Name: optimizing mouse primary lens epithelial cell culture: a comprehensive guide to trypsinization
Uploaded: 2024-06-21T12:30:00.000+00:00
Duration: 3 min 47 s
Description: Lens epithelial cells (LECs) play multiple important roles in maintaining the homeostasis and normal function of the lens. LECs determine lens growth, ...

This work was supported by NEI R21EY033941 (to Hongli Wu); Department of Defense W81XWH2010896 (to Hongli Wu); R15GM123463-02 (to Kayla Green and Hongli Wu)

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
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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&#39;s phosphate buffered saline (DPBS) solution containing 10 &#181;g/mL gentamicin.</li>
	<li>Gently rinse the lenses with DPBS solution containing 10 &#181;g/mL gentamicin to remove any potential debris or contaminants, prepare the lenses for further processing, and maintain a sterile culture environment.</li>
	<li>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.</li>
</ol>
2. LECs isolation
<ol>
	<li>After completing the rinsing process, place the lens on a piece of filter paper, allowing it to dry.</li>
	<li>Once the lens is adequately dry, transfer it carefully to the cover of a Petri dish in preparation for the lens capsule removal.</li>
	<li>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 &#176;C, ensuring synchronized and uniform exposure.</li>
	<li>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.</li>
	<li>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 &#176;C. 
	NOTE: This step facilitates the breakdown of the lens capsule tissue and the subsequent release of individual epithelial cells.</li>
	<li>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.</li>
	<li>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 &#215; g for 5 min.</li>
	<li>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.</li>
	<li>Change the culture medium every 2-3 days.</li>
</ol>
3. LECs subculture
<ol>
	<li>Once the cells achieve confluence, remove the medium from the culture dish. Proceed to wash the cells 2x with 1 mL of DPBS.</li>
	<li>Add 200 &#181;L of trypsin-EDTA solution and place the cells in the incubator for 5 min.</li>
	<li>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.</li>
	<li>Add 1 mL of culture medium and gently pipette the cells 3-5x to detach all the cells.</li>
	<li>Transfer the cells to centrifuge tubes and centrifuge at 1,000 &#215; g for 5 min.</li>
	<li>Carefully remove the supernatant and resuspend the cells in the complete growth medium.</li>
	<li>If needed, count the cell number using a hemocytometer.</li>
	<li>Subdivide the cell suspension at a 1:2 or 1:3 ratio for subculturing purposes.</li>
	<li>When the culture becomes confluent again, repeat the aforementioned procedures. 
	&#8203;NOTE: LECs flourish in high-density culture conditions. Avoid excessively diluting the cells, as this may hinder their growth.</li>
</ol>
4. Storage and shipment
NOTE: The ideal cell number for storage is ~1 &#215; 106.
<ol>
	<li>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.</li>
	<li>Add 2 mL of complete culture medium and transfer the cell suspension to the centrifuge tube and centrifuge at 1,000 &#215; g for 5 min.</li>
	<li>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 &#215; 106 cells/mL. Transfer the cell suspension to the cryovial.</li>
	<li>Immediately move the cells to a -20 &#176;C environment for 1 h, followed by -80 &#176;C overnight, prior to permanent storage in liquid nitrogen. 
	NOTE: If liquid nitrogen is unavailable, the cells can be stored at -80 &#176;C after an initial hour at -20 &#176;C.</li>
	<li>If a shipment is required, ship the cells in the cryovial in a package with dry ice for overnight delivery.</li>
	<li>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. 
	&#8203;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.</li>
</ol>
5. LECs validation
<ol>
	<li>Plate LECs into 35 mm culture dishes with cover glasses and culture them for approximately 48 h.</li>
	<li>Wash the cells 2x with PBS and fix the cells with cold methanol for 10 min at -20 &#176;C.</li>
	<li>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.</li>
	<li>After blocking, incubate the cells overnight at 4 &#176;C with the primary antibodies (&#945;A-crystallin, &#947;-crystallin, and PROX1 antibodies) individually diluted at a 1:50 ratio in diluent buffer.</li>
	<li>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.</li>
	<li>Wash the cells for 3 x 5 min with PBS and stain the cells with 5 &#181;g/mL Hoechst 33342 in PBS for 10 min at room temperature to visualize the nuclei.</li>
	<li>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 &#945;A-, &#947;-crystallins, and PROX1.</li>
</ol>

Cultivation and Characterization of Lens Epithelial Cells

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The authors declare that they have no conflicts of interest.

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 prevalen...

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 development1,2. 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 vision3,4. The precise mechanisms underlying cataract formation are complex and multifactorial, involving various cellular and molecular processes such as UV radiation, oxidative damage, and glycation5,6. LECs have been found to contribute significantly to the development of cataracts, making them a vital focus of research1,2,7,8,9.
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 myofibroblasts11,12,13. 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 techniques14,15,16,17,18,19,20. 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&#39; 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.

<table><tbody><tr><td>0.05% Trypsin-EDTA</td><td>Thermo Fisher</td><td>#25300054</td><td>For LECs dissociation</td></tr><tr><td>Alexa Fluor 488 Secondary Antibody&amp;nbsp;</td><td>Jackson ImmunoResearch</td><td>#715-545-150</td><td>For cell validation</td></tr><tr><td>Alexa Fluor 647 AffiniPure Goat Anti-Rabbit IgG (H+L)</td><td>Jackson ImmunoResearch</td><td>111-605-003</td><td>For cell validation</td></tr><tr><td>Antibody dilution buffer</td><td>Licor</td><td>#927-60001</td><td>For cell validation</td></tr><tr><td>Beaver safety knife</td><td>Beaver-Visitec International</td><td>#3782235</td><td>For lens dissection</td></tr><tr><td>Blocking buffer</td><td>Licor</td><td>#927-60001</td><td>For cell validation</td></tr><tr><td>Capsulorhexis forceps</td><td>Titan Medical Instruments</td><td>TMF-124</td><td>For lens capsule isolation</td></tr><tr><td>DMEM</td><td>Sigma Aldrich</td><td>D6429</td><td>For LECs culture medium</td></tr><tr><td>DMSO</td><td>Sigma Aldrich</td><td>#D2650</td><td>For making freezing medium&amp;nbsp;</td></tr><tr><td>Dulbecco&#039;s Phosphate Buffered Saline&amp;nbsp;</td><td>Thermo Fisher</td><td>#J67802</td><td>For lens dissection</td></tr><tr><td>Dumont tweezers</td><td>Roboz Surgical Instrument</td><td>RS-4976</td><td>For lens capsule isolation</td></tr><tr><td>EpiCGS-a (optional)</td><td>ScienCell</td><td>4182</td><td>For LECs culture medium</td></tr><tr><td>FBS</td><td>Sigma Aldrich</td><td>F2442</td><td>For LECs culture medium</td></tr><tr><td>Gentamicin (50&amp;nbsp;mg/mL)</td><td>Sigma-Aldrich</td><td>G1397</td><td>For LECs culture medium</td></tr><tr><td>Hoechst 33342 solution</td><td>Thermo Fisher</td><td>#62249</td><td>For cell validation</td></tr><tr><td>Micro-dissecting scissors</td><td>Roboz Surgical Instrument&amp;nbsp;</td><td>RS-5983</td><td>For lens dissection</td></tr><tr><td>Micro-dissecting tweezers</td><td>Roboz Surgical Instrument&amp;nbsp;</td><td>RS5137&amp;nbsp;</td><td>For lens dissection</td></tr><tr><td>PROX1 antibody</td><td>Thermo Fisher</td><td>11067-2-AP</td><td>For cell validation</td></tr><tr><td>Vannas micro-dissecting spring scissors</td><td>Roboz Surgical Instrument</td><td>RS-5608</td><td>For lens capsule isolation</td></tr><tr><td>&amp;alpha;A-crystallin antibody</td><td>Santa Cruz</td><td>sc-28306</td><td>For cell validation&amp;nbsp;</td></tr><tr><td>&amp;gamma;-crystallin antibody</td><td>Santa Cruz</td><td>sc-365256</td><td>For cell validation</td></tr></tbody></table>

optimizing mouse primary lens epithelial cell culture: a comprehensive guide to trypsinization

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 &#945;A- and &#947;-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.

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 o...

Author Spotlight: Unlocking the Secrets of Cataracts &#8211; Investigating Redox Repair Enzymes in Lens Epithelial Cells

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.

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

Lens epithelial cells (LECs) play multiple important roles in maintaining the homeostasis and normal function of the lens. LECs determine lens growth, ...

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Research

JoVE Journal

Biology

1.7K Views.  University of North Texas Health Science Center. 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.

Video: Author Spotlight: Unlocking the Secrets of Cataracts – Investigating Redox Repair Enzymes in Lens Epithelial Cells

Generation of Mosaic Mammary Organoids by Differential Trypsinization

Organoids offer self-organizing, three-dimensional tissue structures that recapitulate physiological processes in the convenience of a dish. The murine mammary gland is composed of two distinct epithelial cell compartments, serving different functions: the outer, contractile myoepithelial compartment and the inner, secretory luminal compartment. Here, we describe a method by which the cells comprising these compartments are isolated and then combined to investigate their individual lineage contributions to mammary gland morphogenesis and differentiation. The method is simple and efficient and does not require sophisticated separation technologies such as fluorescence activated cell sorting. Instead, we harvest and enzymatically digest the tissue, seed the epithelium on adherent tissue culture dishes, and then use differential trypsinization to separate myoepithelial from luminal cells with ~90% purity. The cells are then plated in an extracellular matrix where they organize into bilayered, three-dimensional (3D) organoids that can be differentiated to produce milk after 10 days in culture. To test the effects of genetic mutations, cells can be harvested from wild type or genetically engineered mouse models, or they can be genetically manipulated prior to 3D culture. This technique can be used to generate mosaic organoids that allow investigation of gene function specifically in the luminal or myoepithelial compartment.

The mammary gland is a bilayered structure, comprising outer myoepithelial and inner luminal epithelial cells. Presented is a protocol to prepare organoids using differential trypsinization. This efficient method allows researchers to separately manipulate these two cell types to explore questions concerning their roles in mammary gland form and function.

Organoids offer self-organizing, three-dimensional tissue structures that recapitulate physiological processes in the convenience of a dish. The murine ...

Developmental Biology

Transplantation Into the Mouse Ovarian Fat Pad

Orthotopic transplantation assays in mice are invaluable for studies of cell regeneration and neoplastic transformation. Common approaches for orthotopic transplantation of ovarian surface and tubal epithelia include intraperitoneal and intrabursal administration of cells. The respective limitations of these methods include poorly defined location of injected cells and limited space volume. Furthermore, they are poorly suited for long-term structural preservation of transplanted organs. To address these challenges, we have developed an alternative approach, which is based on the introduction of cells and tissue fragments into the mouse fat pad. The mouse ovarian fat pad is located in the immediate vicinity of the ovary and uterine tube (aka oviduct, fallopian tube), and provides a familiar microenvironment for cells and tissues of these organs. In our approach fluorescence-labeled mouse and human cells, and fragments of the uterine tube are engrafted by using minimally traumatic dorsal incision surgery. Transplanted cells and their outgrowths are easily located in the ovarian fat pad for over 40 days. Long-term transplantation of the entire uterine tube allows correct preservation of all principle tissue components, and does not result in adverse side effects, such as fibrosis and inflammation. Our approach should be uniquely applicable for answering important biological questions such as differentiation, regenerative and neoplastic potential of specific cell populations. Furthermore, it should be suitable for studies of microenvironmental factors in normal development and cancer.

We describe an ovarian fad pad transplantation assay suitable for studies of normal and transformed epithelia of the female reproductive tract. The mouse fat pad allows transplantation of large tissue fragments, is easily accessible for surgery and imaging, and provides the most favorable native environment for tissues of M&#252;llerian origin.

Orthotopic transplantation assays in mice are invaluable for studies of cell regeneration and neoplastic transformation. Common approaches for orthotopic ...

Medicine

Cultivating a Three-dimensional Reconstructed Human Epidermis at a Large Scale

A three-dimensional human epidermis model reconstructed from neonatal primary keratinocytes is presented. Herein, a protocol for the cultivation process and the characterization of the model is described. Neonatal primary keratinocytes are grown submerged on permeable polycarbonate inserts and lifted to the air-liquid interface three days after seeding. After fourteen days of stimulation with defined growth factors and ascorbic acid in high calcium culture medium, the model is fully differentiated. Histological analysis revealed a completely stratified epidermis, mimicking the morphology of native human skin. To characterize the model and its barrier functions, protein levels and localization specific for early-stage keratinocyte differentiation (i.e., keratin 10), late-stage differentiation (i.e., involucrin, loricrin, and filaggrin) and tissue adhesion (i.e., desmoglein 1), were assessed by immunofluorescence. The tissue barrier integrity was further evaluated by measuring transepithelial electrical resistance. Reconstructed human epidermis was responsive to proinflammatory stimuli (i.e., lipopolysaccharide and tumor necrosis factor alpha), leading to increased cytokine release (i.e., interleukin 1 alpha and interleukin 8). This protocol represents a straightforward and reproducible in vitro method to cultivate reconstructed human epidermis as a tool to assess environmental effects and a broad range of skin-related studies.

This protocol describes a straightforward method to cultivate three-dimensional reconstituted human epidermis in a reproducible and robust manner. Additionally, it characterizes the structure-function relationship of the epidermal barrier model. The biological responses of the reconstituted human epidermis upon proinflammatory stimuli are also presented.

A three-dimensional human epidermis model reconstructed from neonatal primary keratinocytes is presented. Herein, a protocol for the cultivation process ...

Bioengineering

Characterization and Morphological Dynamics of Cultured Primary Murine RPE

A Simplified Method for Isolation and Culture of Retinal Pigment Epithelial Cells from Adult Mice

Retinal pigment epithelial cells (RPE) are critical for the proper function of the retina. RPE dysfunction is involved in the pathogenesis of important retinal diseases, such as age-related macular degeneration, retinitis pigmentosa, and diabetic retinopathy. We present a streamlined approach for the isolation of RPE from murine adult eyes. In contrast to previously reported methods, this approach enables the isolation and culture of highly pure RPE from adult mice. This simple and fast method does not require extensive technical skill and is achievable with basic scientific tools and reagents. Primary RPE are isolated from C57BL/6 background mice aged 3- to 14-weeks by enucleation of the eye followed by the removal of the anterior segment. Enzymatic trypsinization and centrifugation are used to dissociate and isolate the RPE from the eyecup. In conclusion, this approach offers a quick and effective protocol for the utilization of RPE in the study of retinal function and disease.

Author Spotlight: Efficient Isolation of Primary Retinal Pigment Epithelial Cells from Adult Mice for Retinopathy Research

Retinal pigment epithelium (RPE) acts as a crucial barrier between the choroid and retina, promoting the health and function of retinal cell types, such as photoreceptors. Herein, we describe a simple and effective protocol for isolating and culturing adult murine RPE.

Retinal pigment epithelial cells (RPE) are critical for the proper function of the retina. RPE dysfunction is involved in the pathogenesis of important ...

Neuroscience

Primary Cell Cultures from the Mouse Retinal Pigment Epithelium

The retinal pigment epithelium (RPE) is a highly polarized multi-functional epithelium that is located between the neural retina and the choroid of the eye. It is a single sheet of pigmented cells that are hexagonally packed and connected by tight junctions. The main functions of the RPE include absorption of light, phagocytosis of the shed photoreceptor outer segments, spatial buffering of ions, transport of nutrients, ions and water as well as active involvement in the visual cycle. With such important and diverse functions, it is critically important to study the biology of RPE cells. A number of RPE cell lines have been established; however, passaged and immortalized cells are known to quickly lose some of the morphological and physiological characteristics of natural RPE cells. Thus, primary cells are more suitable for studying different aspects of RPE cell biology and function. Mouse primary RPE cell culture is very useful to researchers since mouse models are widely used in biological studies, however collecting RPE cells from mouse is also very challenging due to their small size. Here, we present a protocol for establishing primary mouse RPE cell cultures which includes enucleation and dissection of the eyes and isolation of the RPE sheets to yield the cells for culturing. This method enables efficient cell recovery. The RPE cells obtained from two mice&#160;can reach confluency on one 12 mm polyester membrane insert pre-loaded in culture plate after one week of culture and display some of the original properties of bona fide RPE cells such as hexagonal shape and pigmentation after two weeks of culture.

The retinal pigment epithelium (RPE) is a multi-functional epithelium of the eye. Here we present a protocol to establish primary cell cultures derived from the murine RPE.

The retinal pigment epithelium (RPE) is a highly polarized multi-functional epithelium that is located between the neural retina and the choroid of the ...

Isolation of Mouse Epidermal Keratinocytes and Their In Vitro Clonogenic Culture

The protocol described here is a reliable method of harvesting primary keratinocytes from adult female mice (54 &#177; 2 days old) yielding approximately 30 x 106 viable cells per mouse. Primary adult mouse keratinocytes are harvested from the dorsal skin of female mice. Male mice (~6 weeks old) can be used for keratinocyte harvesting depending on the requirements of the experiment. Euthanized mice are shaved and sterilized with serial washes in povidone iodine and ethanol solutions (70% alcohol). After disinfecting the mice, the dorsal skin is removed and the subcutaneous fat and muscle are removed with a scalpel and discarded. The skins are cut into small pieces and treated with a mild, low temperature trypsinization to detach the lower dermis from the epidermis. The scraped epidermises are stirred at low speed, filtered to remove the hairs, counted, and re-suspended in culture medium. This method provides an excellent single cell suspension of highly culturable cells for many downstream applications.

The goal of this protocol is to isolate epidermal keratinocytes from the dorsal skin of adult mice for a variety of downstream applications such as molecular biology, biochemistry, fluorescence activated cell sorting, and primary in vitro uses (e.g., clonogenic keratinocytes).

The protocol described here is a reliable method of harvesting primary keratinocytes from adult female mice (54 &#177; 2 days old) yielding approximately ...

Efficient Dissection and Culture of Primary Mouse Retinal Pigment Epithelial Cells

Eye disorders affect millions of people worldwide, but the limited availability of human tissues hinders their study. Mouse models are powerful tools to understand the pathophysiology of ocular diseases because of their similarities with human anatomy and physiology. Alterations in the retinal pigment epithelium (RPE), including changes in morphology and function, are common features shared by many ocular disorders. However, successful isolation and culture of primary mouse RPE cells is very challenging. This paper is an updated audiovisual version of the protocol previously published by Fernandez-Godino et al. in 2016 to efficiently isolate and culture primary mouse RPE cells. This method is highly reproducible and results in robust cultures of highly polarized and pigmented RPE monolayers that can be maintained for several weeks on Transwells. This model opens new avenues for the study of the molecular and cellular mechanisms underlying eye diseases. Moreover, it provides a platform to test therapeutic approaches that can be used to treat important eye diseases with unmet medical needs, including inherited retinal disorders and macular degenerations.

This protocol, which was originally reported by Fernandez-Godino et al. in 20161, describes a method to efficiently isolate and culture mouse RPE cells, which form a functional and polarized RPE monolayer within one week on Transwell plates. The procedure takes approximately 3 hours.

Eye disorders affect millions of people worldwide, but the limited availability of human tissues hinders their study. Mouse models are powerful tools to ...

Primary Culture of Porcine Retinal Pigment Epithelial Cells

The retinal pigment epithelium (RPE) is a monolayer of polarized pigmented epithelial cells, located between the choroid and neuroretina in the retina. Multiple functions, including phagocytosis, nutrient/metabolite transportation, vitamin A metabolism, etc., are conducted by the RPE on a daily basis. RPE cells are terminally differentiated epithelial cells with little or no regenerative capacity. Loss of RPE cells results in multiple eye diseases leading to visual impairment, such as age-related macular degeneration. Therefore, the establishment of an in vitro culture model of primary RPE cells, which more closely resembles the RPE in vivo than cell lines, is critical for the characteristic and mechanistic studies of RPE cells. Considering the fact that the source of human eyeballs is limited, we create a protocol to culture primary porcine RPE cells. By using this protocol, RPE cells can be easily dissociated from adult porcine eyeballs. Subsequently, these dissociated cells attach to culture dishes/inserts, proliferate to form a confluent monolayer, and quickly re-establish key features of epithelial tissue in vivo within 2 wks. By qRT-PCR, it is demonstrated that primary porcine RPE cells express multiple signature genes at comparable levels with native RPE tissue, while the expressions of most of these genes are lost/highly reduced in human RPE-like cells, ARPE-19. Moreover, the immunofluorescence staining shows the distribution of tight junction, tissue polarity, and cytoskeleton proteins, as well as the presence of RPE65, an isomerase critical for vitamin A metabolism, in cultured primary cells. Altogether, we have developed an easy-to-follow approach to culture primary porcine RPE cells with high purity and native RPE features, which could serve as a good model to understand RPE physiology, study cell toxicities, and facilitate drug screenings.

Here, an easy-to-follow method to culture primary porcine retinal pigment epithelial cells in vitro is presented.

The retinal pigment epithelium (RPE) is a monolayer of polarized pigmented epithelial cells, located between the choroid and neuroretina in the retina. ...

A Cost-Effective Method for Isolating Primary RPE Cells from Porcine Eyes

Isolation of Primary Porcine Retinal Pigment Epithelial Cells for In Vitro Modeling

The retinal pigment epithelium (RPE) is a crucial monolayer in the outer retina responsible for supporting photoreceptors. RPE degeneration commonly occurs in diseases marked by progressive vision loss, such as age-related macular degeneration (AMD). Research on AMD often relies on human donor eyes or induced pluripotent stem cells (iPSCs) to represent the RPE. However, these RPE sources require extended differentiation periods and substantial expertise for culturing. Additionally, some research institutions, particularly those in rural areas, lack easy access to donor eyes. While a commercially available immortalized RPE cell line (ARPE-19) exists, it lacks essential in vivo RPE features and is not widely accepted in many ophthalmology research publications. There is a pressing need to obtain representative primary RPE cells from a more readily available and cost-effective source. This protocol elucidates the isolation and subculture of primary RPE cells obtained post-mortem from porcine eyes, which can be sourced locally from commercial or academic suppliers. This protocol necessitates common materials typically found in tissue culture labs. The result is a primary, differentiated, and cost-effective alternative to iPSCs, human donor eyes, and ARPE-19 cells.

Author Spotlight: Modeling Retinal Pathologies with Enhanced RPE Cell-Based Disease Models

This protocol outlines the procedure for obtaining and culturing primary retinal pigment epithelial (RPE) cells from locally sourced porcine eyes. These cells serve as a high-quality alternative to stem cells and are suitable for in vitro retinal research.

The retinal pigment epithelium (RPE) is a crucial monolayer in the outer retina responsible for supporting photoreceptors. RPE degeneration commonly ...

Isolation and Culturing of Cells from Mouse Embryo Dorsolateral Telencephalons

Source: Landeira, B. S., et al. <a href="https://app.jove.com/v/56063">Live Imaging of Primary Cerebral Cortex Cells Using a 2D Culture System.</a> J. Vis. Exp. (2017)
This video demonstrates a protocol for isolating and culturing primary cerebral cortex cells from the dorsolateral telencephalon of mouse embryos.

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