Modeling Cataract Surgery in Mice

Summary

This method models cataract surgery in vivo by removing lens fiber cells from adult mice and leaving behind the capsular bag with attached lens epithelial cells (LECs). The injury response is then assessed at various times post-surgery using molecular and morphological criteria.

Abstract

Cataract surgery (CS) is an effective treatment for cataracts, a major cause of visual disability worldwide. However, CS leads to ocular inflammation, and in the long term, it can result in posterior capsular opacification (PCO) and/or lens dislocation driven by the post-surgical overgrowth of lens epithelial cells (LECs) and their conversion to myofibroblasts and/or aberrant fiber cells. However, the molecular mechanisms by which CS results in inflammation and PCO are still obscure because most in vitro models do not recapitulate the wound healing response of LECs seen in vivo, while traditional animal models of cataract surgery, such as rabbits, do not allow the genetic manipulation of gene expression to test mechanisms. Recently, our laboratory and others have successfully used genetically modified mice to study the molecular mechanisms that drive the induction of proinflammatory signaling and LEC epithelial to mesenchymal transition, leading to new insight into PCO pathogenesis. Here, we report the established protocol for modeling cataract surgery in mice, which allows for robust transcriptional profiling of the response of LECs to lens fiber cell removal via RNAseq, the evaluation of protein expression by semi-quantitative immunofluorescence, and the use of modern mouse genetics tools to test the function of genes that are hypothesized to participate in the pathogenesis of acute sequelae like inflammation as well as the later conversion of LECs to myofibroblasts and/or aberrant lens fiber cells.

Introduction

The lens is a highly organized, transparent tissue that refracts light to produce a clearly focused image onto the retina^1,2,3. This specialized organ is surrounded by an uninterrupted basement membrane (the capsule), which isolates the lens from other parts of the eye. The inner anterior surface of the capsule anchors a monolayer of lens epithelial cells (LECs), which then differentiate at the lens equator into lens fiber cells, which comprise the vast majority of the lens³. A cataract occurs when the lens loses its transparency due to factors like aging, genetic mutation, UV radiation, oxidative stress, and ocular trauma⁴. Cataract is a leading cause of blindness worldwide, particularly in countries with poor medical care⁵. However, this condition is now readily treated by surgical removal of opaque lens cells by phacoemulsification in which the central lens capsule and attached lens epithelium are removed, followed by the use of a vibrating probe to break the cellular mass of the lens into smaller fragments that can be suctioned out; leaving behind a capsular bag with some attached equatorial LECs¹. Vision is then most commonly restored by post-surgical implantation of an artificial intraocular lens (IOL).

While cataract surgery (CS) is a highly effective and minimally invasive treatment for cataracts, a patient's post-surgical recovery can be acutely hampered by the development of ocular inflammation^5,6,8. This inflammation can cause post-surgical pain, retinal edema leading to retinal detachment, as well as exacerbation of other inflammatory and fibrotic conditions like uveitis and glaucoma^4,7,8,9. Ocular inflammation post-CS (PCS) is generally treated with anti-inflammatory eye drops which are plagued by poor patient compliance or dropless cataract surgery that can lead to an increased prevalence of floaters^8,10,11. In the long term, the outcomes of CS can be compromised by the development of posterior capsular opacification (PCO)¹². PCO occurs when residual LECs left behind after surgery undergo a wound healing response, proliferate, and migrate onto the posterior lens capsule at months to years PCS contributing to secondary visual obstruction^13,14.

Understanding the molecular mechanisms by which CS results in such acute and chronic responses represents a major challenge in the field, as much of the literature investigating PCO relies on the induction of LEC conversion to myofibroblasts in culture via treatment with active transforming growth factor beta (TGFβ)^12,15. Human capsular bag models created from cadaver lenses cultured in vitro better reflect the biology of the lens PCS as LECs are cultured on their native basement membrane but are difficult to manipulate mechanistically, do not recapitulate the intraocular environment, and are inherently variable due to lens to lens (or donor to donor) variability^16,17. Rabbit^1,18 and non-human primate^19,20 in vivo models of cataract surgery overcome some of these issues but still are not easy to manipulate for mechanistic studies. Notably, a prior study of mammalian lens regeneration found significant lens regeneration within four weeks after lens fiber cells were removed from mice, leaving the lens epithelial cells and capsule behind, while no lens regrowth occurred when the entire lens was removed^21,22. We subsequently streamlined this procedure and optimized it for the study of the acute response of LECs to lens fiber cell removal and their subsequent conversion to a mixed population of cells with myofibroblast and fiber cell properties.

Using this mouse model of cataract surgery, we have shown that lens epithelial cells drastically remodel their transcriptome by 6 hours PCS to produce numerous proinflammatory cytokines²³, while they begin expressing fibrotic markers as early as 24 h PCS^12,23, prior to the onset of canonical TGFβ signaling. Mechanistic experiments in knockout mice using this model revealed that cellular fibronectin expression by LECs is required for a sustained fibrotic response PCS, likely due to both its role in the assembly of the fibrotic ECM and cell signaling¹². Other studies showed that αVβ8-integrin is required for the transition of LECs to myofibroblasts due to its function in TGFβ activation, and the potential of this approach to identify anti-PCO therapeutic leads was confirmed as a αVβ8-integrin antagonist also blocked LEC EMT¹⁵.

Here, we provide a detailed protocol describing how to remove the lens fiber cells from living mice (Figure 1and Figure 4) while leaving behind the lens capsule and LECs to model the LEC response to cataract surgery. 

Protocol

The following protocol was approved by the University of Delaware Institutional Animal Care and Use Committee under protocol #1039-2021-1. In compliance with the Association for Research in Vision and Ophthalmology (ARVO) for the Use of Animals in Ophthalmic and Vision Research²⁴, all ocular survival surgeries can only be performed in one eye. See Table of Materials for details about reagents and instruments used in this protocol. Alternative approaches for extracting the lens fibers and suturing are shown in Figure 2  and Figure 3.

1. Animals

1. Use any wild-type or genetically modified laboratory mouse strain that has eyes with intact lenses for this procedure.
   1. Use C57BL/6 mice as a starting point, as their responses are highly reproducible at both the morphological and molecular levels^25,26.
      NOTE: C57BL/6 or any mouse strain can be used. In the video protocol, the FVB mouse is used for better visual demonstration.
   2. Perform studies of the response of mutants on other mouse strains with strain-matched controls.
      NOTE: With practice, this procedure may be performed on mice whose lenses have cataracts as long as the lens capsule is intact and the lens has not undergone significant liquefaction. See the study of mice with lens conditional deletion of the transcription factor Sip1²⁷.
2. Perform this surgery on mice of either sex as young as 8 weeks to as old as 24 months^25,26.
   1. Perform this surgery in even younger mice (after eyelid opening) with practice, although it will be more difficult as these eyes will be significantly smaller.
   2. Take particular care for surgeries on elderly mice due to their lower tolerance for anesthesia^25,26.
      ​NOTE: Sex differences in surgery responses are more prevalent in elderly mice²⁶.

2. Preparation of solutions and surgical tools

1. Obtain anesthetizing and analgesic agents, dilation drops, sterile disposable scalpel, #5 Dumont forceps, 2-110 suture forceps, needles (26 G 1/2 and 27 G 45° bent dispensing tip 1"), syringes, and balanced saline solution (BSS).
2. Sterilize surgical instruments in an autoclave.

3. Prior to anesthesia

1. Prior to anesthesia, give the mouse one drop each of 1% atropine and 2.5% phenylephrine to dilate the pupil (see Figure 1A) of the eye undergoing surgery. Apply ophthalmic ointment to the non-operated eye to keep the surface from drying.
   NOTE: The ARVO statement on the use of animals in eye research²⁴ only allows a single eye to undergo survival surgery, leaving the other eye untouched so that the animal retains vision. Thus, survival surgeries are always unilateral; however, the contralateral, unoperated eye can be operated on just prior to sacrifice to create animal-matched 'time zero' controls.
2. Make sure the eye drops do not run down the mouse's face and enter the nose, as this can obstruct its breathing.
3. Prepare a syringe with sterile BSS using a sterile 26 G 1/2 needle. Fit the syringe with a sterile 27 G 45° bent blunt dispensing tip of 1 inch (referred to as a dispensing tip below).

4. Anesthesia

1. Following eye drops, anesthetize the mouse with an intraperitoneal injection of xylazine/ketamine/acepromazine (35 mg/kg, 80 mg/kg, and 0.5 mg/kg) solution (Figure 1B).
   NOTE: Inhaled anesthesia might not be possible unless a specialized nose cone can be utilized to allow for access to the eye needed for surgery.

5. Making the incision

1. Once the mouse is completely under anesthesia, as measured by the lack of toe pinch reflex, and its pupil has dilated completely (2 mm) (Figure 4A), the mouse is ready for surgery.
   1. Move the anesthetized mouse to the stage of a compound dissecting light microscope. Use a heater pad to ensure the mouse has adequate thermal regulation and use sterile drapes.
   2. View the eye under 10x-20x magnification using a dissecting microscope (see Table of Materials for specifics).
2. Make a 1-1.5 mm incision in the center of the cornea using a sterile surgical scalpel. Ensure the incision extends through the cornea and incises the anterior lens capsule (Figure 1C and Figure 4B).

6. Separating the lens fibers from the capsular bag

1. Grasp the syringe with a dispensing tip containing sterile BSS and gently expel 1 drop of BSS above the cornea to wet the surface for further manipulation.
2. Separate the lens fiber mass from the lens capsule.
   1. Insert the dispensing tip through the central corneal incision into the lens capsule incision, then gently move the needle in a circular motion while slowly releasing BSS to separate apical-apical connections between lens epithelial and fiber cells (hydro dissection), ultimately expelling the lens fiber mass (Figure 1D).
   2. In most cases, the fiber mass will eventually adhere to the dispensing tip during this procedure (Figure 2A). If this happens, gently remove the dispensing tip from the inside eye; and clean with a sterile towel to remove this tissue. Repeat until all lens fiber cells are removed.
   3. Alternatively, use micro forceps (number 5 Dumont) to extract the lens nucleus (Figure 2B). If using this method, be careful that the original corneal incision exceeds the diameter of the nucleus to ensure easy removal.
3. Rinse the lens capsule with sterile BSS to remove any remaining fibers.
4. Confirm that the lens capsule is left inside the eye (Figure 1E and Figure 4C) by viewing its translucent/glassy appearance through the microscope.

7. Inflating the anterior chamber and suturing the corneal incision

1. Inflate the anterior chamber to its normal depth by injecting BSS through the incision into the anterior chamber.
   1. Add inhibitors or activators of signaling pathways of interest to the BSS to test their roles in the lens injury response^12,15.
      NOTE: Animals can also be treated systemically with pathway inhibitors, which may be needed depending on the pharmacokinetics of drug turnover as the aqueous humor balance is re-established after surgery¹⁵.
2. Suture the corneal incision with square knots using 10-0 nylon thread with a tapered, curved 5.3 mm 1/2c needle (Figure 1F and Figure 4D-F).
   1. Place either one square knot stitch in the central part of the incision or two square knot stitches at both the upper and lower halves of the original incision (see Figure 3). Number of stitches depends on the length of the corneal incision.
   2. Do not puncture both corneal flaps at one time. Instead, hold one corneal flap using forceps in the non-dominant hand and push the needle/nylon through. Once one side of the cornea has been punctured, push the needle/nylon through the next corneal flap. Close the corneal flaps using square knot stitches.

8. Post-surgical care

NOTE: Topical antibiotic ointment is applied to the eye, and analgesic agents are administered. Mice do not usually show signs of distress after initial surgery. If needed, slow-release version of buprenorphine could be used.

1. Immediately following this surgery, do the following sub-steps.
   1. Apply one drop of 0.5% erythromycin ophthalmic ointment directly above the suture. Make sure the entire incision is covered in ointment.
   2. Administer an intraperitoneal injection of buprenorphine (0.1 mg/kg).
2. Allow the mouse to recover from anesthesia in a cage placed on a heating pad to optimize thermal support. 
   NOTE: Monitor the animal for signs of trauma, distress, or discomfort. Some signs include infection/scratching at the surgical site, displaced suture, loss of fur, etc.

9. Monitoring animal recovery

1. Monitor the animal's recovery after surgery until it awakes from anesthesia.
2. Check mice every day for the first 3 days following surgery.
   NOTE: It is usually not needed to remove stitches because mice are either sacrificed prior to full healing (2 weeks PCS) or stitches spontaneously fall out.
3. Administer buprenorphine orally, as needed for pain management after 4-8 h post-surgery, at a dosage of 0.5 mg/kg (in Jello) or a dosage of 0.1 mg/kg for those not eating. Alternatively, administer buprenorphine extended-release just prior to surgery.
   NOTE: When possible, preemptive analgesics should be utilized, yet this may require adjustments to the analgesic protocol that require review and approval from the institutional animal care and use committee.

10. Euthanasia

1. Euthanize the mouse at a desired time post-surgery
   1. Removed the operated eye using the unoperated eye as control tissue; or re-anesthetize the animal, perform lens fiber cell removal surgery on the previously unoperated contralateral eye, then immediately sacrifice the animal (without allowing it to awake from anesthesia) to provide a time zero operated control.
   2. Ensure quick dissection and tissue harvest.
      NOTE: LECs undergo transcriptomic changes within minutes of surgery so rapid tissue harvest and fixation/freezing are necessary when studying early LEC responses to surgery.

11. Performing further analysis on harvested lens tissues

1. Use the obtained tissue for immunostaining, RNA sequencing, cell culture, or western analysis^{12,15,23,25,26,28}. Flow cytometry is theoretically possible as well¹², although the small number of LECs in the mouse lens (~40,000)²⁹ and the presence of capsule associated cell debris following surgery make this challenging.
2. Immunostaining
   1. Euthanize the mouse in a chamber via CO₂ inhalation (4 L/min) from a gas cylinder. Increase the flow rate to 8-10 L/min once the mouse is unconscious and continue this exposure until complete cessation of breathing plus 1 min.
      NOTE: The mouse can be euthanized using an appropriate method per AVMA Guidelines for the Euthanasia of Animals or per Institutional Animal Care and Use Committee approved guidelines or policies.
   2. Ensure proper sacrifice by employing a secondary method of euthanasia, e.g., cervical dislocation.
      NOTE: The Duncan lab has investigated time points ranging from minutes to months PCS^12,15,23.
   3. After the mouse has been humanely sacrificed, isolate the operated eye for analysis.
      1. Hold micro forceps in the non-dominant hand to grasp the operated eye while holding micro scissors in the dominant hand.
      2. Gently clip the optic nerve and muscles anchoring the eye.
         NOTE: Be particularly careful handling eyes that were recently operated on versus those that have had longer to heal PCS. Recently operated eyes are more structurally fragile.
   4. Embed the dissected eye in OCT media for later cryo-sectioning, immunostaining, and semi-quantitative confocal microscopy^28,30.
3. Isolation of the capsular bag with associated cells for RNA sequencing, cell culture, or western blot analysis
   1. Euthanize the mouse in a chamber via CO₂ inhalation (4 L/min) from a gas cylinder. Increase the flow rate to 8-10 L/min once the mouse is unconscious and continue this exposure until complete cessation of breathing plus 1 min.
   2. Ensure proper sacrifice by employing a secondary method of euthanasia, e.g., cervical dislocation.
   3. If an early timepoint PCS (within 1-2 days) is required, either remove stitches prior to tissue extraction or make another incision in the cornea through which to extract the lens capsule.
      NOTE: At later time points PCS, the stitches will likely have fallen out, so a new incision will be needed.
4. Locate the translucent lens capsule.
   1. Gently insert the micro forceps through the corneal opening, grasp the capsule, and remove it from the anterior chamber.
   2. Place the capsule in a 1.5 mL microcentrifuge tube or culture dish for further analysis.

Results

Based on the results from this surgical method, the Duncan Lab has used this mouse model of cataract surgery to construct an injury response time course of lens epithelial cells post-surgery (Figure 5). By 6 h after surgery, 5% of the lens epithelial transcriptome is differentially expressed (Figure 6A), including the upregulation of numerous immediate early response genes and proinflammatory cytokines²³ (Figure 6B

Discussion

This surgical technique requires advanced mouse handling and micro-surgical skills that take practice to develop. The most difficult to master is the placement of fine sutures in the cornea to close the wound. Suppose the experimenter is a total novice to suturing. In that case, it is recommended to first practice using string and cloth, then move to practice using large-diameter sutures on small fruits to master the needed hand movements to make surgical square knots. Then, the experimenter must practice making incision...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.5% Erythryomycin opthalmic ointment USP	Baush Lomb	24208-910-55
1% Atropine sulfate ophthalmic solution	Amneal	60219-1748-2
1% Tropicamide opthalmic solution USP	Akorn	17478-102-12
10-0 Nylon suture	Ethicon	7707G
2.5% Phenylephrine hydrochloride ophtahlmic solution USP	Akorn	17478-200-12
26 G 1/2 Needles - straight	BD PrecisionGlide	5111
27 G 45° bent dispensing tip 1"	Harfington
Balanced saline solution	Phoenix	57319-555-06
Bupranorphine (0.1 mg/kg)	APP Pharmaceticals	401730D
Chlorhexidine solution
Needle holder 2-110	Duckworth & Kent	2-110-3E
Noyes scissors, straight	Fine Science Tools	12060-02
SMZ800 Nikon model microscope	Nikon
Sterile disposable scalpel No.11	Feather	2975#11
Tweezers #5 Dumont	Electron Microscopy Sciences	72700-D
Xylazine/Ketamine/Acepromazine (35 mg/kg, 80 mg/kg, 0.5 mg/kg) solution	APP Pharmaceticals	401730D