and axonal debris, which would otherwise impede growth of the regenerating axon⁹. In parallel, motor and sensory neurons activate mechanisms needed to promote regeneration of their severed axons. Once the regenerating axon reaches the target, it must transform from a growth cone to a nerve ending capable of properly transmitting (for motor axons) or receiving (for sensory axons) information¹⁰. In this regard, alpha-motor axons undergo a series of well-orchestrated changes that culminate in their growth cone differentiating into a fully functional presynaptic nerve ending that nearly perfectly opposes the post-synaptic site on the target muscle fiber¹¹.

The sciatic, tibial and accessory nerves have been the primary choices for studying axonal and NMJ regeneration^12-14. However, there are a number of drawbacks when using these models to examine cellular and molecular changes associated with regenerating NMJs between animals and under different conditions. Firstly, the sciatic nerve supplies the majority of the muscles of the hind limb, with injury significantly limiting both movement and sensation. It is therefore not possible to use this method to study the impact of exercise alone or in combination with other factors. Additionally, the sciatic nerve is a rather thick structure and thus requires a large amount of compressive force to fully injure all axons. This in turn may result in complete transection of the more superficial axons while leaving the endoneurial tube of deeper lying axons intact, introducing significant variability in the rate and fidelity of regeneration among these axons. Complete transection of this nerve is even less desirable given that many axons will fail to reconnect with the same muscle fibers. Complicating matters, the sciatic nerve possesses intrinsic anatomic variability, both in the number and site of origin of its terminal nerve branches. It is therefore very difficult to lesion the same site. While the tibial nerve is smaller and more amenable to crush injuries, there is also no readily available landmark to serve as a lesion site for this nerve branch.

The accessory nerve branch (part of cranial nerve XI) that supplies the sternocleidomastoid muscle has also been used to study regeneration of NMJs¹⁵. This nerve is particularly attractive because NMJs in the sternocleidomastoid muscle can be more readily imaged in live animals compared to NMJs in other muscles. But similar to the sciatic and tibial nerves, there is no specific landmark that can be used to injure this nerve in the same location, limiting it as a model for comparing regeneration of NMJs among individual animals of an experimental cohort. An inconsistent lesion site introduces variability in the rates of NMJ reinnervation. Due to these shortcomings, the procedure presented here utilizes the injury of a different peripheral nerve branch to examine regenerating NMJs.

The common fibular nerve, also called the common peroneal nerve, contains many features that make it a reliable nerve to examine regeneration of NMJs between animals and across different treatments. The common fibular nerve has a predictable anatomic course as it runs over the tendon of the lateral head of the gastrocnemius muscle in the knee, the intersection serving as a stable landmark for lesions. The nerve is accessed through a small and minimally invasive incision near but anatomically segregated from the muscles of interest. The findings presented here demonstrate that regenerating motor axons begin to reform NMJs in the extensor digitorum longus (EDL) muscle 8 days after crushing the fibular nerve in 70 days old young adult female mice. Importantly, the pattern and rate of reinnervation is consistent among animals of the same age and sex and therefore provide a reliable injury model that will significantly hasten our understanding of the cellular and molecular changes required to maintain and repair NMJs.

Protokół

All experiments were carried out under NIH guidelines and animal protocols approved by the Virginia Tech Institutional Animal Care and Use Committee.

1. Preparing Animals for Surgery

1. Anesthetize mice with a mixture of ketamine (90 mg/kg) and xylazine (10 mg/kg) via subcutaneous inguinal injection with a sterile 1 ml insulin syringe. Carrier solution contains a mixture of 0.9% saline, 17.4 mg/ml ketamine, and 2.6 mg/ml xylazine. Place animals back in cages while waiting for medication to take effect.
   NOTE: If the loading dose does not provide sufficient anesthesia for the duration of the procedure, an additional 25% of the loading dose may be injected.
2. Monitor animals post injection to check for steady respiratory rates and appropriate depression of arousal levels. Check arousal level with a hind foot pinch, which should elicit no response when sufficiently anesthetized.
   NOTE: This usually takes 3-5 min for a young adult mouse averaging 25 to 30 g. If the animal is still responsive after 10 min post injection, an additional 25% of the anesthetic loading dose may be injected.
3. Apply petrolatum and light mineral oil ophthalmic ointment to the animal's eyes to prevent dryness. Remove animals from cage and place on a clean, flat surface. Shave the desired hind limb from foot to pelvis using an electric hair trimmer, exposing only the lateral aspect of the limb.
4. Apply a chemical hair remover to the shaved site for 1 min. Manually remove the hair using laboratory wipes. Clean the depilated area with laboratory wipe soaked in ethanol.

2. Surgical Procedure

1. Sterilize surgical instruments via autoclave or other appropriate method. Clean the surgical site and surgical board with 80% ethanol/H₂0. Disinfect the surgical site with proviodine. Place mouse on surgical board and align with limb restraints. Keep the target hind limb in an anatomically natural position with the knee joint slightly extended without internal or external rotation.
2. Place animal and board under the surgical microscope. Orient to the proper incision site via palpation of superficial landmarks, specifically the bony knee joint and the ridge between the tibialis anterior and gastrocnemius muscles.
3. Make an approximately 3 cm incision through the skin using a scalpel or spring scissors while using general forceps for gripping. Make the incision perpendicular to the underlying course of the common fibular nerve.
4. Continue the incision through the superficial fascia, exposing the biceps femoris and vastus lateralis muscles. Separate these muscles by cutting through the connecting deep fascia. A 1-2 cm cut should be sufficient.
5. Retract the biceps femoris muscle caudally using mechanical retractors, revealing the common fibular nerve.
6. Trace the nerve proximally until its intersection with the tendon of the lateral head of the gastrocnemius muscle is found. Note: Exposure may require additional manipulation of the retracted skin and muscle. This intersection is used as the stable landmark for the nerve injury.
7. Grasp the nerve with a fine forceps, aligning the tips in a parallel fashion to the lateral border of the gastrocnemius tendon. Crush the common fibular nerve by applying steady, hard pressure for 5 sec.
8. Corroborate full crush of the nerve by visual inspection through the surgical scope. It will appear translucent at the site of injury. If using mice expressing fluorescence proteins in peripheral axons, the fluorescence will disappear from the site of injury.
9. Remove retractors and realign muscles in their anatomic positions. Close the incision site with 6-0 silk sutures. 1-3 simple interrupted sutures is sufficient. Place recuperating mouse on a heating pad in a clean cage.
10. Monitor all animals for 2 hr post-operation to check for breathing and any adverse reactions to the anesthesia. Administer an initial dose of buprenorphine 0.05-0.10 mg/kg via subcutaneous inguinal injection immediately following recovery from surgery. Give 3 additional doses every 12 hr over the next 48 hr. After full recovery, return mice to the animal care facility.

3. Isolation and Staining of Extensor Digitorum Longus (EDL) Muscles

1. Sacrifice animals using isoflurane. Dispense 0.5 ml liquid isoflurane into a 50 ml tube packed with absorbent labwipes. Place the uncapped tube with the animal in a sealed 2,500 cm³ chamber. At least 4 min of exposure is sufficient. Test for loss of bilateral palpebral, toe-pinch, and tail-pinch reflexes to ensure that each animal is unconscious before proceeding with perfusion.
2. Transcardially perfuse¹⁶ animals first with 10 ml 0.1M PBS, then 25 ml of 4% paraformaldehyde in 0.1M PBS (pH 7.4). Heparin (30 units/20 g animal weight) may be added with the PBS (10 units/ml) to prevent blood clotting in small capillary beds, improving perfusion results.
3. Remove the skin covering hind limbs by using scissors to cut transversally through the skin around the circumference of the abdomen. Peel down the skin past the hind limbs and feet using forceps.
4. Remove superficial fascia of hind limbs by grasping and peeling with forceps. If using mice expressing fluorescence proteins in peripheral axons, post-fix whole mice overnight in 4% PFA in 50 ml tubes. Rinse three times with PBS.
   NOTE: Fixed mice can be stored in PBS at 4 °C. If not, skip this step and proceed to step 3.6 without post-fixing animals.
5. Dissect out EDL muscles¹⁸ from mouse hind limbs, being sure to keep proximal and distal tendons as intact as possible.
6. Incubate EDL muscles in blocking buffer (1x PBS containing 0.5% Triton X-100, 3% BSA and 5% goat serum) for at least 1 hr.
7. To visualize motor axons and their nerve terminals, place muscles in tubes containing neurofilament (1:1,000) and synaptotagmin-2 (1:250) antibodies diluted in blocking buffer for 3 days. Wash muscles three times with 1x PBS and 10 min each time. Note: Skip this step if using mice expressing fluorescence proteins (XFP) in peripheral axons.
8. Stain the contralateral uninjured EDL as a positive control for complete NMJ innervation. Negative controls should include an EDL obtained at 4 days post-injury, a time point where the NMJ is completely denervated, as well as an EDL stained with secondary antibody only.
9. Incubate muscles with appropriate fluorescently tagged secondary antibodies to detect neurofilament and synaptotagmin-2 for 1 day. Wash muscles three times with 1x PBS and 10 min each time. Note: This step can be carried out together with step 3.10. Skip this step if using mice expressing fluorescence proteins in peripheral axons.
10. To visualize the postsynaptic region of the NMJ, incubate muscles with 5 µg/ml Alexa-555 conjugated alpha-bungarotoxin diluted in blocking buffer for at least 2 hr. Wash muscles three times with 1x PBS and 10 min each time.
11. To mount whole muscles on positively charged glass slides, place the muscle directly on the slide, add a few drops of glycerol based mounting medium on the slide and cover with a coverslip. Press the coverslip against the slide to flatten the muscle. Soak off mounting media from the perimeter of the slide and coverslip using laboratory wipes. Apply nail polish to seal the edges between the coverslip and slide.

4. Imaging and Data Analysis

1. To analyze the structure of NMJs, image the EDL muscle using a confocal laser scanning microscope equipped to excite 488, 555 and 633 nm light and capture the emitted light with 20X and 40X objectives.
2. To visualize whole NMJs, create maximum intensity projection images of optical sections spaced 1 to 2 μm apart extending from the lowest to highest visible regions of the NMJ. Create maximum intensity projections using commercially available imaging software.
3. To determine rates of reinnervation, categorize NMJs as: 1) completely denervated = postsynaptic site is completely devoid of contact with axon, less than 5% colocalization between the axon and AChRs. 2) Partially innervated = the axon partially overlaps the postsynapse, 5-95% colocalization between the axon and AChRs. 3) Full innervation = nearly perfect apposition between the pre- and post-synapse, greater than 95% colocalization between the axon and AChRs. Exclude NMJs that lie perpendicular to the imaging plane or are not fully visualized in the image. Note: In all these experiments, at least 3 animals and 50 NMJs per animal were examined. Results were deemed significant using a student t-test with a P value of less than 0.05.
4. To blind the operator, separate individuals can perform the surgery and image analysis. Without knowledge of the treatment groups, the analyzer can be objective with NMJ scoring. Alternatively, images can be randomized and presented to the operator for analysis without knowledge of the source animal.

5. Quantitative PCR

1. Sacrifice animals using isoflurane and cervical dislocation. Remove skin and superficial fascia covering leg muscles according to step 3.3. Dissect tibialis anterior and EDL muscles according to step 3.4.
2. Flash freeze the entire tibialis anterior and EDL muscles in a 1.5 ml tube over liquid nitrogen. Remove tissue from tube and place in a pre-chilled mortar partially submerged in liquid nitrogen. Grind frozen muscle into a fine powder using a mortar and pestle.
3. Dissolve frozen muscle powder into a commercially available RNA extraction reagent and perform RNA extraction and genomic DNA removal with a commercially available kit according to manufacturer's instructions.
4. Perform reverse transcription with a commercially available reverse transcriptase mix according to manufacturer's instructions.
5. Perform qPCR using a commercially available kit using appropriate housekeeping genes (see table of materials). Use a commercially available quantitative PCR thermal cycler to carry out PCR (see table of materials).
6. Set annealing temperature to 58 °C. Adjust additional cycling parameters to specifications of the manufacturer of taq polymerase/SYBR green mix. Include a final melt curve step in thermal cycler program consisting of 0.5 °C incremental increases from 65 °C to 95 °C to test for primer specificity and primer dimer formation.
7. Determine relative mRNA expression levels by the 2^-ΔΔCT method²¹ using 18S RNA as the control gene.

Wyniki

The common fibular nerve, also called the common peroneal nerve, arises from the sciatic nerve above the popliteal fossa, where it swings around the head of the fibula to the anterior aspect of the leg (Figure 1A). There it branches into the superficial and deep fibular nerves, together supplying the dorsiflexors of the foot and toes (anterior tibialis, extensor digitorum longus and brevis, and extensor halluces longus muscles), and the everters of the foot (peroneus musc...

Dyskusje

The method presented in this manuscript provides unique opportunities to identify mechanisms involved in repairing neuromuscular junctions (NMJ). This method involves crushing the common fibular nerve as it passes over the gastrocnemius tendon near the knee. We show that after only 5 sec of nerve compression with a forceps, complete degeneration is noted by 4 days after injury. In young adult mice, alpha-motor axons begin to reinnervate previous synaptic sites in the extensor digitorum longus muscle (EDL) at 7 days post-...

Materiały

Name	Company	Catalog Number	Comments
Ketamine	VetOne	501072
Xylazine	Lloyd Inc.	003437
Buprenorphine	Zoopharm	1Z-73000-150910
Nair	Nair
Kim-wipes	Kimtech	34155
Electric Razor	Braintree Scientific	CLP-64800
80% EtOH/H20
10% Proviodine
1 mL Insulin Syringe
Spring Scissors	Vannas	91500-09
No. 15 scalpel	Braintree Scientific	SSS 15
#5 Forceps	Dumont	11252-00
6-0 silk suture on reverse cutting needle	Suture Express	752B
Rodent Heating Pad	Braintree Scientific	AP-R-18.5
Alexa 555 conjugated alpha-BTX	Molecular Probes	B35451
Vectashield	Vector Labs	H-1000
Olympus Stereo Zoom Microscope	Olympus	562037192
Zeiss 700 Confocal Microscope	Zeiss
Variable-flow peristaltic perfusion pump	Fisher Scientific	13-876-3
Aurum Total RNA Mini Kit	Bio-Rad	7326820
Bio-Rad iScript RT Supermix	Bio-Rad	1708840
SsoFast Evagreen Supermix	Bio-Rad	1725200
Bio-Rad CFX96	Bio-Rad	1855196
Puralube vet ointment	Puralube	1621
Synaptotagmin-2 antibody	Antibodies-Online	ABIN401605
Neurofilament antibody	Antibodies-Online	ABIN2475842