A Revised Surgical Approach to Induce Endolymphatic Hydrops in the Guinea Pig

Summary

This article demonstrates an extradural approach to obliterate the guinea pig endolymphatic sac and injure the endolymphatic duct with a fine pick in order to induce experimental endolymphatic hydrops.

Abstract

Endolymphatic hydrops is an enlargement of scala media that is most often associated with Meniere's disease, though the pathophysiologic mechanism(s) remain unclear. In order to adequately study the attributes of endolymphatic hydrops, such as the origins of low-frequency hearing loss, a reliable model is needed. The guinea pig is a good model because it hears in the low-frequency regions that are putatively affected by endolymphatic hydrops. Previous research has demonstrated that endolymphatic hydrops can be induced surgically via intradural or extradural approaches that involve drilling on the endolymphatic duct and sac. However, whether it was possible to create an endolymphatic hydrops model using an extradural approach that avoided dangerous drilling on the endolymphatic duct and sac was unknown. The objective of this study was to demonstrate a revised extradural approach to induce experimental endolymphatic hydrops at 30 days post-operatively by obliterating the endolymphatic sac and injuring the endolymphatic duct with a fine pick. The sample size consisted of seven guinea pigs. Functional measurements of hearing were made and temporal bones were subsequently harvested for histologic analysis. The approach had a success rate of 86% in achieving endolymphatic hydrops. The risk of cerebrospinal fluid leak was minimal. No perioperative deaths or injuries to the posterior semicircular canal occurred in the sample. The presented method demonstrates a safe and reliable way to induce endolymphatic hydrops at a relatively quick time point of 30 days. The clinical implications are that the presented method provides a reliable model to further explore the origins of low-frequency hearing loss that can be associated endolymphatic hydrops.

Introduction

Endolymphatic hydrops is an enlargement of scala media. The presence of endolymphatic hydrops can be measured using the cross-sectional area of scala media. It is thought that clinical endolymphatic hydrops can be associated with low-frequency sensorineural hearing loss, such as that seen in Meniere's disease. But the origin(s) of the hearing loss remain unclear. To adequately study the origins of low-frequency hearing loss associated with endolymphatic hydrops, a reliable model is needed.

In 1965, Kimura and Schuknecht described how to induce endolymphatic hydrops in the guinea pig using an intradural approach¹. Their technique involved using a posterior cranial fossa approach to access the operculum and subarcuate fossa. The steps involved incising dura, retracting the cerebellum with a Ringer's solution soaked cotton pad, and drilling across the endolymphatic duct and the intermediate portion of the endolymphatic sac. Bone wax was then placed into the operculum to separate the endolymphatic duct from the distal endolymphatic sac. The craniotomy defect was closed by placing absorbable gelatin powder (e.g., Gelfoam) and reapproximating the overlying muscles. Histologic evidence of endolymphatic hydrops was consistently found at post-operative days 1, 3, 7, 14, 21, and 30, demonstrating that the intradural approach was a reliable method to induce histologically-confirmed endolymphatic hydrops. Using the same intradural approach as Kimura and Schuknecht, but with different time points, Salt and DeMott confirmed that scala media in the second turn of the cochlea was significantly enlarged at day 4 and beyond². While the actual morbidity of inducing a cerebrospinal fluid (CSF) leak using Kimura and Schuknecht's intradural approach was not reported in the original study, the presence of a CSF leak could increase the risk of meningitis. It has been suggested that loss of CSF could lead to an outflow of perilymph, resulting in a simultaneous temporary expansion of the endolymphatic volume in the guinea pig³. An extradural approach to inducing endolymphatic hydrops would be a safer option.

In 1989, Andrews and Bohmer described two extradural surgical approaches to reach the endolymphatic sac and duct, via either a middle cranial fossa approach or posterior cranial fossa approach, to obliterate the endolymphatic sac⁴. They described removing the operculum with a diamond drill, and then either drilling off the intermediate portion of the endolymphatic sac or using a fine pick to disrupt the endolymphatic sac and duct. In 1993, Lee, Wright, and Meyerhoff described a similar approach, which included drilling through the endolymphatic sac and duct, but differed in that they also simultaneously obstructed the cochlear aqueduct⁵. They demonstrated the presence of endolymphatic hydrops, as assessed via histology, at four weeks after obliterating the endolymphatic sac and obstructing the cochlear aqueduct. Megerian et al. was the first to publish a video article demonstrating an extradural obliteration of the endolymphatic sac and duct that involved drilling directly on the medial portion of the operculum to enter into the endolymphatic sac and duct⁶. They demonstrated histologic evidence of endolymphatic hydrops in a guinea pig sacrificed at 28 weeks after surgery, as well as hearing loss in the 16 kHz region⁶. Whether it was possible to induce histologically confirmed endolymphatic hydrops and low-frequency hearing loss at an early time point using extradural approaches was unknown.

The overall goal of this report is to demonstrate an extradural approach to induce experimental endolymphatic hydrops at 30 days post-operatively by obliterating the endolymphatic sac and injuring the endolymphatic duct with a fine pick. The rationale behind the use of this technique is the advantage of avoiding the need to drill on the petrous temporal bone, thereby removing the risk of accidentally injuring the dura and causing a CSF leak, mitigating the possibility of injuring the posterior semicircular canal, and reducing the risk of injury to the sigmoid sinus.

Protocol

All procedures listed immediately below in the Protocol section were conducted as described in protocols approved by the Washington University in St. Louis Institutional Animal Care and Use Committee.

1. Anesthetic Induction and Monitoring of Vital Signs

NOTE: This study used pigmented NIH-strain guinea pigs obtained from an in-house breeding colony.

1. Use guinea pigs of either sex, weighing at least 350 g.
2. Place the guinea pig in a neonatal warming isolette and give a ketamine/xylazine mixture intraperitoneally (50 mg/kg ketamine and 10 mg/kg xylazine) for induction anesthesia. Observe the guinea pig until it loses the toe-pinch reflex.
3. Once loss of toe-pinch reflexes occurs, shave the posterior neck and head of the guinea pig with a hair trimmer typically advertised for human use.
4. Inject a subcutaneous bolus of 12 mL of lactated Ringer's solution into the back of the animal.
5. Place the guinea pig supine on a warming pad with legs raised and place 27.5 G butterfly needle intraperitoneally. Verify that the butterfly needle is in the correct position in the intraperitoneal space by ensuring only air is aspirated. If blood or fluid is aspirated, there is concern for delivery into the vascular or bowel system. The butterfly needle is used for repeated administration of anesthesia.
6. Flip the guinea pig over to the prone position and secure the head to a stereotactic holder.
7. Secure a pulse oximeter to the foot. If using pigmented guinea pigs, pigmented paws can prevent reading oxygen saturation. Therefore, place the pulse oximeter on any paw that is not pigmented.
8. Insert a rectal temperature probe to monitor body temperature. The rectal probe is part of a warming blanket system that maintains body temperature at 38 °C. Do not turn on the warming blanket until the rectal probe is in place to avoid overheating the warming blanket. If having difficulty placing the rectal probe, it can be laid alongside the guinea pig's body.
9. Apply lubricant to both eyes of the guinea pig to prevent corneal abrasions.
10. Administer supplemental oxygen as needed via a rubber tubing positioned near the nose to maintain oxygen saturation levels above 90%.
11. Give enrofloxacin 0.5 mg/kg subcutaneously as an antibiotic prophylaxis.
12. Give 0.25 mg/kg bupivacaine with 1:100,000 epinephrine subcutaneously at the anticipated incision site for local anesthesia and vasoconstrictive effects.
13. Provide maintenance anesthesia every 20 min for 4 cycles and then only as needed based on vital signs. Routinely monitor the depth of anesthesia by body temperature, respiration rate, oxygen saturation, and heart rate.
14. Monitor vital signs every 15 minutes (temperature, respiratory rate, heart rate, and oxygen saturation).

2. Surgical Preparation

1. Once the head of the guinea pig is positioned securely in a stereotactic holder, place a piece of masking tape over the back to provide adequate tension along the skin overlying the occiput. Secure the ends of the tape to the stereotactic holder.
2. Thoroughly prepare the skin overlying the occiput and posterior neck with povidone-iodine followed by 70% ethanol using aseptic technique, with three alternating antiseptic applications.
3. At this point, use sterile precautions and autoclaved instruments. Place sterile drapes over the guinea pig.

3. Surgical Procedure

1. Using a 15 blade, make a small, midline incision along the posterior occiput extending down into the posterior neck. Once under the skin, use iris scissors to detach the right posterior cervical muscles from the occipital bone. If any bleeding occurs while cutting the muscles, control by applying pressure with a sterile cotton ball.
2. Using a combination of a #3 mm, #2 mm, and #1 mm diamond burr with a 5-0 suction and sterile irrigation, perform a craniotomy that is bounded by the external occipital crest, lamboidal ridge, the occipitomastoid suture line, and the dorsal margin of the foramen magnum.
   1. Gently place a small piece of saline-moistened cotton ball under the bone while separating the occipital bone from the dura.
3. Skeletonize the sigmoid sinus with a #0.5 mm diamond burr and carefully remove the bone overlying it.
4. Once the sigmoid sinus is exposed, gently retract the sigmoid sinus medially using a cotton ball and switch to using a 3-0 suction.
5. Identify the operculum as a slit like structure that is located within the petrous temporal bone. The subarcuate fossa will be situated superiorly and the sigmoid sinus will be medial to it. The extra-osseous portion of the endolymphatic sac is then visualized as a clear sac entering the operculum and attached to the dura overlying the sigmoid sinus. The operculum is oval shaped, approximately 3 to 4 mm by 1.5 to 2 mm. However, as seen from the surgical view, the operculum appears as an approximate 1 mm slit. The visible portion of the sac from the surgical view is approximately the same size as the visible portion of the operculum, if not smaller.
6. Apply gentle retraction to the sigmoid sinus medially in order to clearly visualize the extra-osseous portion of the endolymphatic sac and increase the tension between the extraosseous and intraosseous portions of the endolymphatic sac.
   1. Use a fine angled pick to gently expunge the intermediate portion of the endolymphatic sac. It is critical that the expungement process leave no visible connection between the dura and the operculum; then place a fine pick inside the operculum to broadly scrape along the inside of the bone to injure it.
   2. Turn the fine pick in the direction of the endolymphatic duct and blindly disrupt the lining. At this point some bleeding may occur from a vessel within the operculum. It can be controlled with a small piece of cotton.
7. Dry the empty operculum with a small piece of cotton. Using the 3-0 suction as needed to keep the cotton dry.
8. Obtain bone dust by using a small curette to scrape along the squamosal portion of the temporal bone. Generously pack the operculum with bone dust. Use a cotton ball and suction to keep the area dry while packing it with bone dust.
9. Apply bone wax to the operculum to seal it. Ensure that there is no excess bone wax dislodged into the skull.
10. Use bone wax to cover the skull defect.
11. Approximate the posterior cervical muscles with 4-0 braided, absorbable suture in an interrupted fashion.
12. Perform a subcuticular closure using a 4-0 braided, absorbable suture.

4. Post-procedure Care

1. Remove the guinea pig from the custom stereotactic holder and transfer to a warming isolette.
2. Give 2 mg/kg Atipamezole and 24 mL of lactated Ringer's solution (subcutaneously away from the incision). Give lactated Ringer's solution due to the diuretic effects of xylazine. Administer 0.2 mg/kg meloxicam subcutaneously for post-operative analgesic coverage.
3. Obtain vital signs every 15 minutes until the guinea pig fully emerges from anesthesia.
4. Give an additional 12 mL fluid bolus of lactated Ringer's solution about 2 hours from end of surgery during the recovery period.
5. Once the guinea pig is alert, ambulating, voiding, and having bowel movements, return the guinea pig to the animal facility. Approximately 2 to 4 hours are needed for the guinea pig to emerge completely from anesthesia.
6. Monitor guinea pigs twice daily for the first three post-operative days. If signs of discomfort are observed, administer 0.2 mg/kg meloxicam subcutaneously every 24 hours as needed. Alternatively, buprenorphine (0.05 mg/kg) may be administered subcutaneously if symptoms are not sufficiently managed by meloxicam. 
7. Give a 12 mL fluid bolus of lactated Ringer's solution subcutaneously twice a day for up to three days until the guinea pig reaches the pre-operative weight. If the guinea pig reaches its pre-operative weight prior to the third post-operative day, then stop fluid boluses. If the guinea pig continues to lose weight after the first three days, use a supplement nutrition shake typically advertised for human consumption mixed with crushed guinea pig food pellets.
8. Monitor guinea pigs weekly until their end point.

Results

The presented method used an extradural approach to obliterate the endolymphatic sac and injure the endolymphatic duct with a fine pick in seven guinea pigs consisting of two males and five females. The average duration of surgery was 2 hours from incision to closure. The total drill time ranged from 5-10 minutes. Up to 4 hours was needed for the guinea pig to fully emerge from anesthesia. There were no intra-operative or post-operative deaths in the sample. There were no injuries to the ...

Discussion

The presented extradural method had a success rate of 86% in achieving histologically confirmed endolymphatic hydrops and low-frequency hearing loss. The method reliably achieved histological evidence of endolymphatic hydrops by post-operative day 30, consistent with prior studies that used an intradural approach². The significance of the method with respect to existing methods is that a CSF leak is not required, thus removing a potential confounding variable that has been suggested to result in a...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
12 mL syringe	Henke-Sass Wolf	5100-X00V0
1 mL and 3 mL syringe	BD Precision(Ordered from Fischer Sci)	14-826-87 15859152
27.5 butterfly gauge needle	Terumo Surflo Winged Infusion Set, Terumo Corporation, Japan) (Ordered from McKesson)	448407
4-0 suture	McKesson	1034507
4 x 4 gauze sponges	Dukal (Ordered form McKesson)	374454
60 mL syringe	Fisher Sci	22-031-375
Anspach otologic drill	Anspach	SC2100
atipamezole	Zoetis	107204-6
autoclave	Fisher sci	15-103-0508
autoclave bags	McKesson	524881
bayonet separator	Olympus	AL 130564
bupivicaine	auro Medics Pharma	555150-169-10
clear sterile drape	3M	1020
cotton balls	Fisherbrand (ordered from Fisher Sci)	22-456-885
cotton swabs	McKesson	508716
diamond burrs #3, #2, #1, and #0.5 mm	Anspach	QD8-3SD; QD8-2SD; QD8-1SD; QD8-05SD
diaper pad	McKesson	945330
disposable 15 blade	Swann-Morton	0305
enrofloxacin	Hospira	0409-4888-01
epinephrine	McKesson	63739-0456
eye ointment	Dechra Vet Products	17033-211-38
Freer elevator	Grace Medical	215100FX
gelfoam	Pfizer (Ordered from McKesson)	82830
hair trimmers	Oster Power Pro Cordless (ordered from Amazon)	078400-020-000
iodine scrub	Purdue Pharma (ordered from mcKesson)	521243
iris scissors	Olympus	CL-542114
ketamine	Henry Schein Animal Health	55853
lactated ringers	B. Braun Medical (ordered from McKesson)	186662
lancet knife by Rosen	Grace Medical	151100FX	referred to as curette in the text
lubricant	Milex (ordered from Cooper Surgical)	MX5030
masking tape	3M (ordered from fisher sci	19047259
metal rectangle basin	Amazon	B07NQDBC6T
needle holder	Olympus	CR 213015-ENT
needles: 27 gage, 18 gauge	BD Precision(Ordered from Fischer Sci)	14-826-48 14-826-5D
neonatal warming isollete	Air Borne Life Support Systems	731-1800
operating microscope	Carl Zeiss	OPMI pico
oxygen tank	AirGas	OX USP200
pulse ox	CapnoTrue (Ordered from Medacx)	M-3090112001
rectal probe with heating blanket	Harvard Apparatus	probe: PY2 50-7217 Heating Blanket: PY2 50-7214
red body holder	Lichtenhan Lab	N/A	In-house product
right angle	Olympus	BV-230337
rosen needle	Olympus	AM-130566	customized, it is the instrument I use to tear the sac
rubber tubing for O2 administration	Fisher Sci	14-171-104
saran wrap	Fisher Sci	NC9617977
stereotactic head holder	WUSTL Instrument Machine Shop	N/A	In-house product
sterile drapes	Cardinal Health	7553
suction tube by Baron	Grace Medical	034903FX 034905FX	#3 and #5 Suction
tissue forceps adson brown	Grace Medical	325112FX
Weitlander retractor	Olympus Grace Medical	BL200011 100313FX
xylazine	Akorn	59399-110-20