Optimizing Mouse Primary Lens Epithelial Cell Culture: A Comprehensive Guide to Trypsinization

Summary

This manuscript outlines a detailed video protocol for culturing primary lens epithelial cells (LECs), aiming to improve reproducibility and aid research in cataracts and posterior capsule opacification (PCO). It offers step-by-step instructions on lens dissection, LECs isolation, and validation, serving as a valuable guide, especially for newcomers in the field.

Abstract

Lens epithelial cells (LECs) play multiple important roles in maintaining the homeostasis and normal function of the lens. LECs determine lens growth, development, size, and transparency. Conversely, dysfunctional LECs can lead to cataract formation and posterior capsule opacification (PCO). Consequently, establishing a robust primary LEC culture system is important to researchers engaged in lens development, biochemistry, cataract therapeutics, and PCO prevention. However, cultivating primary LECs has long presented challenges due to their limited availability, slow proliferation rate, and delicate nature.

This study addresses these hurdles by presenting a comprehensive protocol for primary LEC culture. The protocol encompasses essential steps such as the formulation of an optimized culture medium, precise isolation of lens capsules, trypsinization techniques, subculture procedures, harvest protocols, and guidelines for storage and shipment. Throughout the culture process, cell morphology was monitored using phase-contrast microscopy.

To confirm the authenticity of the cultured LECs, immunofluorescence assays were conducted to detect the presence and subcellular distribution of critical lens proteins, namely αA- and γ-crystallins. This detailed protocol equips researchers with a valuable resource for cultivating and characterizing primary LECs, enabling advancements in our comprehension of lens biology and the development of therapeutic strategies for lens-related disorders.

Introduction

The lens of the eye plays a crucial role in vision by focusing incoming light onto the retina. It consists of a transparent, avascular structure composed of specialized cells, among which lens epithelial cells (LECs) are key players. LECs are located at the anterior surface of the lens and are responsible for maintaining its transparency, regulating water balance, and participating in lens growth and development^1,2. LECs are a unique type of cells located at the anterior part of the lens, playing a critical role in maintaining lens clarity and function by continuously producing lens fibers throughout life.

Cataracts are characterized by the progressive clouding of the lens, resulting in the distortion and scattering of light, leading to compromised vision^3,4. The precise mechanisms underlying cataract formation are complex and multifactorial, involving various cellular and molecular processes such as UV radiation, oxidative damage, and glycation^5,6. LECs have been found to contribute significantly to the development of cataracts, making them a vital focus of research^1,2,7,8,9.

Furthermore, one of the most pressing issues in ophthalmology today is the relatively high incidence of posterior capsule opacification (PCO), also known as secondary cataract. PCO remains the most common complication after cataract surgery, affecting up to 20-40% of adult patients and 100% of children within 5 years post surgery¹⁰. PCO is primarily caused by the residual LECs that remain in the capsular bag following cataract extraction. These cells undergo a multifaceted pathophysiological transformation involving not only epithelial-to-mesenchymal transition (EMT) but also the differentiation of LECs to lens fibers, resulting in a cell population that is a mixture of LECs, fibers, and myofibroblasts^11,12,13. The transformed cells proliferate and migrate across the posterior lens capsule, leading to visual impairment. Understanding the behavior and control mechanisms of LECs in culture models can provide valuable insights into the prevention and management of PCO. Therefore, this protocol of culturing LECs presents a vital tool for ophthalmic researchers aiming to study, understand, and ultimately combat this prevalent postoperative complication.

To unravel the intricacies of LEC biology and its role in cataract formation and PCO, it is essential to establish robust and reproducible in vitro primary cell culture systems. Primary LEC culture provides researchers with a controlled environment to study the functions, signaling, and molecular characteristics of LECs. Furthermore, it allows for the investigation of cellular processes and the effects of different experimental conditions, providing valuable insights into lens physiology and pathology.

Prior research has enriched our understanding of LEC culture techniques^{14,15,16,17,18,19,20}. Although these studies have employed various methodologies and yielded significant findings on LEC behavior and characteristics, a comprehensive and accessible video recording protocol for culturing LECs is absent in the current literature. This limitation can hinder novice researchers' ability to accurately reproduce the techniques and can lead to inconsistencies and variations in experimental results. By providing a video recording protocol, this research paper aims to bridge this gap and provide a standardized resource that can enhance reproducibility and facilitate knowledge transfer in the field of LEC culture.

Protocol

All animal experiments were performed in accordance with the Association for Research in Vision and Ophthalmology guidelines for the Use of Animals in Ophthalmic and Vision Research. Procedural approval was granted by the University of North Texas Health Science Center Animal Care and Use Committee (protocol number: IACUC-2022-0008). Young C57BL/6J mice, typically under 2 weeks of age, were used in these studies.

1. Culture medium preparation and lens dissection

1. Prepare culture medium by adding 50 mL of fetal bovine serum (FBS) and 0.1 mL of 50 mg/mL gentamicin to 450 mL of DMEM.
2. Humanely euthanize C57BL/6J mice younger than 2 weeks.
   NOTE: We euthanized using the CO₂ inhalation method. An optimal flow rate for CO₂ euthanasia systems should displace 30% to 70% of the chamber or cage volume/min.
3. Gently remove the eyelids using surgical scissors and apply delicate pressure with curved tweezers on opposite sides of the eye socket, causing the eye to protrude outward. Make a careful incision on the cornea using the cataract knife and carefully extract the lens using curved tweezers, ensuring no damage to the lens or its capsule.
   NOTE: Exercise caution while performing these steps to maintain the integrity of the lens capsule. Due to the delicate nature of the lens, it is important to use dissecting tools with curved and blunt tips to minimize the risk of lens damage.
4. Utilize curved tweezers with blunt tips to transfer the lenses into a 60 mm plastic tissue culture dish filled with 5 mL of prewarmed and sterile Dulbecco's phosphate buffered saline (DPBS) solution containing 10 µg/mL gentamicin.
5. Gently rinse the lenses with DPBS solution containing 10 µg/mL gentamicin to remove any potential debris or contaminants, prepare the lenses for further processing, and maintain a sterile culture environment.
6. To obtain an adequate number of LECs, pool four lenses for a 24-well culture plate and six lenses for a 6-well culture plate.

2. LECs isolation

1. After completing the rinsing process, place the lens on a piece of filter paper, allowing it to dry.
2. Once the lens is adequately dry, transfer it carefully to the cover of a Petri dish in preparation for the lens capsule removal.
3. Rotate the lens upward, ensuring that the anterior segment is facing upward. While using the tweezers to hold the anterior capsule, employ the capsulorhexis forceps in the dominant hand to create a small tear in the capsule. Gently pull the two tools in opposite directions to remove the capsule and put it in DPBS until all the lens dissections are completed.
   NOTE: To avoid any discrepancies, researchers should promptly dissect each lens epithelial capsule and temporarily store them in DPBS. Only after completing all dissections are the capsules collectively transferred to trypsin maintained at 37 °C, ensuring synchronized and uniform exposure.
4. Carefully transfer the lens capsule to a 6-well plate. Add 1 mL of 0.05% trypsin solution to each well to initiate the enzymatic digestion process.
5. Gently agitate the trypsin solution to ensure even permeation. Place the plate in a cell culture incubator and allow the capsule to be digested for 8-10 min at 37 °C.
   NOTE: This step facilitates the breakdown of the lens capsule tissue and the subsequent release of individual epithelial cells.
6. After the incubation, carefully mince the digested lens capsule using dissecting scissors to break down any remaining tissue clumps and promote cell separation.
   NOTE: Thoroughness in tissue mincing is emphasized to ensure efficient cell release from the digested lens capsules.
7. Add 0.5 mL of the culture medium containing 10% FBS to quench the trypsin. Transfer the tissue samples to a centrifugation tube and centrifuge at 1,000 × g for 5 min.
8. Carefully remove the supernatant without disturbing the cell pellet. Use 1 mL of culture medium to resuspend the cells and seed the cells in a 24-well plate.
9. Change the culture medium every 2-3 days.

3. LECs subculture

1. Once the cells achieve confluence, remove the medium from the culture dish. Proceed to wash the cells 2x with 1 mL of DPBS.
2. Add 200 µL of trypsin-EDTA solution and place the cells in the incubator for 5 min.
3. After incubation, remove the cells from the incubator and inspect them under a microscope to confirm that they have detached from the culture dish and begun to float.
4. Add 1 mL of culture medium and gently pipette the cells 3-5x to detach all the cells.
5. Transfer the cells to centrifuge tubes and centrifuge at 1,000 × g for 5 min.
6. Carefully remove the supernatant and resuspend the cells in the complete growth medium.
7. If needed, count the cell number using a hemocytometer.
8. Subdivide the cell suspension at a 1:2 or 1:3 ratio for subculturing purposes.
9. When the culture becomes confluent again, repeat the aforementioned procedures.
   ​NOTE: LECs flourish in high-density culture conditions. Avoid excessively diluting the cells, as this may hinder their growth.

4. Storage and shipment

NOTE: The ideal cell number for storage is ~1 × 10⁶.

1. Thoroughly wash the cells 3x with 1 mL of DPBS. After washing, add 1 mL of trypsin-EDTA solution and place the cells in the incubator for 5 min.
2. Add 2 mL of complete culture medium and transfer the cell suspension to the centrifuge tube and centrifuge at 1,000 × g for 5 min.
3. Discard the supernatant and resuspend the cells in a freezing medium composed of 90% FBS and 10% DMSO, aiming for a cell density of 1 × 10⁶ cells/mL. Transfer the cell suspension to the cryovial.
4. Immediately move the cells to a -20 °C environment for 1 h, followed by -80 °C overnight, prior to permanent storage in liquid nitrogen.
   NOTE: If liquid nitrogen is unavailable, the cells can be stored at -80 °C after an initial hour at -20 °C.
5. If a shipment is required, ship the cells in the cryovial in a package with dry ice for overnight delivery.
6. Upon receipt of the cells, ensure swift recovery and place the cells in the subculture. If immediate culture is not feasible, transfer the cells to liquid nitrogen for prolonged storage.
   ​NOTE: If the cells are to be shipped, ensure the samples are deeply buried in the dry ice to prevent potential damage from temperature fluctuations.

5. LECs validation

1. Plate LECs into 35 mm culture dishes with cover glasses and culture them for approximately 48 h.
2. Wash the cells 2x with PBS and fix the cells with cold methanol for 10 min at -20 °C.
3. Wash the fixed cells for 3 x 5 min with PBS and incubate the fixed cells with blocking buffer for 1 h at room temperature to prevent non-specific binding.
4. After blocking, incubate the cells overnight at 4 °C with the primary antibodies (αA-crystallin, γ-crystallin, and PROX1 antibodies) individually diluted at a 1:50 ratio in diluent buffer.
5. Wash the cells for 3 x 5 min with PBS and incubate the cells with the secondary antibody diluted 1:100 in diluent buffer for 1 h.
6. Wash the cells for 3 x 5 min with PBS and stain the cells with 5 µg/mL Hoechst 33342 in PBS for 10 min at room temperature to visualize the nuclei.
7. Wash the cells for 2 x 5 min with PBS to remove excess staining solution and capture fluorescent images of the cells using a fluorescence microscope using the DAPI channel for nuclei and FITC channel for αA-, γ-crystallins, and PROX1.

Representative Results

As shown in Figure 2, by following this protocol, primary LECs from C57BL/6J mice adhered to the dishes within a period of 4 h. Notably, there were visible remnants of other tissues such as sections of the posterior capsule and lens fiber cells. However, these unintended elements did not attach to the dish and could, therefore, be removed by changing the culture medium. Subsequently, between the third and fifth day, the LECs initiated their proliferation phase. Rapid growth, characteristic of the logarithmic proliferation phase, could then be observed between the seventh and tenth day. In instances when the cells do not progress to the rapid growth phase, it may be advisable to incorporate growth supplements, such as EpiCGS-a, into the culture medium. Furthermore, we observed that cells exhibit a faster growth rate in the medium containing 20% FBS compared to 10% FBS. A lower FBS concentration typically fosters slower growth, mirroring the traits of the lens's central epithelium, while a 20% FBS concentration replicates the attributes of the lens's proliferative zone.

According to Reddy et al. and Andley et al., who developed the immortalized human lens epithelial cell line B3, it is expected that LECs should express various lens-specific proteins such as αA- and γ-crystallins^2,21. Hence, in this study, we employed αA- and γ-crystallins as cellular markers to validate the identity of the cells as LECs. LECs within the initial three passages were cultured on glass coverslips. The primary antibodies specific to αA- and γ-crystallins were used to incubate the cells overnight. As shown in Figure 3, these cells exhibited robust expression of both αA- and γ-crystallins, providing definitive evidence that they are lens-derived epithelial cells. Additionally, we used PROX1 as a well-established marker for fiber cells to label the primary LECs. The data indicated that these LECs showed negative staining for PROX1, confirming that these cells are epithelial cells and had not undergone differentiation into fiber cells yet.

Figure 1: Step-by-step workflow for LEC culture. The figure depicts the sequential steps involved in the culture of primary lens epithelial cells. Step 1 involves creating a small tear in the anterior capsule. Step 2 entails gently removing the lens capsule. In Step 3, the lens capsule is carefully transferred to a 24-well plate and incubated with 0.05% trypsin solution at 37 °C for 10 min. Following incubation, the digested lens capsule is minced using dissecting scissors to promote cell separation. Step 4 involves quenching the trypsin by adding 0.5 mL of culture medium containing 20% FBS to the centrifugation tube and centrifuging the tissue samples at 1,000 × g for 5 min. Step 5 requires resuspending the cells in 1 mL of culture medium and seeding them in a 24-well plate. Finally, Step 6 involves changing the culture media every 2-3 days to support cell growth and maintenance throughout the experiment. Abbreviations: LECs = lens epithelial cells; FBS = fetal bovine serum. Please click here to view a larger version of this figure.

Figure 2: Growth pattern of LECs over time. The morphological changes of primary lens epithelial cells at various time points during their growth were recorded under a phase-contrast microscope. The images captured on days 1, 2, 3, 5, 7, and 10 depict the evolving cell morphology, providing insight into the development and behavior of lens epithelial cells over time. Red arrow indicating the location of actively proliferating lens epithelial cells. Scale bars = 100 µm. Abbreviation: LECs = lens epithelial cells. Please click here to view a larger version of this figure.

Figure 3: Validation of LECs. (A) Immunostaining of αA-crystallin in mouse LECs. Primary mLECs were stained with Hoechst 33342 (blue) and αA-crystallin antibody (green). (B) Immunostaining of γ-crystallin in mouse LECs. (C) Immunostaining of PROX1 in mouse LECs. Primary mLECs were stained with Hoechst 33342 (blue) and PROX1 antibody (green). Scale bars = 50 µm. Abbreviation: LECs = lens epithelial cells. Please click here to view a larger version of this figure.

Discussion

The protocol presented in this paper provides a comprehensive, step-by-step guide to the successful isolation, culture, and subculture of primary LECs, complete with accompanying video documentation. The detailed visual guide alongside the written instructions enhances the clarity and accessibility of the protocol, promoting its use and reproducibility among researchers in the field. The ultimate aim is to contribute to the expanding body of knowledge surrounding the role of LECs in cataract formation and PCO, a prevalent complication following cataract surgery.

When comparing primary LECs with lens epithelial cell lines, such as HLE-B3 and SRA01/04, each presents unique advantages and challenges in a research context. The HLE-B3 cell line, along with the SRA01/04, represent a category of cell lines that, while being easy to handle and durable, often exhibit genetic and phenotypic changes due to their continuous replication and prolonged culture conditions. This can lead to significant differences from the function of the original LECs, thereby reducing the authenticity of their responses. Conversely, primary LECs, directly isolated from living tissue from patients or research animals, more accurately mirror the natural cellular environment and inherent responses of the lens in vivo. Despite the added complexity of their extraction and culture, they are often preferred in studies demanding high physiological relevance, as they deliver more accurate and reliable results.

Some key points must be considered while following this protocol. Young C57BL/6J mice, typically under 2 weeks of age, were used in these studies. We observed that LECs harvested from these young mice demonstrated more vigorous growth compared to LECs from mice aged 2 months or older. This indicates a negative correlation between the age of the mice and cell growth.

The dissection of the lens from a mouse eye is a delicate task, necessitating the careful use of dissecting tools to preserve the integrity of the lens and its capsule. The procedure outlined in protocol section 1 ensures that the lens is successfully extracted with minimal damage. While maintaining the health and integrity of the lens capsule is challenging, its importance cannot be understated as any damage could potentially impact the quality and quantity of LECs isolated. It is noteworthy that the majority of cell divisions typically occur in the germinative zone near the equatorial region in the lens, an area not easily accessible for culturing. According to Zetterberg et al., although the central part of the lens epithelium showcases minimal mitotic activity under normal circumstances, experiments utilizing tritiated thymidine (³H-Tdr) labeling have marked these centrally positioned lens epithelium cells as potential stem cells^17,22. Stem cells exhibit distinct characteristics, such as limitless proliferation capacity despite having a low proliferation rate under standard conditions. As such, the central part of the lens epithelium may provide a better representation of the proliferative capacity of the LECs than the germinative zone.

The isolation of LECs, as detailed in protocol section 2, constitutes a pivotal step in this experimental procedure. Integral actions including the removal of the lens capsule, enzymatic digestion, and tissue fragmentation are carefully performed to separate individual epithelial cells from the capsule. One of the critical elements within this process is the duration of trypsin digestion. It is recommended that the digestion period be set between 8 and 10 min. If this period is shortened to less than 5 min, it may result in incomplete cell separation. Conversely, overextension of this time frame may significantly compromise cell viability. The strategic choice of employing trypsin-EDTA as a cell dissociation reagent, in conjunction with a DMEM culture medium supplemented with 20% FBS and 10 µg/mL gentamicin, optimizes cell release and subsequent proliferation whilst mitigating potential contamination risks.

Antibiotics are frequently used to prevent bacterial contamination during primary LEC cultures. However, the choice of antibiotics should be made carefully. Commonly utilized antibiotics such as penicillin and streptomycin, as well as antifungal agents, have the potential to impact the viability of LECs. As an alternative, it is recommended to employ a gentamicin solution at a concentration of 10 µg/mL for optimal LEC growth.

Furthermore, maintaining a high cell density is another crucial factor for successful primary LEC culture. Unlike established cell lines, primary LECs need a higher cell density for optimal growth. This is primarily due to their reliance on effective intercellular communication, facilitated by direct contact or paracrine signaling, which helps maintain their differentiation status and function. High cell density also mitigates the adverse effects of "culture shock," a condition experienced by primary cells when isolated and placed into an in vitro environment significantly different from their in vivo origin²³. By mimicking a more in vivo-like environment, a high cell density increases survival rates. Additionally, this density helps establish a concentration gradient of growth factors and cytokines that supports cell growth and function. Given that many primary cells are anchorage-dependent, requiring surface attachment for proliferation, a high cell density offers an adequate number of neighboring cells for adhesion, thus promoting healthy growth. Consequently, the regulation of cell density is a crucial consideration in the cultivation of primary cells due to its significant impact on cell communication, survival, and proliferation. Opting for smaller culture dishes, such as 24-well or 6-well plates, is recommended to create an ideal environment for LECs. Following this protocol typically results in LECs reaching a confluent state ~10-14 days post cultivation. We recommend utilizing LECs at a low number of passages, ideally between P0 and P6, to ensure the most natural behavior in experiments. Beyond 7-10 passages, LECs may exhibit diminished growth and may not react to experimental conditions in the same manner as cells in lower passages.

The serum concentration is directly related to the rate of cell growth. Lower levels of FBS are more likely to induce slower growth, resembling the characteristics of the central epithelium. In contrast, higher FBS levels mimic the conditions of the proliferative zone. If cell growth is slow, as may occur when LECs need to be isolated from older or genetically modified animals, it may be beneficial to increase FBS to 20% or add a growth supplement like EpiCGS-a (5 mL, see Table of Materials) to the culture medium. This serum and supplement enrichment can enhance cell proliferation and promote the optimal growth of epithelial cells.

The storage and shipment protocol (protocol section 5) considers the challenges associated with the preservation and transport of live cells. The choice of freezing medium is critical for the survival of LECs during storage and transportation. We have experimented with different freezing mediums, including 70% complete culture medium + 20% FBS + 10% DMSO; 90% FBS + 10% DMSO; and DMSO- and serum-free freezing medium. Evidence from our experiments suggests that a solution comprising 10% DMSO and 90% FBS exhibits superior performance in preserving cell viability. Utilizing this specific formulation, we have successfully maintained primary LECs in storage at -80 °C for durations exceeding 10 years, demonstrating the robustness of this approach and its ability to facilitate cell revival post storage.

αA-crystallin and γ-crystallin were used as markers for LECs. PROX1 was utilized as a marker for lens fiber cells. Additional markers such as PAX6, FOXE3, and E-cadherin may also be employed to characterize the LEC phenotype. If researchers are interested in exploring EMT, αSMA should be used as a marker. It is essential to adjust incubation times and dilutions in accordance with the specific requirements of the experiment and the recommendations provided for the respective antibodies utilized.

While this study presents a robust methodology for culturing primary LECs, it is important to acknowledge its limitations. While the cells isolated initially are primary LECs, any subculturing, trypsinization, and reanimation procedures will lead to alterations in their status. The protocol we have designed primarily employs mouse lens as the source of LECs; however, LECs can also be cultured from other resources, including cataract patient samples, eye bank eyes, or patients with different ocular diseases including glaucoma and diabetic retinopathy^17,24. LECs harvested from older individuals or those with specific ocular conditions might not comply with the protocol as effectively as cells from younger, healthier counterparts. Culturing LECs from older or genetically modified individuals or animals might require optimizing the culture medium, potentially by increasing FBS to 20% or incorporating additional growth factors. Additionally, previous studies by Menko et al., conducted on primary embryonic chick lens epithelial cell cultures, demonstrated spontaneous differentiation occurring after the second day of culture²⁵. Therefore, researchers should exercise caution and consider the differences in methodologies, especially if studying differentiation is their main research goal.

Various methods can be utilized to culture primary LECs. For instance, methods developed by Ibaraki et al., Sundelin et al., and Andjelic et al., involve culturing primary LECs directly on the Petri dish using the anterior portion of the lens capsule collected during cataract surgery^16,17,19,20. This approach maintains natural cell-to-cell contacts and the extracellular matrix, providing high physiological relevance crucial for conditions like PCO. Alternatively, the lens explant method, such as outlined by Zelenka et al. preserves native tissue architecture, allowing for studies that are more physiologically relevant, especially beneficial for exploring the terminal differentiation of LECs, cellular interactions, and lens development processes, granting a detailed understanding of sequential cellular and molecular events during differentiation²⁶.

In contrast, the trypsinization method described in this protocol produces a uniform single-cell suspension, simplifying uniform seeding and precise cell counting, which is advantageous for certain experiments such as cell viability and proliferation assays, drug screening, and cell signaling pathway analysis. However, it is critical to acknowledge that while this method facilitates controlled and precise studies due to its uniform cell population, it may alter cellular behavior and compromise the physiological relevance seen in the explant and direct-culture methods due to enzymatic processing. For researchers focused exclusively on studying epithelial cells, this method proves highly applicable and convenient, offering dependable insights into the characteristics and behaviors of these cells. However, for those whose scientific inquiries extend to cellular differentiation, it becomes essential to contemplate alternative methods such as explant techniques, or adapt and optimize the current one to encompass these aspects.

Overall, this protocol is designed with careful consideration of the specific needs and requirements of LECs. Every step, from dissection to validation, is thoughtfully crafted to maintain the viability and functionality of the cells. As a result, it can be a valuable guide for researchers studying LECs and their role in ocular physiology and pathology. Future studies can adapt and modify this protocol to explore different aspects of LEC biology, providing a platform for further advancements in the understanding of lens-related diseases and the development of new therapeutic strategies for cataract and PCO prevention.

Materials

Name	Company	Catalog Number	Comments
0.05% Trypsin-EDTA	Thermo Fisher	#25300054	For LECs dissociation
Alexa Fluor 488 Secondary Antibody	Jackson ImmunoResearch	#715-545-150	For cell validation
Alexa Fluor 647 AffiniPure Goat Anti-Rabbit IgG (H+L)	Jackson ImmunoResearch	111-605-003	For cell validation
Antibody dilution buffer	Licor	#927-60001	For cell validation
Beaver safety knife	Beaver-Visitec International	#3782235	For lens dissection
Blocking buffer	Licor	#927-60001	For cell validation
Capsulorhexis forceps	Titan Medical Instruments	TMF-124	For lens capsule isolation
DMEM	Sigma Aldrich	D6429	For LECs culture medium
DMSO	Sigma Aldrich	#D2650	For making freezing medium
Dulbecco's Phosphate Buffered Saline	Thermo Fisher	#J67802	For lens dissection
Dumont tweezers	Roboz Surgical Instrument	RS-4976	For lens capsule isolation
EpiCGS-a (optional)	ScienCell	4182	For LECs culture medium
FBS	Sigma Aldrich	F2442	For LECs culture medium
Gentamicin (50 mg/mL)	Sigma-Aldrich	G1397	For LECs culture medium
Hoechst 33342 solution	Thermo Fisher	#62249	For cell validation
Micro-dissecting scissors	Roboz Surgical Instrument	RS-5983	For lens dissection
Micro-dissecting tweezers	Roboz Surgical Instrument	RS5137	For lens dissection
PROX1 antibody	Thermo Fisher	11067-2-AP	For cell validation
Vannas micro-dissecting spring scissors	Roboz Surgical Instrument	RS-5608	For lens capsule isolation
αA-crystallin antibody	Santa Cruz	sc-28306	For cell validation
γ-crystallin antibody	Santa Cruz	sc-365256	For cell validation