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Summary

This manuscript describes a detailed protocol for isolating retinal glial Müller cells from mouse eyes. The protocol starts with enucleation and dissection of mouse eyes, followed by isolation, seeding, and culturing of Müller cells.

Abstract

The primary supporting cell of the retina is the retinal glial Müller cell. They cover the entire retinal surface and are in close proximity to both the retinal blood vessels and the retinal neurons. Because of their growth, Müller cells perform several crucial tasks in a healthy retina, including the uptake and recycling of neurotransmitters, retinoic acid compounds, and ions (like potassium K+). In addition to regulating blood flow and maintaining the blood-retinal barrier, they also regulate the metabolism and the supply of nutrients to the retina. An established procedure for isolating primary mouse Müller cells is presented in this manuscript. To better understand the underlying molecular processes involved in the various mouse models of ocular disorders, Müller cell isolation is an excellent approach. This manuscript outlines a detailed procedure for Müller cell isolation from mice. From enucleation to seeding, the entire process lasts about a few hours. For 5-7 days after seeding, the media shouldn't be changed in order to allow the isolated cells to grow unhindered. Cell characterization using morphology and distinct immunofluorescent markers comes next in the process. Maximum passages for cells are 3-4 times.

Introduction

Müller cells (MCs) are the main and most abundant glial cells found in the retinal tissue. They are the key players in providing structural integrity and metabolic functions within the retina¹. The strategic structure of MCs is spread across the entire retina thickness, thus providing support to the retina. In addition to their scaffold-like properties, they have metabolic functions for retinal neurons, supplying them with energy substrates, including glucose and lactate. These functions are crucial in order to maintain healthy neuronal function. Impaired MCs have been reported to contribute to various retinal diseases, including age-related macular degeneration, diabetic retinopathy, and glaucoma^2,3. MCs can be endogenous cellular sources for regenerative therapy in the retina¹. They also comprise a significant portion of the retina, and strong evidence suggests that in several species, these cells can be stimulated to substitute missing neurons². They collaborate beneficially with neurons, and the conically branching Müller cells' endfeet densely unsheathe the blood vessels and connect the retina's neural components. In order to maintain neuronal development and neuronal plasticity, Müller cells act as a soft substrate for neurons, protecting them from mechanical trauma³. Additionally, under pathological circumstances, Müller cells may differentiate into neural progenitors or stem cells that replicate or regenerate the lost photoreceptors and neurons^2,3,4. Müller cells retain the characteristics of retinal stem cells, including different levels of potential for self-renewal and differentiation^5,6. The Müller glial cell has a significant retinal lineage that produces neurotrophic factors, uptakes and recycles neurotransmitters, spatially buffers ions, and maintains the blood-retinal barrier in order to keep the retina in homeostasis^7,8,9. This highlights the potential of Müller cells as a promising tool in cell-based therapies for treating diseases related to retinal degeneration. Müller cells are the primary glia distributed throughout the retina, connecting to both neurons and blood vessels. They play a crucial protective role, providing essential structural and metabolic support to maintain the viability and stability of retinal cells. Unfortunately, very few protocols are found in the literature for primary Müller cell isolation from the retina^10,11.

We present an enhanced approach to reliably isolating and culturing mouse primary Müller cells. This protocol was used in our group to isolate Müller cells from the wild-type C57BL/6 mice and transgenic mice^12,13. Mice aged between 5 and 11 days, with no sex preference, are used for this protocol. Cells have been passaged up to 4 times; however, at P4 they stop adhering to the flask, and it becomes difficult to grow a healthy culture. The culture is often contaminated with Retinal Pigmented Epithelial (RPE) cells, so the cells should be passaged at least once before performing any additional experiments on the cell line. Passaging allows for further isolation from contaminants. Therefore, the protocol presented offers a quick and effective way to isolate mouse Müller cells, which can then be used as a reliable platform to research therapeutic targets and evaluate potential treatments for retinal diseases¹⁴.

Protocol

All experiments with animals conformed to the ARVO statement for the Use of Animals in Ophthalmic and Vision Research and were done following our animal protocol approved by the Institute for Animal Care and Use Committee (IACUC) and Oakland University policies (protocol number 2022-1160)

1. Media and solution preparation

1. Prepare Müller cell eye solution by supplementing the Dulbecco Modified Eagle Medium (DMEM, details in the material table) with penicillin/streptomycin at the dilution of 1:1000. For instance, for 10 mL of DMEM, add 10 µL of penicillin/streptomycin.
   NOTE: This is a simple serum-free media prepared. Also, warm the complete media in a 37 °C water bath before using it for cell culture work.
2. Prepare complete primary Müller cell growth media by supplementing DMEM with 10% Fetal Bovine Serum (FBS, heat-inactivated), and 1% penicillin/streptomycin. For example (500 mL of DMEM media + 50 mL of FBS + 5 mL of penicillin/streptomycin).
3. Prepare a 0.05% Trypsin-EDTA solution with Collagenase IV at a concentration of 200 U/mL (Trypsin-EDTA is used to dissolve the calculated amount of Collagenase IV powder to reach the needed concentration).

2. Enucleation

1. Ethically euthanize the mouse using CO₂ inhalation following the IACUC guidelines.
   1. Briefly, place the animal(s) in the CO₂ chamber to introduce 100% CO₂ at a fill rate of 30-70% of the chamber vol/min with CO₂ to achieve the goal of rapid unconsciousness within 2-3 min with minimal distress to the mice.
   2. Since mice are at a young age (~10 days), perform cervical dislocation as a secondary method to ensure proper euthanasia.
2. After sterilizing the working area with 70% ethanol, place the mouse on a sterile under-pad.
3. Extract the eyes by placing the tweezers' arms at the back of the eye, gently pulling from the orbital area, and enucleating the eyes by applying pressure with 5/45 style tweezers to the orbital area to cause proptosis.
   NOTE: Sterilize the dissection tools with 70% ethanol followed by phosphate buffer saline prior to the dissection.
4. Ensure that the optic nerve is still connected to the eye by enucleating the eye slowly. Make sure to pull the optic nerve (visually identified as a white cord) behind the eye.
   NOTE: This step does not need to be performed under a microscope, and can be performed on a sterilized bench.

3. Treatment of the enucleated eyes

1. Cover a 15 mL centrifuge tube with aluminum foil and fill the tube with 10 mL of the Müller cell eye solution.
2. Place the dissected eyes in the 15 mL tube containing Müller cell eye solution and allow them to sit overnight, or about 18 h, at room temperature.
3. After 18 h, decant the eye solution and briefly rinse the eyes with 2-3 mL of warmed (37 °C) phosphate buffered saline (PBS).
4. Decant the PBS and move the eyes to a 100 mm Petri dish containing 10 mL of trypsin solution. (prepared in step 1.3).
   NOTE: Trypsin is used on the whole enucleated eye, acting as a dissociative agent to facilitate cell separation from the retinal layers during dissection. The dissected retina has many cells, including RPE cells, which are sticky and most likely to contaminate Müller cell culture¹⁴. Microscopic examination clearly demonstrated that after 5 days of isolation (during the first media change), RPE cells detached from the plate and died¹³.The media used for RPE isolation is DMEM/F-12, with a different composition than DMEM (used in Müller cells isolation), containing sodium pyruvate and low glucose.
5. Place the dish in an incubator and incubate for 1.5 to 2 h at 37 °C and 5% CO₂.
6. After incubating, transfer the eyes to a 60 mm Petri dish containing roughly 5 mL of complete primary Müller cell growth media.

4. Dissection

NOTE: This procedure must be carried out within the sterile environment of the culture hood. Hence, it is essential to thoroughly sterilize the hood surfaces with 70% alcohol, along with all the tools and the dissection microscope. It is worth noting that the video was recorded outside the hood for better visibility and clearer demonstration purposes.

1. Place a small piece of sterilized lab-grade tissue in the dish to help stabilize the eyes during dissection. Place the eyes under a dissecting microscope to start the dissection.
   NOTE: The use of lab-grade tissue facilitates the maneuvering of eyes in a fluid-filled dish (complete media), ensuring precise dissection.
2. Use Vannas scissors and M5S-style tweezers to remove the connective tissue, extraocular muscles, and optic nerve.
   1. Grasp the eye with M5S style tweezers and make an incision in the ora serrata section with Vannas scissors or a needle of a syringe.
      NOTE: There is a light blue circular line between the anterior and posterior part of the eye called ora serrata, which can be used as a landmark for accurate direction during dissection.
   2. Grasp the incision with M5S style tweezers and pull or tear the cornea from the posterior part of the eye. Pull in small increments to ensure the integrity of the retina remains intact.
      NOTE: These steps are to remove the anterior part of the eye (Cornea, Iris, and Lens) from the posterior part of the eye cup (neural and pigmented retinas).
   3. To maintain the integrity of the retinal layers, pull gently and remove the retinal pigmented epithelium from the neural retina. The neural retina should be free of any black specks to ensure the purity of the isolation as much as possible, as the RPE layer is usually sticky and attached to the neuronal retina.
   4. Cut the dissected neural retina into small pieces before collecting it in a plate to ensure a homogenous spread of the cells on the plate.
3. Plate the neural retinas in a T25 flask containing 4 mL of complete primary Müller cell growth media.
4. Finally, place the flask in an incubator and incubate at 37 °C and 5% CO₂.

5. Culturing of primary Müller glial cells

1. Do not move the flask for 3-5 days to promote cell growth.
2. Change the media after the fifth day and every other day thereafter until the cells are 90% confluent.

6. Passaging of primary Müller glial cells

1. Aspirate the Müller cell culture media, followed by 2 mL of wash in PBS, and aspirate out the PBS. Add 2 mL of 0.25% Trypsin-EDTA (1x), or enough to cover the plate.
2. Incubate for 5-10 min at room temperature.
3. Ensure the cells are detached from the plate under the microscope. Then, collect the cell suspension and add 4 mL of pre-warmed (37 °C) complete primary Müller cell growth media (prepared in step 1.2) to the collection tube.
4. Centrifuge for 7 min at 300 x g. After aspirating the supernatant, resuspend the pellet in 10 mL of complete primary Müller cell growth media. Because the cell number can vary according to the number of eyes dissected, therefore, it is better to count the cells before seeding and incubation. The recommended seeding density is 3.0 x 10⁶ cells in a T75 flask.
   NOTE: Cells can be passaged up to 4-5 times. However, the most consistent experiment results are found after 1-2 passages.

7. Immunofluorescence

NOTE: Use the immunofluorescence protocol to stain and validate Müller cell specificity. Here is a brief overview of the immunofluorescent protocol. This step is performed after the first passage¹².

1. Seed the cells into a Cell Culture Chamber Slide (30,000 cells per well for an 8-well chamber slide).
2. Culture the isolated cells for 24-48 h in a complete Müller cell culture medium (prepared in step 1.2) at 37 °C and 5% CO₂.
3. Aspirate out the media and wash the chamber slide with 250-300 μL of 1x PBS on each well when the cells are 80-90% confluent.
4. Fix the slide for 10 to 15 min in 250-300 μL of 4% paraformaldehyde on each well, followed by washing with PBS/TX-100 (5 min each/3x).
5. Add blocking buffer for 1 h at 37 °C (see the Table of Materials).
6. Aspirate the blocking solution, then add the primary antibodies specific for Müller cells to confirm the purity of the isolated cells using glutamine synthetase and vimentin^5,12. Incubate for 3 h at 37 °C (using an antibody concentration of 1:100-1:200 in blocking buffer in a volume of 150-200 μL/slide). Wash the slide with PBS/TX-100 washes (5 and 10 min each).
   NOTE: Those antibodies were chosen as they are hallmarks for identifying Müller cells and ensuring the success and purity of the Müller cell culture.
7. Add the secondary antibody (with an antibody concentration ranging from 1:1000 in blocking buffer in a 150-200 μL/slide) and incubate for 1 h at 37 °C. Then, wash with PBS/Triton X-100, three times (5-10 min each).
8. Apply a drop of DAPI mounting medium to label the nuclei before covering the slide with a coverslip. Avoid air bubbles when placing the coverslip.
9. Use a fluorescent microscope to check the slide and capture images (see Table of Materials).
   NOTE: Cells are located with DAPI at ex/em 359/461 at 10x magnification to locate cells; higher magnification can capture cells. Other wavelengths would depend on the chosen color. Green color (GFP)- ex/em 488/510. Red color (Cy3)- ex/em 555/569.

Representative Results

Validation of the specificity, purity, and barrier function of isolated Müller cells
To confirm the viability, morphology, and distinctive qualities of the isolated Müller cells, the cells were examined under a light microscope. P0 and P1 images were recorded (Figure 1A). To check the contamination of isolated Müller cells with RPE cells and confirm it's purity, immunofluorescence staining (IF) was performed using antibodies specific to RPE cell marker, RPE65. Human retinal pigmented cells (ARPE -19) were used as a positive control. RPE65 stained RPE cells (red and blue for nuclear staining) are shown in Figure 1B (lower panel). RPE65 specific antibodies did not stain the isolated Müller cells (Figure 1B, upper panel). The fact that the RPE65 stained the ARPE-19 cells (red) and didn't stain the isolated cells, indicates that the isolated cells are not RPE cells. The presence of pyruvate and low glucose in the media establishes unfavorable conditions for the growth of RPE cells. Consequently, even in the event of RPE cell contamination, they will eventually detach from the flask.

Furthermore, to confirm the identity of isolated cells, a Müller cells immunofluorescence staining for Müller cells specific markers were used to confirm the successful isolation of Müller cells. The cytoskeleton intermediate filament protein called vimentin was used^12,13. Additionally, glutamine synthetase (GS), that catalyzes the reaction of condensation of glutamate and ammonia to form glutamine, as a key function of retinal glial Müller cells, was also used⁵.

Collectively, Isolated cells stained negative for RPE65 (red, Figure 1B), positive for vimentin (green, Figure 1C) and GS (green, Figure 1D). IF staining confirms successful isolation (purity and specificity)of the isolated Müller cells. Figure 1E, is a negative control for vimentin (upper panel) and GS (lower panel), confirming the specificity of the antibodies.

Figure 1: Validation of Müller cell isolation (purity and specificity). (A) Light microscopy image for passages:passage zero(P0)and(P1)morphology. (B) RPE65 immunostaining combined with DAPI nuclear staining. (C) Vimentin immunostaining with DAPI nuclear staining at high magnification. (D) GS immunostaining at the same high magnification. (E) negative controls to confirm the specificity of vimentin and GS antibody staining. scale bar = 300 μm, 300 μm, 50 μm, 20 μm, 20 μm, 50 μm, 50 μm, respectively. Please click here to view a larger version of this figure.

Discussion

The isolation of primary retinal pigmented epithelium (RPE) from mice was previously documented by our lab. This manuscript describes a detailed demonstration protocol for primary Müller cells isolation. This procedure involves enucleation, treatment, dissection, collection, seeding, culture, and characterization of Müller cells isolated from mouse eyes. It is based on a previously successful protocol found in earlier publications and our modified protocol that we used in a recent publication^12,13. While the steps are straightforward, successful Müller cell isolation requires adherence to specific restrictions and essential requirements. After numerous trial and error attempts to optimize this protocol, multiple factors influencing its success were identified.

The ideal mouse age, as observed in our experiments, was found to be between 5-11 days in order to grow the isolated cells up to four passages. The culture is also influenced by how many eyes are isolated and cultured. For single isolation, at least two eyes are required to obtain sufficient number of cells to be able to grow and multiply. Another important aspect is the duration between enucleation and plating; therefore, this step needs to be performed relatively quickly. It was observed that prolonged manipulation of the eye tissue has a detrimental effect on the cells' ability to adhere to the flask and multiply. To make sure that the proximity of isolated cells is close enough to encourage proliferation, we advise using a T25 flask. Once the first passage has been made, the cells can be plated on a T75. Also, cells should not be disturbed after isolation and seeding by removing the tissue culture flask from the incubator or by switching the media for at least 5 days. Finally, cells can only be passed about four times before they stop adhering. Although this procedure is straightforward and easy to perform, it has some limitations including the mice ages, limited number of cell passages, etc.

There aren't many protocols for Müller cell isolation, but there are some notable differences from this protocol to the protocols that are available^5,14. First, the time between enucleation and isolation is significantly different. Other protocols suggest dissection soon after enucleation, typically within 10 min. Whereas the current protocol suggests a resting period of at least 18 h as this provided the best results for isolation in this protocol. Another significant difference between this protocol and other protocols is the point in which the eyes are treated with enzymes. In this protocol, the eyes are treated before any dissection has been performed, even before the removal of connective tissue and extraocular muscles. While other protocols isolate the retinas from the eye prior to enzyme treatment. Furthermore, the enzyme recipe is also different. In this protocol, the enzyme solution consists of trypsin and collagenase type IV. In similar protocols, papain and DNase are used for tissue digestion. Additionally, the culture vessel is different, this protocol suggests culturing on a T25 then on a T75 after the first passage. While other protocols use Petri dishes or multi-well plates. Finally, this protocol suggests that the flask should not be disturbed (for 5-7 days) to promote cell growth. Whereas in other protocols media change is performed after one day.

Materials

Beaker : 100mL	KIMAX	14000
Collagenase IV	Worthington	LS004188
Disposable Graduated Transfer Pipettes :3.2mL Sterile		13-711-20
DMEM (1X)	Thermo Scientific	11885084	Media to grow Müller cells
Fetal Bovine Serum (FBS)	gibco	26140079	For complete Muller cell culture media
Glutamine synthase	Cell signalling	80636
Heracell VISO 160i CO2 Incubator	Thermo Scientific	50144906
Kimwipes	Kimberly-Clark	34155
Luer-Lok Syringe with attached needle 21 G x 1 1/2 in., sterile, single use, 3 mL	B-D	309577
Micro Centrifuge Tube: 2 mL	Grainger	11L819
Pen Strep	gibco	15140-122	For complete Müller cell culture media
Phosphate Buffer saline (PBS)	Thermo Scientific	J62851.AP
Positive Action Tweezers, Style 5/45	Dumont	72703-DZ
Scissors Iris Standard Straight 11.5cm	GARANA INDUSTRIES	2595
Sorvall St8 Centrifuge	ThermoScientific	75007200
Stemi 305 Microscope	Zeiss	n/a
Surgical Blade, #11, Stainless Steel	Bard-Parker	371211
Suspension Culture Dish 60mm x 15mm Style	Corning	430589
Tissue Culture Dish : 100x20mm style	Corning	353003
Tornado Tubes: 15mL	Midsci	C15B
Tornado Tubes: 50mL	Midsci	C50R
Tweezers 5MS, 8.2cm, Straight, 0.09×0.05mm Tips	Dumont	501764
Tweezers Positive Action Style 5, Biological, Dumostar, Polished Finish, 110 mm OAL	Electron Microscopy Sciences Dumont	50-241-57
Underpads, Moderate : 23" X 36"	McKesson	4033
Vannas Spring Scissors – 2.5mm Cutting Edge	FST	15000-08
Vimentin	invitrogen	MA5-11883
Zeiss AxioImager Z2	Zeiss	n/a
Zeiss Zen Blue 2.6	Zeiss	n/a