Delivery of a Fluorescent Dye into the Subretinal Space of a Mouse

Abstract

Source: Parikh, S., et al. An Alternative and Validated Injection Method for Accessing the Subretinal Space via a Transcleral Posterior Approach. J. Vis. Exp. (2016)

This video demonstrates the procedure for delivering a fluorescent dye into the subretinal space of a mouse. The process involves anesthetizing the mouse, preparing the eye, making precise incisions to access the subretinal space, and injecting the dye to create a controlled retinal detachment. This technique allows for targeted delivery of the dye, enabling detailed imaging and analysis of retinal structures.

Protocol

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

Animals: Wild type C57Bl/6J mice bred at the University of California at Los Angeles (UCLA). All animals were between 11 - 17 weeks old and included male and female mice. All mice were group-housed, maintained in a 12:12 light/dark cycle with food and water ad libitum.

NOTE: All drugs and injectable agents are United States Pharmacopeia (USP) grade.

1. Surgical Preparation

1. Anesthetize the mouse with an intraperitoneal injection of 100 mg/kg ketamine and 8 mg/kg xylazine in a saline mix. Administer anesthesia to a depth such that the mouse has no toe pinch or corneal touch reflexes.
2. Maintain body temperature at 37.0 °C with a circulating water pad.
3. Dilate pupils with 2.5% phenylephrine eye drops and trim whiskers to facilitate visualization. Whiskers provide significant sensory input to the mouse; therefore, whisker trimming should remove only the portion that blocks clear access to the eye, and not to the base of the whisker. In our experience, mice show normal recovery after this procedure. Apply methylcellulose eye drops to prevent dryness and minimize anesthetic induced transient cataracts.
4. Sterilize instruments prior to surgery (i.e., betadine and ethanol or hot beads).
5. Prepare diluted fluorescein (0.01% using 0.9% saline) in a sterile environment (i.e., biosafety cabinet) if visualization will be performed (see section 3 below).

2. Injection Site Preparation

1. Prepare a syringe (e.g., 5 µl syringe) with the appropriate injection volume (e.g., 0.3 to 1.0 µl).
2. Position the mouse so the eye is facing up and clearly visible in the dissecting microscope.
3. Gently pinch the temporal conjunctiva with fine-tipped forceps. Make a circumferential incision of approximately 90 degrees using curved Vannas scissors.
4. Repeat step 2.3 with the underlying Tenon's capsule.
5. Resect the surrounding connective tissue with fine-tipped forceps while rotating the globe nasally. Work towards the injection site approximately 0.5 mm temporal to the optic nerve. Use great care to avoid disrupting the retro-orbital sinus.

3. Sclerotomy and Subretinal Injection

NOTE: It is recommended that the injection of 0.01% fluorescein in 0.9% Saline be used to assist with visualization while learning this procedure. The topographic distribution of fluorescein can be effectively documented with fundus imaging (see section 4 below).

1. Make a small scleral incision at the injection site by gently scratching the eyecup with a 22.5-degree ophthalmic blade. This incision should only be large enough to allow the tip of the needle to pass through the sclera.
2. Insert the beveled 33 G needle (angled 5 - 10° into the sclerotomy with the bevel facing and angled parallel to the retina. Inject desired volume (e.g., 0.3 to 1.0 µl of 0.01% Fluorescein for learning purposes).
   NOTE: Maintain sterility of the syringe by thoroughly cleaning with successive washes of a suitable solvent and DI water before each injection.
3. Depress the plunger slowly (over ~ 3 sec) without moving the needle and with even pressure.
   NOTE: When the needle is in the subretinal space, a slight resistance will be felt while depressing the plunger. There will be no to minimal resistance if the needle punctures the retina, and high resistance if the needle does not penetrate the sclera or retinal pigment epithelium (RPE).
4. Wait several seconds before withdrawing the needle to minimize backflow.
5. Rinse the eye with sterile buffered saline and ensure the eye has rotated back to its normal position.

4. Assessment of Retinal Detachment by Optical Coherence Tomography (OCT) and Fundus Imaging

1. Perform OCT imaging immediately after injection to evaluate the quality of the injection and at appropriate time points post-injection as needed to evaluate retinal structure.
   1. Adjust and align the OCT image to target the site of injection. The injection site should be midline and 0.5 mm temporal to the optic nerve head. Repeat as needed if the detachment is out of frame or not optimally centered.
2. Visualize retinal detachment and dye injection area with en-face fundus imaging.
   NOTE: If an OCT imaging system is not available, injection of a small amount of fluorescein with a vector for practice will allow visualization with any fundus camera that performs fluorescein angiography using the same excitation wavelengths and blocking filters. Localized areas of hyperfluorescence will appear underneath the vasculature, and the vasculature will have sharp and distinct boundaries if the subretinal space is targeted correctly. The edge of the bleb from the injection will be demarcated by the transition from hyper- to hypo-fluorescence. Several instruments provide this capability for the mouse; the instrumentation used here is described elsewhere.

5. Post-operative Care

1. Apply a thick coat of triple antibiotic ophthalmic cream to the corneal surface of the injected eye.
2. Place mice in clean solitary cages for recovery. Do not combine mice that have undergone surgery until they are fully recovered.
3. Monitor respiration and temperature during anesthesia recovery. Monitor animals until they can maintain sternal recumbency.
4. Perform additional appropriate post-operative monitoring and treatment, including a subcutaneous injection of carprofen (5 mg/kg) for post-surgical pain management.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Hamilton Model 62 RN SYR	Hamilton	87942	Syringe x 1
Hamilton Needle 33 G, 1.0", 20 DEG, point 3 (304 stainless steel)	Hamilton	7803-05	Needles x 6
Vannas Curved Scissors	Ted Pella, INC.	1347	5 mm Blade
22.5 Degree Microsurgery Knife	Wilson Ophthalmic Corp.	91204
Ketaject	Phoenix	NDC 57319-609-02	Ketamine
Anased	Lloyd Laboratories	NDC 61311-482-10	Xylazine
Fluorescein 10% AK-Fluor	Akorn	NDC 17478-253-10	100 mg/ml
0.9% Saline USP	Hospira	NDC 0409-4888-50	0.9% NaCl
Antibiotic Ointment	Akorn	NDC 17478-235-35	Ophthalmic
Water Circulating Pump	Gaymar	TP-500 T/Pump	P/N 07999-000
sd-OCT	Bioptigen	R-Series	Commercial
Fundus Camera	Phoenix Research Laboratories	MICRON III
Tweezers Type 3	Ted Pella, INC.	5385-3SU
2.5% Phenylephrine	Paragon BioTeck	NDC 42702-102-15	Ophthalmic
Hypromellose 2.5%	Goniovisc	AX0401	Methylcellulose
Eye Drops (Rinse)	Bausch & Lomb		Saline Solution
Microscope	Zeiss	Stemi 2000	Microscope
Light source	Fostec	P/N 20520	Light source