MOUSE RETINA – IHC Cx36

Euthanasia / Fixation / Sectionning

Animals are euthanized by 100/10 mg/kg of K/X (IP) followed by cervical dislocation.

Eyes are rapidly removed and placed in 1X PBS (from 10X PBS, Fisher Scientific).

Eyes are opened (midline cut), either the posterior part of the eye (eyecup, used for sections) or the neural retina (whole-mounted retinas) is kept and rapidly fixed.

Fixation: 4% PFA/1X PBS for 2hrs (eyecup, sections) or for 30 min (whole-mounted retinas). Fixation must be done at room temperature (RT). Following fixation, keep the tissue overnight in PBS (1X) at 4℃ (fridge).

For sections with a cryotome: increase sucrose concentration in PBS (1X): 5%, 10%, 20%, 30% sucrose, at least 1h each, overnight for the last one (eyeballs must sink to the bottom of the recipient). Sections (20-um thick) are placed on slides. When all sections have been placed on a slide (up to 10), the slide is warmed for 30 min (in an hybridization oven) and then stored at -20℃, until processed for IHC.

For whole-mounted retinas or for floating 40-um thick sections cut with a microtome, keep the tissue in PBS (1X) at 4℃ until sectioned and/or processed.

Staining procedure (2-7 days)

Blocking step: take the slides out of the fridge/freezer and let them warm up to RT. Prepare the blocking solution.

Solutions: 1X PBS; 10% sodium azide (Na-az; in 1X PBS); normal donkey serum (NDS, from Jackson Immunoresearch); Triton-X100 (Tx)

Prepare 300 uL of blocking solution / slide (or per retina, floating sections and whole-mounts are placed in wells, in 48- or 96-well plates):

[blocking solution] = 0.2% Tx + 2.5% NDS + 0.1% Na-az + 1X PBS

[blocking solution / 300 uL] = 0.6 uL Tx + 7.5 uL NDS + 3 uL Na-az + 300 uL 1X PBS

Sequence of the steps:

Once sections/retinas have come to RT, place them in 1X PBS & prepare pieces of parafilm (will cover sections)
Initial washing = wash sections or retinas 3 x 5min in 1X PBS (in slide cup holder for sections on slides or in wells for wholemounts or floating sections)
Blocking = 2 hrs (sections) or overnight (wholemounts), RT - 300uL.
1. For sections on slides, use a box such as a slide box, place some PBS at the bottom to saturate the atmosphere, place the blocking solution on the sections (avoid direct pressure on the sections), use a piece of parafilm to cover the sections, and place the slides horizontally in the box above the PBS solution. Close the box. For floating sections or whole mounts, fill the empty wells with 1X PBS and close the lid of the plate (fill the empty wells with water or PBS).
After the incubation, delicately remove the parafilm or remove the blocking solution from the wells, rinse the slides/tissue in 1 X PBS, 3 x 5 min

Primary antibodies: Prepare 300 uL of solution / slide or retina, according to the following composition:

[primary Abs solution] = 0.2% Tx + 1% NDS + 0.1% Na-az + 1X PBS

[primary Abs solution / 300 uL] = 0.6 uL Tx + 3 uL NDS + 3 uL Na-az + 300 uL 1X PBS

Add primary Abs to the solution, for instance:

Anti-Cx36 monoclonal mouse (Chemicon, MAP3045; Loop) use 1:600

Anti-cArr polyclonal rabbit (Millipore, Ab15282) use 1:250

Anti-VGLUT1 polyclonal guinea pig (Synaptic Systems, 135304) use 1:3,000

5. Apply 300 uL / slide or retina, cover with parafilm (sections) or place in wells (wholemounts, floating sections) and incubate overnight (sections) or for 5 days (wholemounts) at RT

6. After incubation, delicately remove the parafilm, rinse the slides/tissue in 1 X PBS, 10 x 5 min (important step - do no speed it up!)

Secondary antibodies: Prepare 300 uL of solution / slide or retina, according to the following composition:

[secondary Abs solution] = 0.2% Tx + 1X PBS

[secondary Abs solution / 300 uL] = 0.6 uL Tx + 300 uL 1X PBS

Add secondary Abs to the solution, for instance:

Donkey-anti-mouse CY3 (IgG, JIR 715-165-150) use 1:600

Donkey-anti-rabbit Dylight488 (IgG, JIR 711-485-152) use 1:600

Donkey-anti-guinea pig (IgG, JIR 706-605-148) use 1:600

7. Apply 300 uL/slide, cover with parafilm (sections) or place in wells (wholemounts, floating sections) and incubate for 2h (sections) or 0.5-1 day (wholemounts) at RT IN THE DARK.

8. Rinse in 1X PBS, 3 x 10 min

9. Mount with coverslip, in medium with DAPI (Vector labs), 75-85 uL / slide, keep in the dark.

10. After 2 hrs, you can seal the coverslip with nail polish.

11. Wait until the nail polish is fully dry before imaging.

Tricks:

Prepare 10% sodium azide solution (about 1 mL) and keep it in the fridge (will be good forever).
For antibodies that need to be kept at -20℃, dilute the solution in ½ glycerol so that the solution doesn't freeze. But remember to adjust the concentration/volume of Ab solution to add (x2....).
Use slide cup holder/jars where you can place up to 10 slides vertically.
Use small plastic tubes or glass pieces to place between the coverslip and the slide when you mount wholemounts to prevent tissue damage.
Nail polish is best to seal (buy cheap one at your local pharmacy).
"In the dark" means that you need to prevent overexposure to light, not that you actually need to work in the dark
Immunosignal/tissue integrity is always better when PFA is used fresh.
Mount the wholemounts photoreceptor-side down (slide side) and ganglion cells up (coverslip side) for best imaging results of gap junctions in the OPL.
Adjust the gain/power/saturation of the microscope on the staining in the OPL and NOT in the IPL (IPL staining is usually about 8 times brighter). Use 63x to image rod/cone gap junctions.
Advise: z-stack 2-5 consecutive optical sections and collapse them.
Before applying the Ab solution or mounting medium, hold the slide vertically to let PBS fall down, and place the corner of the slide on a piece of filter paper while holding the slide vertically so that it absorbs the drop at the base of the slide. Do not try to remove all of remaining liquid on the slide, there is no need. DO NOT let the sections dry!!!!
Sodium-azide is a health hazard, proceed with care and in accordance with local rules/policies.