Subcutaneous Neurotrophin 4 Infusion Using Osmotic Pumps or Direct Muscular Injection Enhances Aging Rat Laryngeal Muscles

Summary

Here, we present a protocol to describe the use of neurotrophin 4 (NTF4) systemically and directly to remodel rat aging laryngeal muscles.

Abstract

Laryngeal dysfunction in the elderly is a major cause of disability, from voice disorders to dysphagia and loss of airway protective reflexes. Few, if any, therapies exist that target age-related laryngeal muscle dysfunction. Neurotrophins are involved in muscle innervation and differentiation of neuromuscular junctions (NMJs). It is thought that neurotrophins enhance neuromuscular transmission by increasing neurotransmitter release. The neuromuscular junctions (NMJs) become smaller and less abundant in aging rat laryngeal muscles, with evidence of functional denervation. We explored the effects of NTF4 for future clinical use as a therapeutic to improve function in aging human laryngeal muscles. Here, we provide the detailed protocol for systemic application and direct injection of NTF4 to investigate the ability of aging rat laryngeal muscle to remodel in response to NTF4 application. In this method, rats either received NTF4 either systemically via osmotic pump or by direct injection through the vocal folds. Laryngeal muscles were then dissected and used for histological examination of morphology and age-related denervation.

Introduction

Laryngeal muscles contract rapidly and consistently, and are vulnerable to the adverse effects of aging. This constant activity is thought to contribute to voice problems or dysphagia observed in persons over 65 years of age ^{1,2,3,4,5,6,7}. Several molecular and pathophysiologic mechanisms contribute to this age-related dysfunction. These mechanisms can include remodeling of laryngeal mucosa, muscle fiber atrophy or loss, lack of muscle fiber regeneration or atrophy which causes bowing of the vocal folds and inability of glottis closure ^8,9,10,11. There is no proven medical therapy at this time that can completely prevent or rehabilitate these age-related changes in these muscles.

Modulation of the effectiveness of neuromuscular transmission can greatly influence neuromotor performance. The family of neurotrophins include nerve growth factor (NGF), brain derived nerve growth factor (BDNF), neurotrophin 3 (NTF3) and NTF4 ^12,13. Neurotrophins have been shown to modulate synaptic efficacy^1,4. Hepatocyte growth factor, transforming growth factor beta and fibroblast growth factor have recently been used in humans for the treatment of vocal fold scarring ^15-17. NTF4 also regulates NMJ effectiveness; mice lacking NTF4 show disassembled NMJs ^11,18,19. These studies lead to promising effects of treatment of aging laryngeal muscle disorders and denervation with growth factors.

Direct injection therapeutics to the tissues of the vocal folds are not unprecedented in humans. For example, local injections of botulinum toxin are currently used as an effective treatment for neurological movement disorders that affect the muscles in the larynx, such as spasmodic dysphonia and bilateral recurrent laryngeal nerve paralysis ^20,21. Hyaluronic acid hydrogel is another injectable, which is used to treat vocal fold scaring and glottal insufficiency ^22,23. Injection laryngoplasty can be used to treat a variety of communication disorders ²⁴. These direct injection methods hold promise to improve vocal function and swallowing in aging populations.

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Protocol

Use male Fischer 344-Brown Norway rats at 6 and 30 months of age for this protocol. Rats were obtained from the National Institute of Aging rodent colony. We used rats for this study because the structure of the rat larynx is similar to that of the human, functionally serving for airway protection and species-specific vocalizations This study was performed in accordance with the PHS Policy on Humane Care and Use of Laboratory Animals, the NIH Guide for the Care and Use of Laboratory Animals, and the Animal Welfare Act (7 U.S.C. et seq.); the animal use protocol was approved by the Institutional Animal Care and Use Committee (IACUC) of the University of Kentucky.

1. Anesthesia of Rats

1. Prepare the anesthetics by mixing ketamine hydrochloride (dissociative anesthetic) and xylazine hydrochloride (sedative and analgesic) in buffered saline. The concentrations of ketamine and xylazine in the final solution are 100 mg/8 mg per kg body weight respectively.
2. Inject the anesthetics into the rat by intraperitoneal administration using a syringe with a 25 G needle.
3. Determine that the rat is sufficiently anesthetized by pinching the toe or foot with forceps. If the rat does not react to the pinch, then surgery can begin. If the rat reacts to the toe pinch with reflex or muscle contractions, then wait 1-2 min and repeat the pinch test. If the rat reacts again, replace the rat with a new animal and repeat procedure beginning from step 1.2.
4. Apply ophthalmic ointment to the rat's eyes, after the rat is immobile, to prevent the corneas from drying out.

2. Osmotic Pump Implantation

1. Place the rat ventral on the aseptic surgical area.  Administer meloxicam as a preanesthetic medication. Administer intraperitoneally at a dosage of 1-4 mg/kg of body weight using a syringe with a 25 G needle.
2.  Use clippers to remove an approximately 1" x 1" square of fur from back of neck, and approximately 1" caudal of space between shoulders. Shave as close to the skin as possible.
3. Wet the back and neck with disinfecting ethanol (70%).
4. After shaving, scrub the dorsal aspect of the neck with 3 scrubs in succession of iodine-alcohol finishing with alcohol.
5. Maintain the body temperature of the rat by placing it on a heating pad set to 34 °C.
6. Fill aseptically prepared osmotic pumps with either 50 µL of NTF4 or saline for systemic NTF4 treatment (Figure 1).
   1. Use a scalpel to make a horizontal incision approximately 2 cm wide through the skin, just cranial to the space between the scapulae. Lift the posterior edge of the incision with forceps with one hand while inserting the tip of hemostats and gently pushing posterior to the incision.
   2. After the tip of the hemostats is approximately 2 cm cranial to the incision, open the handles on the hemostats, expanding the tips to form a hollow "pocket" subcutaneous to the incision site. This will be the placement site for the pump.
7. Orient the pump delivery portal end first upon insertion to minimize any interaction of the NTF4 and the healing of the pocket incision site.
8. Deliver 50 µL of NTF4 saline for either 7 14 days. The 7 day group received 6.72 mg/day of NTF4 for a total dose of 47.04 mg. The 14-day group received 6.72 mg/day for a total dose of 94.08 mg of NTF4²⁵.
9. Use 5-0 nylon suture thread, hemostats and forceps to close the incision made for pump placement.
10. Observe the rats for a minimum of 30 min as they recover from anesthesia. Criteria for completion of monitoring include the animal becoming active, moving about the cage, drinking water, and beginning other normal activities such as grooming.
11. Monitor animals daily for the first week by observing the healing of surgical site, normal feed and water consumption and passing of urine/feces, and any abnormal behavioral signs of stress, pain, or other post-operative complications.
12. If the rat appears to be in pain or distress, provide the rat with a 5 mg/kg subcutaneous injection of carprofen once every 24 h for up to 5 days to relieve pain.
13. If there appears to be an infection, consult a veterinarian to ensue that the wound heals properly.
14. Depending on which experimental group the rat in in, remove the 5-0 nylon suture 7-10 days following surgery to prevent irritation from the thread.

3. Anesthesia of Rats for Direct Injection

1. Withhold food from the rats the night before the procedure. This ensures that there is no food to block the endoscope or injection needle.
2. Weigh rats and prepare acepromazine 1-2 mg/kg body weight. Inject intramuscularly (the IM location is the left thyroarytenoid muscle).
3. Place the rat in the induction box. Induce anesthesia in the induction box with 5% isoflurane and 1 L O₂.
4. Move the rat to a nose cone with 2% isoflurane and 600 mL O₂.
5. Determine that the rat is sufficiently anesthetized by pinching the toe or foot with forceps. If the rat does not react to the pinch, then the injection protocol can begin. If the rat reacts to the toe pinch with reflex or muscle contractions, then wait 1-2 min and repeat the pinch test. If the rat reacts again, replace the rat with a new animal and repeat procedure beginning from step 3.4.

4. Direct Injection and Visualization

1. Place aseptically prepared 50-µL dosages containing NTF4 or saline in a H₂O bath set to 25° C for 30 min before injection.
2. Place the rat in a supine and reclined position on a plexiglass platform (Figure 2). Suspend the rat in the reclined posture from their frontal top incisors via a guide wire strung across the top of the platform.
3. Attach a 50 mm, 30 gauge, 100 µL syringe to a 1.9 mm, 30° sinus endoscope (Figure 3).
   NOTE: The syringe assembly is attached via a jig that holds the cannula firmly to the outer wall of the endoscope. The endoscope allows for visualization of the vocal folds and guidance of the syringe intraorally. The position of the cannula tip is adjusted prior to each animal to ensure that the tip is fully and clearly visible via the endoscopic view (Figure 4).
4. Use a rubber-tipped pair of forceps to extend the tongue and move it laterally. Afterward, insert a plastic speculum to maintain oral patency. Make the speculum from a 5 mL plastic syringe barrel that is cut to a length of 1.5 to 2 cm, with the cut edges deburred and polished smooth.
5. Turn off the lights in the room and attach a halogen light source to the endoscope. Turn on the video recorder to capture the procedure.
6. Immerse the distal end of the endoscope in warm water for a few seconds to minimize the development of condensation on the glass tip when inserted into the mouth of the rat.
7. Using visual feedback from the monitor, carefully guide the needle to the area of the left vocal fold.
8. Time the injection of the solution with the inspiratory phase of the animals' respiration cycle to fully access the vocal fold. During the inspiratory phase of respiration, the vocal fold is fully exposed.
   1. Once the vocal fold is fully visible, insert the needle into the left thyroarytenoid, found lateral to the white medial edge of the vocal fold. With the needle in place, deliver the injectate through depression of the syringe.
9. Turn off the halogen light source on the endoscope and the video player, and turn back on the room lights.
10. Return the rat to its home cage and place on a heating pad.
11. Allow the rat to recover before removal from the heating pad. Replace food and water in the cage.
12. Monitor rats for 7 days after the injection and then euthanize. Remove the larynges for cryosectioning²⁴.

5. Euthanization of Rats

1. Anesthetize rats with ketamine hydrochloride and xylazine hydrochloride (100 mg/8 mg per kg body weight injected intraperitoneal injection).
2. Euthanize by exsanguination following a medial thoracotomy.

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Results

The rats were euthanized after 2 weeks of osmotic pump infusion or 1 week after direct injection of NTF4. Larynges were harvested, placed in cryoprotectant (30% sucrose and 70% phosphate buffered saline) and then serially sectioned in 10- µm widths with a cryostat. Aging laryngeal muscles are affected by administration of NTF4 ²⁵. In addition to young and old rat, we compared the injected and non-injected side of the thyroarytenoid muscles. Typically we see a ...

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Discussion

Laryngeal muscles are vulnerable to the unfavorable effects of aging. Previous studies have demonstrated changes in aging laryngeal muscles that include changes in fiber size, total number of fibers, regenerative ability, NMJ size and quantity changes, in addition to variations in contractile function and myosin isoform shifts ^{4,11,27,30,31}. Aging laryngeal mu...

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Materials

Name	Company	Catalog Number	Comments
Neurotrophin 4	Pepro Tech	450-04	200 ng in 50 μL
Alzet Osmotic Pump	DURECT Corporation	2001D
30 ° endoscope	Stoltz	61029D
50 mm 30 gauge 100-μL syringe	Hamilton	84850 and 201812
saline (sodium chloride solution)	Sigma-Aldrich	S8776
ketamine hydrochloride	Henry Schein	56344
xylazine hydrochloride	Henry Schein	33198
25 G 5/8 needle	Becton-Dickinson	305901
1 mL syringe	Becton-Dickinson	309659
ophthalmic ointment	Henry Schein	8897
clippers	Oster	44-018
ethanol	Decon	2716
iodine (Betadine)	Purdue Pharma L.P.	606404
heating pad	Sunbeam	731-5
5-0 nylon suture thread	AD Surgical	PMN-518R6
crile hemostat	Fine Science Tools	13005-14
delicate suture tying forceps	Fine Science Tools	11063-07
meloxicam	Henry Schein	49756
carprofen	Merritt Veterinary Supplies	148700
antibiotic ointment	Henry Schein	57110
acepromizine Aceproject	Henry Schein	3845
isoflurane Isothesia	Henry Schein	50033
induction box (anesthetizing box)	Harvard Apparatus	50-0116
oxygen compressed tank	Scott Gross	UN1072
plexiglas platform	Small Parts Inc (Amazon)
rubber tipped forceps	Fine science tools rubber	11075-00
liquid rubber for forceps above	Lowe's	42518
plastic spectula (BD syringe cut to length)	Becton-Dickinson	309659
halogen light source rhino-laryngeal stroboscope	Kay-Pentax	RLS 9100 B
video recorder	Kay-Pentax
sucrose	Sigma-Aldrich	S0389-500G
phosphate buffered saline	Sigma-Aldrich	P4417-100TAB
cryostat Mictotom HM525	Thermo Scientific	HM 525
Gill 1 hematoxylin	VWR	10143-142
Shandon eosin-Y alcoholic	Thermo Fisher Scientific	6766007
anti-sodium channel Nav1.5 antibody produced in rabbit	Sigma-Aldrich	S0819
Texas red-X phalloidin	Sigma-Aldrich	T7471
alpha- bungarotoxin alexa fluor 488 conjugate	Thermo Fisher Scientific	B-13422
Small animal anaesthesia machine	Smiths Medical	CDS 9000