Протокол для оценки региональной патологии оптического нерва грызунов с использованием продольных криосекций и поперечной электронной микроскопии.

Before you begin

We published that T cells infiltrating the central nervous system (CNS) upstream of optic nerve pathology is concurrent with spinal cord white matter pathology, where experimental autoimmune encephalomyelitis (EAE) is traditionally studied. 1 This protocol describes the processing, imaging, and analysis of mouse optic nerves for the purposes of assessing whole-nerve pathology by immunofluorescence/immunohistochemistry and myelin/axon changes using electron microscopy in a model of autoimmune demyelination. This protocol can be adapted for any histochemical assessment of longitudinal mouse optic nerve sections or analysis of cross-sectional electron microscopy (EM) samples. Additionally, investigators may want to assess changes in CNS immune cell infiltration to explore whether genetic or pharmacological approaches are altering peripheral immune responses or directly impacting CNS pathology (please refer to our publication for further explanation 3 ).

Institutional permissions

All animal use and conditions prior to dissection were performed in accordance with National Institutes of Health (NIH) and the Institutional Animal Care and Use Committee guidelines of the Cleveland Clinic. Before performing any live animal work, all investigators must acquire permissions from their relevant institutions.

Transcardiac perfusion protocol for immunohistochemistry

Timing: ∼10–20 min per adult mouse

Prepare perfusion solutions. a. 1x phosphate buffered saline (PBS) pH 7.4 (included with 10x PBS recipe, Materials and Equipment ). i. Use a clean pH meter to check pH level and adjust with HCl or NaOH solutions if needed to reach desired pH. ii. Filter PBS with 0.2 μm filter before use. Note: Prepare a 10x solution of PBS (see 10x PBS, Materials and Equipment ) by dissolving reagents in 800 mL distilled water. Once all reagents are dissolved, add distilled water to a final volume of 1000 mL. Stock solution can be stored at 20°C–25°C for up to 12 months. A new solution should be prepared if precipitate is observed. 4 b. 4% paraformaldehyde (PFA) in pH 7.4 1x PBS. i. Dissolve 4 g PFA powder (Sigma Aldrich, St. Louis, MO; 158127) in 100 mL 1x PBS on a hot plate at 50°C–60°C with a stir rod. 1. Take care that the solution does not boil. ii. Cool PFA on ice. Use a clean pH meter to check pH level and adjust with HCl or NaOH solutions if needed to reach desired pH. iii. Filter cooled PFA with 0.2 μm filter before use. Note: For safety, always use powdered PFA in a ventilated fume hood (see materials and equipment ). Solution will not dissolve immediately—do not filter solution until it changes from a cloudy appearance to completely clear. Larger volumes may take a longer time to dissolve. Final PFA solution may be stored at 4°C for up to 7 days, however, for best perfusion results it is recommended to use within 24 hours.

1x phosphate buffered saline (PBS) pH 7.4 (included with 10x PBS recipe, Materials and Equipment ). i. Use a clean pH meter to check pH level and adjust with HCl or NaOH solutions if needed to reach desired pH. ii. Filter PBS with 0.2 μm filter before use. Note: Prepare a 10x solution of PBS (see 10x PBS, Materials and Equipment ) by dissolving reagents in 800 mL distilled water. Once all reagents are dissolved, add distilled water to a final volume of 1000 mL. Stock solution can be stored at 20°C–25°C for up to 12 months. A new solution should be prepared if precipitate is observed. 4

Use a clean pH meter to check pH level and adjust with HCl or NaOH solutions if needed to reach desired pH.

Filter PBS with 0.2 μm filter before use. Note: Prepare a 10x solution of PBS (see 10x PBS, Materials and Equipment ) by dissolving reagents in 800 mL distilled water. Once all reagents are dissolved, add distilled water to a final volume of 1000 mL. Stock solution can be stored at 20°C–25°C for up to 12 months. A new solution should be prepared if precipitate is observed. 4

Note: Prepare a 10x solution of PBS (see 10x PBS, Materials and Equipment ) by dissolving reagents in 800 mL distilled water. Once all reagents are dissolved, add distilled water to a final volume of 1000 mL. Stock solution can be stored at 20°C–25°C for up to 12 months. A new solution should be prepared if precipitate is observed. 4

4% paraformaldehyde (PFA) in pH 7.4 1x PBS. i. Dissolve 4 g PFA powder (Sigma Aldrich, St. Louis, MO; 158127) in 100 mL 1x PBS on a hot plate at 50°C–60°C with a stir rod. 1. Take care that the solution does not boil. ii. Cool PFA on ice. Use a clean pH meter to check pH level and adjust with HCl or NaOH solutions if needed to reach desired pH. iii. Filter cooled PFA with 0.2 μm filter before use. Note: For safety, always use powdered PFA in a ventilated fume hood (see materials and equipment ). Solution will not dissolve immediately—do not filter solution until it changes from a cloudy appearance to completely clear. Larger volumes may take a longer time to dissolve. Final PFA solution may be stored at 4°C for up to 7 days, however, for best perfusion results it is recommended to use within 24 hours.

Dissolve 4 g PFA powder (Sigma Aldrich, St. Louis, MO; 158127) in 100 mL 1x PBS on a hot plate at 50°C–60°C with a stir rod.

1. Take care that the solution does not boil.

Cool PFA on ice. Use a clean pH meter to check pH level and adjust with HCl or NaOH solutions if needed to reach desired pH.

Filter cooled PFA with 0.2 μm filter before use. Note: For safety, always use powdered PFA in a ventilated fume hood (see materials and equipment ). Solution will not dissolve immediately—do not filter solution until it changes from a cloudy appearance to completely clear. Larger volumes may take a longer time to dissolve. Final PFA solution may be stored at 4°C for up to 7 days, however, for best perfusion results it is recommended to use within 24 hours.

Note: For safety, always use powdered PFA in a ventilated fume hood (see materials and equipment ). Solution will not dissolve immediately—do not filter solution until it changes from a cloudy appearance to completely clear. Larger volumes may take a longer time to dissolve. Final PFA solution may be stored at 4°C for up to 7 days, however, for best perfusion results it is recommended to use within 24 hours.

Prepare anesthesia, perfusion, and dissection equipment. Note: If using inhaled anesthesia, equipment should include a vaporizer (such as VetEquip, Livermore, CA), an induction chamber, and tubing connected to a nose cone to maintain sedation during the procedure. Otherwise, prepare appropriate dosing of injectable anesthetic to allow sustained deep anesthesia throughout the procedure. 5 a. Attach a 25-gauge butterfly needle (BD Vacutainer, Fisher, 02-664-8 to the end of 20–30 mL syringes or tubing of a peristaltic pump (Fisher Scientific, Hampton, NH). i. For syringes: Fill one syringe with 20 mL cold 1x PBS and a separate syringe with PFA—the blood will first be flushed with PBS followed by fixation with PFA. 1. Prime the tubing of the butterfly needles with fluid to avoid air bubbles. ii. For peristaltic pump: provide separate beakers or jars containing 1x PBS and PFA, respectively. 1. Ensure that tubing is primed with fluid (1x PBS to start) to prevent air bubbles during perfusion. Note: There are several methods to perform transcardiac perfusion. We have listed two commonly used preparations (syringes and peristaltic pump). Another commonly used technique is gravity perfusion. Additional details can be found for this method. 6 b. Keep 1x PBS and PFA on ice. Note: Dissection equipment should consist of sharp dissection scissors and forceps. Hemostats and other preferred dissection equipment may be used as required.

Note: If using inhaled anesthesia, equipment should include a vaporizer (such as VetEquip, Livermore, CA), an induction chamber, and tubing connected to a nose cone to maintain sedation during the procedure. Otherwise, prepare appropriate dosing of injectable anesthetic to allow sustained deep anesthesia throughout the procedure. 5

Attach a 25-gauge butterfly needle (BD Vacutainer, Fisher, 02-664-8 to the end of 20–30 mL syringes or tubing of a peristaltic pump (Fisher Scientific, Hampton, NH). i. For syringes: Fill one syringe with 20 mL cold 1x PBS and a separate syringe with PFA—the blood will first be flushed with PBS followed by fixation with PFA. 1. Prime the tubing of the butterfly needles with fluid to avoid air bubbles. ii. For peristaltic pump: provide separate beakers or jars containing 1x PBS and PFA, respectively. 1. Ensure that tubing is primed with fluid (1x PBS to start) to prevent air bubbles during perfusion. Note: There are several methods to perform transcardiac perfusion. We have listed two commonly used preparations (syringes and peristaltic pump). Another commonly used technique is gravity perfusion. Additional details can be found for this method. 6

For syringes: Fill one syringe with 20 mL cold 1x PBS and a separate syringe with PFA—the blood will first be flushed with PBS followed by fixation with PFA.

1. Prime the tubing of the butterfly needles with fluid to avoid air bubbles.

For peristaltic pump: provide separate beakers or jars containing 1x PBS and PFA, respectively.

1. Ensure that tubing is primed with fluid (1x PBS to start) to prevent air bubbles during perfusion. Note: There are several methods to perform transcardiac perfusion. We have listed two commonly used preparations (syringes and peristaltic pump). Another commonly used technique is gravity perfusion. Additional details can be found for this method. 6

Note: There are several methods to perform transcardiac perfusion. We have listed two commonly used preparations (syringes and peristaltic pump). Another commonly used technique is gravity perfusion. Additional details can be found for this method. 6

Keep 1x PBS and PFA on ice. Note: Dissection equipment should consist of sharp dissection scissors and forceps. Hemostats and other preferred dissection equipment may be used as required.

Note: Dissection equipment should consist of sharp dissection scissors and forceps. Hemostats and other preferred dissection equipment may be used as required.

Anesthetize animal (e.g., isoflurane or preferred anesthetic). a. Confirm with toe pinch that animal is fully anesthetized before beginning. CRITICAL: Ensure that all procedures on live animals are performed in accordance with institutional guidelines for animal care and use. b. Place anesthetized animal in the supine position (on its back) on an aluminum tray with wax. Secure each paw to the wax with pins to ensure the animal remains lying flat. Note: For safety, perform all perfusion procedures in a fume hood. If using inhaled anesthetic, ensure that a nose cone is used and that the animal remains under deep anesthesia when secured to the wax.

Confirm with toe pinch that animal is fully anesthetized before beginning. CRITICAL: Ensure that all procedures on live animals are performed in accordance with institutional guidelines for animal care and use.

CRITICAL: Ensure that all procedures on live animals are performed in accordance with institutional guidelines for animal care and use.

Place anesthetized animal in the supine position (on its back) on an aluminum tray with wax. Secure each paw to the wax with pins to ensure the animal remains lying flat. Note: For safety, perform all perfusion procedures in a fume hood. If using inhaled anesthetic, ensure that a nose cone is used and that the animal remains under deep anesthesia when secured to the wax.

Note: For safety, perform all perfusion procedures in a fume hood. If using inhaled anesthetic, ensure that a nose cone is used and that the animal remains under deep anesthesia when secured to the wax.

Perform transcardiac perfusion in a fume hood. a. Pinch up skin of the abdomen with forceps and make a cut with scissors to expose underlying tissue. b. Cut along each side to completely open the abdominal cavity, revealing the rib cage. c. Lift the xiphoid process of the ribs and carefully cut the connected diaphragm to expose the heart and lungs. d. Cut up the center of the rib cage so that the heart becomes fully accessible. e. Carefully snip the animal’s right atrium of the heart—blood will begin flowing, allowing the release of perfused solutions. f. Quickly insert the perfusion needle (25-gauge, see key resources table ) into the left atrium of the heart and slowly perfuse 10–25 mL of cold 1x PBS (flow rate = 5–10 mL/min). g. After 1x PBS perfusion is complete, perfuse with 20 mL PFA.

Pinch up skin of the abdomen with forceps and make a cut with scissors to expose underlying tissue.

Cut along each side to completely open the abdominal cavity, revealing the rib cage.

Lift the xiphoid process of the ribs and carefully cut the connected diaphragm to expose the heart and lungs.

Cut up the center of the rib cage so that the heart becomes fully accessible.

Carefully snip the animal’s right atrium of the heart—blood will begin flowing, allowing the release of perfused solutions.

Quickly insert the perfusion needle (25-gauge, see key resources table ) into the left atrium of the heart and slowly perfuse 10–25 mL of cold 1x PBS (flow rate = 5–10 mL/min).

After 1x PBS perfusion is complete, perfuse with 20 mL PFA.

CRITICAL: Poor perfusion technique can decrease the quality of tissue for further processing ( Troubleshooting Problem 1 ). Ensure that the needle does not traverse the heart and use an appropriate flow rate to maintain the integrity of blood vessels and allow complete fixation of the tissue.

Transcardiac perfusion protocol for electron microscopy

Timing: ∼10–20 min per adult mouse

Prepare perfusion solutions in a fume hood. a. Prepare Modified Karnovsky’s fixative solution pH 7.4 (see materials and equipment ). i. Mix EM-grade PFA, EM-grade glutaraldehyde, and sodium cacodylate buffer to final concentrations of 4% PFA, 2% glutaraldehyde, and 0.1 M sodium cacodylate buffer in sterile Milli-Q H 2 O. ii. Use a clean pH meter to check pH level and adjust with HCl or NaOH solutions if needed to reach desired pH. iii. Filter modified Karnovsky’s fixative solution with 0.2 μm filter before use.

Prepare Modified Karnovsky’s fixative solution pH 7.4 (see materials and equipment ). i. Mix EM-grade PFA, EM-grade glutaraldehyde, and sodium cacodylate buffer to final concentrations of 4% PFA, 2% glutaraldehyde, and 0.1 M sodium cacodylate buffer in sterile Milli-Q H 2 O. ii. Use a clean pH meter to check pH level and adjust with HCl or NaOH solutions if needed to reach desired pH. iii. Filter modified Karnovsky’s fixative solution with 0.2 μm filter before use.

Mix EM-grade PFA, EM-grade glutaraldehyde, and sodium cacodylate buffer to final concentrations of 4% PFA, 2% glutaraldehyde, and 0.1 M sodium cacodylate buffer in sterile Milli-Q H 2 O.

Use a clean pH meter to check pH level and adjust with HCl or NaOH solutions if needed to reach desired pH.

Filter modified Karnovsky’s fixative solution with 0.2 μm filter before use.

Note: Final Karnovsky’s fixative may be stored at 4°C for up to 7 days, however, for best perfusion results it is recommended to use within 24 hours. For safety, use all hazardous chemicals such as paraformaldehyde, glutaraldehyde, and sodium cacodylate in a fume hood and wear proper personal protective equipment such as gloves, a lab coat, and safety goggles.

Prepare anesthesia, perfusion, and dissection equipment as detailed above (Step 2). a. In this step, Karnovsky’s fixative should be heated to 37°C for optimal perfusion.

In this step, Karnovsky’s fixative should be heated to 37°C for optimal perfusion.

Perform transcardiac perfusion. a. Open abdominal cavity and expose heart according to Steps 4a-b. b. Inject 100 μL 1% lidocaine into the left ventricle of the heart at this point to enhance capillary opening. 7 , 8 c. Carefully snip the animal’s right atrium of the heart—blood will begin flowing, allowing the release of perfused solutions. d. Quickly insert the perfusion needle into the left atrium of the heart and perfuse with 20 mL Karnovsky’s fixative at a flow rate of 5–10 mL/min.

Open abdominal cavity and expose heart according to Steps 4a-b.

Inject 100 μL 1% lidocaine into the left ventricle of the heart at this point to enhance capillary opening. 7 , 8

Carefully snip the animal’s right atrium of the heart—blood will begin flowing, allowing the release of perfused solutions.

Quickly insert the perfusion needle into the left atrium of the heart and perfuse with 20 mL Karnovsky’s fixative at a flow rate of 5–10 mL/min.

CRITICAL: High tissue quality is essential for EM. Injecting lidocaine before perfusion, using heated fixative, and keeping a steady flow rate will help preserve tissue integrity. A peristaltic pump is highly recommended to maintain flow rate. Indicators of successful fixation include muscle contractions and clearing of the liver (becomes pale) during perfusion, and complete stiffness of the body when perfusion is complete. This also applies for tissue perfused for IHC (Steps 1–4).

Key resources table

REAGENT or RESOURCE SOURCE IDENTIFIER Antibodies Rat anti-MBP (1:500) Abcam Cat# ab7349, RRID: AB_305869 Mouse anti-GFAP (1:1,000) BioLegend Cat# 835301, RRID: AB_2565344 Rabbit anti-IBA1 (1:500) Wako Cat# 019-19741, RRID: AB_839504 Biotinylated goat anti-rat IgG (1:200) Vector Labs Cat# BA-9400, RRID: AB_2336202 Alexa Fluor 488 goat anti-rabbit IgG (1:1,000) Invitrogen Cat# A11034, RRID: AB_2576217 Alexa Fluor 647 goat anti-mouse IgG (1:1,000) Invitrogen Cat# A21235, RRID: AB_2535804 Chemicals, peptides, and recombinant proteins Paraformaldehyde MilliporeSigma Cat# 158127 16% paraformaldehyde aqueous solution, EM grade Electron Microscopy Sciences Cat# 15711 50% aqueous glutaraldehyde EM grade Electron Microscopy Sciences Cat# 16320 0.2 M Sodium cacodylate buffer Electron Microscopy Sciences Cat# 11652 1% lidocaine Pfizer Hospital US/Hospira UoS NDC# 0409-4276-01 Sodium chloride (NaCl) MilliporeSigma Cat# 71376 Potassium chloride (KCl) MilliporeSigma Cat# P9541 Sodium phosphate dibasic dihydrate (Na 2 HPO 4 ·2H 2 O) MilliporeSigma Cat# 71643 Potassium phosphate monobasic (KH 2 PO 4 ) MilliporeSigma Cat# P9791 Sucrose MilliporeSigma Cat# S0389 O.C.T. compound Fisher Healthcare Cat# 23-730-571 Goat serum MilliporeSigma Cat# S26-100ML Triton X-100 MilliporeSigma Cat# T8787 Hydrogen peroxide Fisher Scientific Cat# H323-500 VECTASTAIN ABC kit Vector Laboratories Cat# PK-4000 DAB peroxidase substrate kit Vector Laboratories Cat# SK-4100 Xylene Fisher Scientific Cat# X3P-1GAL Fisher Chemical Permount mounting medium Fisher Scientific Cat# SP15 Fluoromount-G SouthernBiotech Cat# 0100-01 Bisbenzimide (1:1,000) Invitrogen Cat# H3569, RRID: AB_2651133 Immersion oil Immersion oil N/A Methanol Fisher Scientific Cat# A452-4 200-proof ethanol Fisher Scientific Cat# BP2818-4 Isopentane Fisher Scientific Cat# O3551-4 Pure Eponate 12 resin Ted Pella, Inc. 18005 Maleate buffer Electron Microscopy Sciences 18150 Osmium tetroxide Electron Microscopy Sciences 19190 Uranyl acetate Electron Microscopy Sciences 22400 Lead citrate Sigma L6258 Software and algorithms Fiji ImageJ https://imagej.net/Fiji Other Peristaltic perfusion pump Fisher Scientific Cat# 13-876-1 Aluminum tray with wax AutoMate Scientific Cat# 11-117 Isoflurane vaporizer system for small animal anesthesia VetEquip http://vetequip.com Cryostat Leica Biosystems N/A Leica SCN400 slide scanner Leica Biosystems N/A Microtome with diamond knife Leica Microsystems UC7 Tecnai G2 SpiritBT transmission electron microscope FEI Company N/A Confocal fluorescence microscope with 40x oil immersion objective lens Nikon N/A Experimental models : Organisms/strains Experimental autoimmune encephalomyelitis MOG 35-55 (50 μg, customized) Pertussis toxin (200 ng on days 0 and 2 of induction) Hooke Laboratories EK-2110 Mouse: C57BL/6J 8- to 10-week-old males Jackson Laboratory Strain #:000664 RRID:IMSR_JAX:000664

Materials and equipment

10x PBS Reagent Final concentration Amount NaCl 1.37 M 80 g KCl 27 mM 2 g Na 2 HPO 4 100 mM 17.8 g KH 2 PO 4 18 mM 2.4 g Dissolve all ingredients completely in 800 mL distilled H 2 O, then add more distilled H 2 O to final volume of 1000 mL.

Modified Karnovsky’s fixative Reagent Final concentration Amount 16% paraformaldehyde 4% 25 mL 50% glutaraldehyde 2% 4 mL 0.2 M sodium cacodylate buffer 0.1 M 50 mL 10x PBS 1x 10 mL Mix reagents and add distilled H 2 O to final volume of 100 mL. Heat to 37°C before use.

Blocking buffer (immunohistochemistry/Immunofluorescence) Reagent Final concentration Amount Goat Serum 10% 1 mL Triton-X 100 0.1% 10 μL 10x PBS 1x 1 mL Mix Triton-X 100 completely in serum and PBS, then bring to final volume of 10 mL with distilled H 2 O.

Step-by-step method details

Dissection of optic nerve and post-fixation

Timing: 1–3 days

This step describes how to carefully dissect the optic nerve followed by post-fixation and preparation for the next processing steps ( Figure 1 ).

Decapitate the mouse immediately following transcardiac perfusion (for either IHC or EM).

Using scissors, cut the scalp from the brainstem toward the snout to expose the skull.

Make small, shallow cuts up the middle of the skull. a. Using forceps, gently peel back each half of the skull to expose the brain.

Using forceps, gently peel back each half of the skull to expose the brain.

Gently push the brain up and locate the optic chiasm on the underside of the brain.

Cut the two optic nerves at the optic chiasm to separate them.

Enucleate the eyes and extract the optic nerves. a. Place forceps between the eye and the skull. b. Gently pull the eye (with optic nerve attached) out of the eye socket. c. Repeat for both eyes. d. Place in a 1.5 mL tube containing 4% PFA (IHC) or Karnovsky’s fixative (EM). Be sure to label and keep separate the left and right eyes/optic nerves.

Place forceps between the eye and the skull.

Gently pull the eye (with optic nerve attached) out of the eye socket.

Repeat for both eyes.

Place in a 1.5 mL tube containing 4% PFA (IHC) or Karnovsky’s fixative (EM). Be sure to label and keep separate the left and right eyes/optic nerves.

CRITICAL: Dissections should be performed carefully but quickly so as not to let the tissue dry at any point. This is especially important for tissue being processed for EM, as maintaining tissue integrity is essential for obtaining high-quality images.

Incubate in respective fixative overnight (12–16 hrs.) at 4°C.

The next day, wash the eyes with optic nerves in 3 x 15 min. in ∼5 mL 1x PBS. a. For IHC, after washing place the eyes/optic nerves in 30% sucrose solution in 1x PBS. This step is necessary for cryopreservation to avoid freezing artifacts. i. For 30% sucrose, dissolve 150 g sucrose (Sigma Aldrich, St. Louis, MO; S0389) in 500 mL 1x PBS. b. For EM, after washing place the eyes/optic nerves in 1x PBS and keep at 4°C until ready for next processing step.

For IHC, after washing place the eyes/optic nerves in 30% sucrose solution in 1x PBS. This step is necessary for cryopreservation to avoid freezing artifacts. i. For 30% sucrose, dissolve 150 g sucrose (Sigma Aldrich, St. Louis, MO; S0389) in 500 mL 1x PBS.

For 30% sucrose, dissolve 150 g sucrose (Sigma Aldrich, St. Louis, MO; S0389) in 500 mL 1x PBS.

For EM, after washing place the eyes/optic nerves in 1x PBS and keep at 4°C until ready for next processing step.

Note: Placing tissue in 30% sucrose is necessary for cryopreservation to avoid freezing artifacts. The eye and nerve will float when placed in sucrose solution. Once the tissue has been saturated with the sucrose solution it will sink to the bottom of the tube (∼48 hrs.). Tissue is now cryopreserved and ready for the embedding step.

Note: Tissue processed for EM can also be kept long-term at 4°C in EM fixative.

Dissection of optic nerve

(A) Following transcardiac perfusion and decapitation, an incision is made from the brainstem toward the snout to remove the scalp and expose the skull. Small cuts are made up the middle of the skull from the brainstem toward the snout, and each half is peeled back to expose the brain (B). The brain is gently lifted to reveal the optic nerves and tract, and the nerves are severed at the brain followed by separating the two nerves at the optic chiasm (C). The eyes are then enucleated with the nerves attached for post-fixation and processing (D).

Sample preparation and analysis for IHC

Timing: ∼1 h per sample

This step describes how to appropriately embed the optic nerve so that complete longitudinal cryosections can be obtained (Troubleshooting Problem 2 ).

Set up a freezing station to cryopreserve the optic nerves. a. Create a flat surface over dry ice.

Create a flat surface over dry ice.

Optional: The embedding mold can be placed in isopentane over dry ice to facilitate freezing. Be sure the isopentane is shallow enough in the freezing chamber so that it does not overflow into the embedding mold during freezing.

Detach the optic nerve from the eye and place in an embedding mold. a. Cut the optic nerve from the eye. i. Take care not to damage the back of the eye when cutting the nerve (if keeping eye for other analysis). ii. Keep track of which side of the nerve was attached to the eye versus the side projecting into the brain. iii. Label the embedding mold to maintain the orientation. b. Place freezing medium on the optic nerve. i. Mix 1:2 v/v 30% sucrose and Optimal Cutting Temperature (OCT) medium (23-730-571, Fisher HealthCare, Waltham, MA). For 30% sucrose, see Step 8ai. ii. Place enough freezing medium to completely cover the nerve iii. Ensure the nerve does not float and lies flat against the bottom of the mold. c. Freeze the optic nerve in the small amount of freezing medium. i. Once the small amount has turned completely white, fill the embedding mold with more freezing medium and allow to freeze completely (∼5 min.).

Cut the optic nerve from the eye. i. Take care not to damage the back of the eye when cutting the nerve (if keeping eye for other analysis). ii. Keep track of which side of the nerve was attached to the eye versus the side projecting into the brain. iii. Label the embedding mold to maintain the orientation.

Take care not to damage the back of the eye when cutting the nerve (if keeping eye for other analysis).

Keep track of which side of the nerve was attached to the eye versus the side projecting into the brain.

Label the embedding mold to maintain the orientation.

Place freezing medium on the optic nerve. i. Mix 1:2 v/v 30% sucrose and Optimal Cutting Temperature (OCT) medium (23-730-571, Fisher HealthCare, Waltham, MA). For 30% sucrose, see Step 8ai. ii. Place enough freezing medium to completely cover the nerve iii. Ensure the nerve does not float and lies flat against the bottom of the mold.

Mix 1:2 v/v 30% sucrose and Optimal Cutting Temperature (OCT) medium (23-730-571, Fisher HealthCare, Waltham, MA). For 30% sucrose, see Step 8ai.

Place enough freezing medium to completely cover the nerve

Ensure the nerve does not float and lies flat against the bottom of the mold.

Freeze the optic nerve in the small amount of freezing medium. i. Once the small amount has turned completely white, fill the embedding mold with more freezing medium and allow to freeze completely (∼5 min.).

Once the small amount has turned completely white, fill the embedding mold with more freezing medium and allow to freeze completely (∼5 min.).

Pause Point: Samples may be stored at −80°C until ready to be sectioned.

Note: Cryosectioning can be performed once the samples are completely frozen. Freezing is complete when the medium has turned completely white. As the initial freezing temperature is ∼−80°C, allow samples to acclimate in the cryostat chamber at -18°C−20°C for 20−30 min. before sectioning (if the embedded tissue is too cold, it will not section smoothly).

Section optic nerves on a cryostat (Leica Microsystems, Wetzlar, Germany). a. Set the cryostat to desired thickness (10−15 μm). b. Section the bottom face of the optic nerve block to obtain longitudinal sections. i. Be sure to maintain the orientation of the nerve. ii. If sections are retrieved from −80°C, allow to acclimate to cryostat temperature for 20–30 min. before beginning.

Set the cryostat to desired thickness (10−15 μm).

Section the bottom face of the optic nerve block to obtain longitudinal sections. i. Be sure to maintain the orientation of the nerve. ii. If sections are retrieved from −80°C, allow to acclimate to cryostat temperature for 20–30 min. before beginning.

Be sure to maintain the orientation of the nerve.

If sections are retrieved from −80°C, allow to acclimate to cryostat temperature for 20–30 min. before beginning.

CRITICAL: Be careful not to squeeze the optic nerve during transfer or detach from the eye to preserve tissue integrity. The nerve must also lay completely flat against the bottom of the embedding cup so that the sectioned surface cuts the full length of the optic nerve.

Pause Point: Sectioned slides may be stored at −80°C until ready to be further processed.

Myelin detection with 3′-3′-diaminobenzidine

Timing: 2 days

Myelin protein was targeted in optic nerve sections to evaluate areas of demyelination in experimental autoimmune encephalomyelitis (EAE).

Wash optic nerve sections 3 x 15 min. in 1x PBS + 0.1% Triton X-100. a. If sections are retrieved from −80°C, allow to come to room temperature (RT, 20°C–22°C) and dry completely. b. Bake slides for 10 min. on a hot plate at 37°C. i. After 10 min., turn off heat and allow.

If sections are retrieved from −80°C, allow to come to room temperature (RT, 20°C–22°C) and dry completely.

Bake slides for 10 min. on a hot plate at 37°C. i. After 10 min., turn off heat and allow.

After 10 min., turn off heat and allow.

Optional: If needed, antigen retrieval can be performed to promote better antibody detection. Incubate slides in pre-warmed sodium citrate buffer at 37°C for 30 minutes. Wash slides 3 x 15 min. in 1x PBS before proceeding to the next step.

Apply hydrophobic pen around the sections and completely cover sections in blocking buffer for 1 hr. at RT (20°C–22°C) in a humidified chamber. a. For blocking buffer recipe, see materials and equipment .

For blocking buffer recipe, see materials and equipment .

Tap off blocking solution and add primary antibody diluted in blocking buffer at desired concentration. a. Incubate overnight (12–16 hrs.) at 4°C in a humidified chamber.

Incubate overnight (12–16 hrs.) at 4°C in a humidified chamber.

Note: Blocking buffer should be stored at 4°C for up to 3 days. While the hydrophobic pen should retain its barrier throughout the staining process, be sure to check between incubation steps that no liquid is seeping across the barrier as this can lead to tissue drying. If there are issues, try gently aspirating the buffer rather than tapping it off.

The next day, tap off primary antibody and wash optic nerve sections 3 x 15 min. in 1x PBS + 0.1% Triton X-100.

Add horseradish peroxidase (HRP)-conjugated secondary antibody diluted in blocking buffer for 1 hr. at RT (20°C–22°C) in a humidified chamber. a. Near end of secondary incubation, prepare the avidin-biotin complex (ABC) solution (see key resources table ). b. Allow to mix gently on a rocker at RT (20°C–22°C) for 30 min. before use.

Near end of secondary incubation, prepare the avidin-biotin complex (ABC) solution (see key resources table ).

Allow to mix gently on a rocker at RT (20°C–22°C) for 30 min. before use.

Wash slides 3 x 15 min. in 1x PBS+0.1% Triton X-100.

Incubate slides in a Coplin jar with ∼45 mL 0.3% hydrogen peroxide in methanol for 10 min. at RT (20°C−22°C) to quench endogenous peroxidase activity.

Wash slides 2 x 15 min. in 1x PBS+0.1% Triton X-100 followed by 1 x 15 min in 1x PBS without Triton X-100 to wash away detergent.

Completely cover sections in ABC solution for 30 min. at RT (20°C–22°C) in a humidified chamber.

Wash slide 2 x 15 min. in 1x PBS and 1x 15 min. in distilled H 2 O.

Completely cover sections on one slide with DAB solution (see key resources table ) and record the time it takes for the stain to be sufficiently developed ( Troubleshooting Problem 3 ). a. Place slide in distilled water as soon as developed. b. Incubate each subsequent slide for the same amount of time and place in distilled water when finished.

Place slide in distilled water as soon as developed.

Incubate each subsequent slide for the same amount of time and place in distilled water when finished.

CRITICAL: In order for slides to be compared across a cohort, each must be incubated for the same amount of time in DAB solution. Incubation is complete when the very edges of the tissue become slightly darker than the rest of the sample.

Wash slides 3 x 5 min. in distilled water.

Dehydrate slides in a series of 200-proof ethanol (ETOH) and xylene washes in a Coplin jar at RT (20°C–22°C). Prepare each dehydration step in a separate jar and transfer slides quickly from one to the next. a. 70% ETOH for 2 min. b. 95% ETOH for 2 min. c. 100% ETOH for 2 min. d. 1:1 v/v ETOH and xylene for 2 min. e. 100% xylene for 2 min.

70% ETOH for 2 min.

95% ETOH for 2 min.

100% ETOH for 2 min.

1:1 v/v ETOH and xylene for 2 min.

100% xylene for 2 min.

Tap off excess xylene and coverslip slides using permount. a. Place a coverslip on top of sections, being careful to avoid forming air bubbles on tissue. b. Allow slides to dry on a flat surface overnight (12−16 hrs.) before imaging.

Place a coverslip on top of sections, being careful to avoid forming air bubbles on tissue.

Allow slides to dry on a flat surface overnight (12−16 hrs.) before imaging.

Pause Point: Slides can be stored at RT (20°C–22°C) until ready for imaging.

Slide scanning of myelin basic protein using DAB

Timing: ∼30 min per slide

This step provides instructions for acquiring images of entire longitudinal optic nerve section ( Figure 2 A and 2B).

Scan optic nerve sections from DAB detection steps (Steps 12–25) on a bright-field slide scanner (Leica SCN400, Leica Microsystems, Wetzlar, Germany). a. Use 40x magnification for high-powered imaging. b. Outline each optic nerve for faster scanning. c. Add focal points across the optic nerves for easier detection and focusing ( Troubleshooting Problem 4 ).

Use 40x magnification for high-powered imaging.

Outline each optic nerve for faster scanning.

Add focal points across the optic nerves for easier detection and focusing ( Troubleshooting Problem 4 ).

Note: Because the nerve sections take little space on the entire slide, outlining the individual sections as regions of interest will aid with scanner detection and focusing.

Analyzing demyelinated area after DAB staining in longitudinal optic nerve sections

(A) A slide scanner was used to image entire longitudinal sections of the optic nerve that were stained against myelin basic protein at 40x magnification from naïve mice and mice induced with EAE (B). Scale bars = 500 μm.

(C) These images were adjusted to 8-bit files for analysis and then traced in ImageJ/FIJI using the polygon tool (yellow circle), excluding meningeal tissue and any folds or damaged tissue (D). The traced area was saved as a region of interest (ROI) specific to each image for record.

(E and F) Thresholds were adjusted manually by an investigator blinded to treatment condition to analyze the percent demyelinated (unstained) area. Scale bars = 250 μm.

(G) Measurements were set to calculate the area fraction of demyelination based on the threshold and calculated by clicking “Analyze”→”Measure”. Percent area was recorded from the “Results” window for analysis.

Analysis of demyelination using DAB detection

Timing: ∼10 min per image

This step describes how to analyze whole optic nerve sections to measure demyelinated area between naïve and EAE-induced animals ( Figures 2 C–2G).

Open image in ImageJ/FIJI (NIH free software) and set the scale. a. Ensure that image is an 8-bit file, which is necessary to adjust the threshold. i. If needed, adjust by clicking “Image”→”Type→ “8-bit” ( Figure 2 C). b. Click “Analyze”→”Set Scale”. i. Use the straight line tool to draw a line of known distance (for example, either across a scale bar or across the length of the image). ii. Input the known distance and set the unit as microns. Click “OK”.

Ensure that image is an 8-bit file, which is necessary to adjust the threshold. i. If needed, adjust by clicking “Image”→”Type→ “8-bit” ( Figure 2 C).

If needed, adjust by clicking “Image”→”Type→ “8-bit” ( Figure 2 C).

Click “Analyze”→”Set Scale”. i. Use the straight line tool to draw a line of known distance (for example, either across a scale bar or across the length of the image). ii. Input the known distance and set the unit as microns. Click “OK”.

Use the straight line tool to draw a line of known distance (for example, either across a scale bar or across the length of the image).

Input the known distance and set the unit as microns. Click “OK”.

Draw a region of interest (ROI) to trace the entire optic nerve ( Troubleshooting Problem 5 ). a. Use the polygon tool to trace around the section ( Figure 2 C). b. Leave out the meningeal tissue of the nerve at the edges ( Figure 2 D). i. Do not include any tears or folds in the tissue. c. Add the ROI to the ROI manager by pressing the “t” key on the keyboard. d. Save ROI with a name specific to its respective image ( Figure 2 C)

Use the polygon tool to trace around the section ( Figure 2 C).

Leave out the meningeal tissue of the nerve at the edges ( Figure 2 D). i. Do not include any tears or folds in the tissue.

Do not include any tears or folds in the tissue.

Add the ROI to the ROI manager by pressing the “t” key on the keyboard.

Save ROI with a name specific to its respective image ( Figure 2 C)

Set the threshold of the image to detect the empty (unstained) space. This is indicative of the demyelinated area. a. Go to “Image”→”Adjust”→”Threshold” in the ImageJ/FIJI task bar ( Figure 2 E). i. Thresholds are set individually for each image by an investigator blinded to the treatment condition. ii. Make sure “dark background” is unchecked in the threshold window. iii. Fill all white pixels to assess demyelinated (unstained) area ( Figure 2 F).

Go to “Image”→”Adjust”→”Threshold” in the ImageJ/FIJI task bar ( Figure 2 E). i. Thresholds are set individually for each image by an investigator blinded to the treatment condition. ii. Make sure “dark background” is unchecked in the threshold window. iii. Fill all white pixels to assess demyelinated (unstained) area ( Figure 2 F).

Thresholds are set individually for each image by an investigator blinded to the treatment condition.

Make sure “dark background” is unchecked in the threshold window.

Fill all white pixels to assess demyelinated (unstained) area ( Figure 2 F).

Set measurements to include area fraction and other measurements that are desired. a. Go to “Analyze”→”Set Measurements” ( Figure 2 G). b. Click the relevant boxes that you would like to analyze (i.e., area fraction).

Go to “Analyze”→”Set Measurements” ( Figure 2 G).

Click the relevant boxes that you would like to analyze (i.e., area fraction).

Click “Analyze”→“Measure” and record the percentage of the demyelinated area from the “Results” window that is generated ( Figure 2 G).

Note: Although optic nerves from control conditions are not anticipated to have a “demyelinated” area, the thresholding will detect pixels containing no pigment, and since the optic nerve is not 100% myelin, there will be a small percentage of unstained area that is measured.

Fluorescent antibody staining to detect microgliosis and astrocytosis

Timing: 2 days

Traditionally, markers for microgliosis and astrocytosis have been used to evaluate inflammation in response to damage in the CNS. This protocol also utilizes area fraction analysis of staining for GFAP and IBA1 to evaluate changes in astrocytosis and microgliosis, respectively ( Figure 3 ). Of note, IBA1, which was used to identify resident microglia will also label infiltrating myeloid cells.

Wash slides containing optic nerve sections 3 x 15 min. in 1x PBS+0.1% Triton X-100. a. If sections are retrieved from −80°C, allow to come to RT (20°C–22°C) and dry completely before washing.

If sections are retrieved from −80°C, allow to come to RT (20°C–22°C) and dry completely before washing.

Optional: If needed, antigen retrieval can be performed to promote better antibody detection. Incubate slides in pre-warmed sodium citrate buffer at 37°C for 30 min. Wash slides 3 x 5 min. in 1x PBS before proceeding to the next step.

Apply hydrophobic pen around the sections and completely cover sections in blocking buffer for 1 hr. at RT (20°C–22°C) in a humidified chamber.

Tap off blocking solution and completely cover sections with primary antibody diluted in blocking buffer at desired concentration. a. Incubate overnight (12−16 hrs.) at 4°C in a humidified chamber.

Incubate overnight (12−16 hrs.) at 4°C in a humidified chamber.

The next day, tap off primary antibody and wash optic nerve sections 3 x 15 min. in 1x PBS+0.1% Triton X-100.

Add secondary antibody diluted in blocking buffer for 1 hr. at RT (20°C–22°C) in a humidified chamber protected from light.

Wash optic nerve sections 3 x 15 min. in 1x PBS+0.1% Triton X-100 protected from light.

Tap off excess wash buffer and add 75-100 μL mounting medium (Fluoromount-G (0100–01, SouthernBiotech, Birmingham, AL).

Place a coverslip on top of sections, being careful to avoid forming air bubbles on tissue.

Allow slides to dry for at least 1 hr. (protected from light) before imaging.

Pause Point: Fluorescent slides can be stored at 4°C protected from light until ready for imaging.

Image fluorescently stained optic nerve sections (Steps 33–40) on a fluorescent microscope ( Figure 3 A). a. Use 40x magnification for high-powered imaging. b. Capture at least 4 fields across the nerve. Note which end of the optic nerve from which the images were captured ( Figure 3 B).

Use 40x magnification for high-powered imaging.

Capture at least 4 fields across the nerve. Note which end of the optic nerve from which the images were captured ( Figure 3 B).

Perform area fraction analysis on reactive gliosis markers. a. Perform background subtraction as necessary using “Process”→”Subtract background” in ImageJ/FIJI before analyzing. Choose desired rolling ball radius for adequate background subtraction ( Figure 3 C). i. Background subtraction settings should be applied equally across all images. b. Ensure to check the “dark background” box in the threshold window ( Figure 3 D) c. Use the measurement function (Steps 30–31) to calculate the area fraction of reactive gliosis markers ( Figure 3 E).

Perform background subtraction as necessary using “Process”→”Subtract background” in ImageJ/FIJI before analyzing. Choose desired rolling ball radius for adequate background subtraction ( Figure 3 C). i. Background subtraction settings should be applied equally across all images.

Background subtraction settings should be applied equally across all images.

Ensure to check the “dark background” box in the threshold window ( Figure 3 D)

Use the measurement function (Steps 30–31) to calculate the area fraction of reactive gliosis markers ( Figure 3 E).

Note: When multiple markers have been stained together in a single section and are present in the same image, the area fraction analysis must be performed separately for each marker (e.g., IBA1 measurement is calculated, then step 42 is repeated for the channel containing GFAP to get the next area fraction measurement).

Analyzing reactive gliosis by immunofluorescence in optic nerve fields

(A) Images were taken on a fluorescent confocal microscope at 40x magnification.

(B) Four (4) image fields were selected to span the length of the longitudinal optic nerve section, including the region closest to the optic nerve head (1), the center regions of the nerve (2-3), and the end closest to the optic chiasm (4).

(C) Background was subtracted from images by adjusting the rolling ball radius, performed equally for all images.

(D) Threshold with the “dark background” setting was adjusted manually by an investigator blinded to treatment condition in order to measure the area fraction of positive staining for (E) IBA1+ microglia (green) and GFAP+ astrocytes (pink) in naïve optic nerves and nerves from mice induced with EAE. Scale bars = 20 μm.

Sample preparation and analysis for transmission electron microscopy

Timing: 3 days

EM tissue was processed in collaboration with the Cleveland Clinic Learner Research Institute Microscopy Core. The steps below outline the protocol in brief that was used to obtain optic nerve electron microscopy samples ( Troubleshooting Problem 6 ). Ensure that coronal cross-sections are collected from the sample for optimal axon and myelin analysis. More detailed information can be found for preparation and staining of optic nerves for EM. 9

Prepare optic nerves for EM. a. Cut optic nerve from the eye. b. Post-fix optic nerve in 1% osmium tetroxide in water for 60 min. at 4°C. i. Following incubation, wash samples 2 x 5 min. in 0.2 M sodium cacodylate buffer. ii. Rinse 1 x for 5 min. in 2% maleate buffer (pH 5.1). c. Incubate optic nerve samples in 1% uranyl acetate in maleate buffer (pH 5.1) for 60 min. at RT (20°C–22°C). i. Following incubation, wash samples 3 x 5 min. in 2% maleate buffer (pH 5.1). d. Dehydrate samples with ethanol. i. 30% ethanol for 5 min. at 4°C. ii. 50% ethanol for 5 min. at 4°C. iii. 75% ethanol for 5 min. at 4°C. iv. 95% ethanol for 5 min. at 4°C. v. 100% ethanol 3 x 10 min. at RT (20°C–22°C). e. Embed optic nerve Pure Eponate 12 resin (Ted Pella Inc., Redding, CA). i. Carefully decant or aspirate 100% ethanol and replace with 1:1 v/v 100% ethanol/Pure Eponate 12. ii. Incubate overnight (12–16 hrs.) at RT (20°C–22°C). iii. Following overnight incubation, change samples to 100% Pure Eponate 12 resin for 4–6 hrs. at RT (20°C–22°C). iv. Place samples in 100% Pure Eponate 12 resin in a rubber mold. Allow to polymerize for 24 hrs. at 62°C. f. Using a microtome with a diamond blade, cut ultrathin 85-nm section. g. Stain sections with 10% uranyl acetate and lead citrate. h. Mount sections onto an EM grid for imaging.

Cut optic nerve from the eye.

Post-fix optic nerve in 1% osmium tetroxide in water for 60 min. at 4°C. i. Following incubation, wash samples 2 x 5 min. in 0.2 M sodium cacodylate buffer. ii. Rinse 1 x for 5 min. in 2% maleate buffer (pH 5.1).

Following incubation, wash samples 2 x 5 min. in 0.2 M sodium cacodylate buffer.

Rinse 1 x for 5 min. in 2% maleate buffer (pH 5.1).

Incubate optic nerve samples in 1% uranyl acetate in maleate buffer (pH 5.1) for 60 min. at RT (20°C–22°C). i. Following incubation, wash samples 3 x 5 min. in 2% maleate buffer (pH 5.1).

Following incubation, wash samples 3 x 5 min. in 2% maleate buffer (pH 5.1).

Dehydrate samples with ethanol. i. 30% ethanol for 5 min. at 4°C. ii. 50% ethanol for 5 min. at 4°C. iii. 75% ethanol for 5 min. at 4°C. iv. 95% ethanol for 5 min. at 4°C. v. 100% ethanol 3 x 10 min. at RT (20°C–22°C).

30% ethanol for 5 min. at 4°C.

50% ethanol for 5 min. at 4°C.

75% ethanol for 5 min. at 4°C.

95% ethanol for 5 min. at 4°C.

100% ethanol 3 x 10 min. at RT (20°C–22°C).

Embed optic nerve Pure Eponate 12 resin (Ted Pella Inc., Redding, CA). i. Carefully decant or aspirate 100% ethanol and replace with 1:1 v/v 100% ethanol/Pure Eponate 12. ii. Incubate overnight (12–16 hrs.) at RT (20°C–22°C). iii. Following overnight incubation, change samples to 100% Pure Eponate 12 resin for 4–6 hrs. at RT (20°C–22°C). iv. Place samples in 100% Pure Eponate 12 resin in a rubber mold. Allow to polymerize for 24 hrs. at 62°C.

Carefully decant or aspirate 100% ethanol and replace with 1:1 v/v 100% ethanol/Pure Eponate 12.

Incubate overnight (12–16 hrs.) at RT (20°C–22°C).

Following overnight incubation, change samples to 100% Pure Eponate 12 resin for 4–6 hrs. at RT (20°C–22°C).

Place samples in 100% Pure Eponate 12 resin in a rubber mold. Allow to polymerize for 24 hrs. at 62°C.

Using a microtome with a diamond blade, cut ultrathin 85-nm section.

Stain sections with 10% uranyl acetate and lead citrate.

Mount sections onto an EM grid for imaging.

Note: Coronal cross-sections of the optic nerve are used for electron microscopy so that the myelin rings and axon structures can be easily visualized and measured. Therefore, the ultrathin samples that are obtained will be from a consistent region of the optic nerve across samples rather than a comprehensive regional analysis of the whole length of the nerve. For example, we consistently observe damage in the center of optic nerves during EAE and take random images of this region across samples.

Transmission electron microscopy

Timing: ∼30 min per sample

This step describes how to acquire images of optic nerves by TEM for use in determining ultrastructural myelin and axonal changes ( Figures 4 A and 4B).

Insert grid containing EM samples into electron microscope (Tecnai G2 SpiritBT, FEI Company, Hillsboro, OR).

Adjust the voltage to 60 kV and focus the microscope so that the myelin (which will appear black) is clearly visible.

Take 5-6 images sampling the entire cross-section at desired magnification.

Counting myelinated and unmyelinated axons in EM images of optic nerve cross-sections

(A) Samples were inserted into a transmission electron microscope and 5-6 fields were taken throughout the section at 6800x magnification.

(B) Representative images of electron micrographs from a naïve optic nerve and an optic nerve from an animal induced with EAE. Scale bars = 1 μm.

(C) To count myelinated and unmyelinated axons, the multi-point tool was used in ImageJ/FIJI.

(D) Multiple counter types can be used to count myelinated and unmyelinated axons separately.

(E) Representative images indicating myelinated axons (M, yellow) with dark myelin rings containing microtubules and neurofilaments in the oval-shaped axon, and unmyelinated axons (U, blue) which do not have any myelin. Scale bars = 1 μm.

Analysis of myelinated and unmyelinated axons

Timing: ∼15 min per image

This step describes the criteria for counting viable myelinated and unmyelinated axons in electron micrographs of optic nerve cross-sections ( Figures 4 C–4E).

Open image in ImageJ/FIJI (NIH free software) and set the scale (Step 27).

Use the image area to measure axon numbers per square millimeter. a. Ensure image area is in microns instead of pixels. i. Use the scale bar to set the measurement scale in ImageJ to microns. ii. Convert the image area to square millimeters.

Ensure image area is in microns instead of pixels. i. Use the scale bar to set the measurement scale in ImageJ to microns. ii. Convert the image area to square millimeters.

Use the scale bar to set the measurement scale in ImageJ to microns.

Convert the image area to square millimeters.

Use the multipoint tool to count myelinated and unmyelinated axons ( Figure 4 C). a. Separate markers can be used to denote myelinated and unmyelinated axons ( Figure 4 D).

Separate markers can be used to denote myelinated and unmyelinated axons ( Figure 4 D).

Note: For an axon to be counted in the image, it must be an oval shape and contain microtubules and neurofilaments in the axoplasm ( Figure 4 E). For myelinated axons, myelin will be a dark color (in contrast to the lighter gray color of the axon). Myelin must be intact to count the axon. The axon must not touch the outer edges of the image area. For standardized quantifications across a defined sampling size (if the entire image area is not desired), the Sampling Window plugin in ImageJ can be used to provide a grid area to aid with analysis. Additionally, the Cell Counter plugin is another tool to count multiple types of items (e.g., myelinated and unmyelinated axons) in the same image.

Record number of myelinated and unmyelinated axons. a. Divide the number of axons counted by the image area to calculate axons per square millimeter.

Divide the number of axons counted by the image area to calculate axons per square millimeter.

Repeat steps 47–49 for all images. Calculate the average number of axons per square millimeter for each animal (5-6 images per animal) to use for statistical analysis.

Expected outcomes

The visual system is frequently affected in MS, and visual system pathology has been shown to correlate with neuroaxonal damage in the brain and spinal cord in MS and preclinical MS disease models 1 , 10 , 11 , 12 It is therefore highly relevant to study components of the visual circuit when investigating mechanisms of neurodegeneration in MS, such as performed in our previous work. 1 , 2 This protocol allows for the analysis of changes across the entire length of the optic nerve. In this case, detection of areas of demyelination and reactive gliosis were captured to compare naïve animals with those induced with EAE. It is anticipated that ∼10 slides containing two (2) serial sections of full-length optic nerve per slide can be obtained when following this protocol. Additionally, high-powered images can be acquired for accurate detection and subsequent measurement of proteins and markers across the full length of the optic nerve or in smaller fields of view spreading the length of the nerve. By maintaining the orientation of the optic nerve, it is possible to draw conclusions about regional changes occurring near the optic nerve head versus the optic chiasm, as well as areas in between them.

When analyzing demyelination during EAE in C57BL6/J mice, the degree of demyelination will vary across optic nerves and experiments depending on how EAE is induced, the strain of mouse, how severely each mouse is affected, and at what time point the analysis is performed. However, it is anticipated that the area fraction of demyelinated (unstained) area in EAE mice will increase to approximately 7-12% of the analyzed area on average, based on our previous work. 1 , 2 It is also possible to obtain at least four (4) clear fields of view at 40x magnification that can be taken at the optic nerve head end, the optic chiasm end, and two areas in between to assess pathology (such as reactive gliosis) at high power and zoomed into particular regions of interest. This sampling also allows for detectable changes as long as it is not anticipated that only a small or specific part of the optic nerve will be affected.

For electron microscopy, it is recommended to take 5-6 random fields of view in a consistent area of the optic nerves across samples. While the sample will only be a small portion of the optic nerve, it is anticipated that in robust disease models such as EAE, alterations in myelin and axon integrity will be detected between experimental groups as shown in our previously published work. 1 , 2 For example, we consistently observe damage in the center of optic nerves during EAE and take random images of this region across samples. Numbers of axons that are able to be analyzed will depend on the magnification used. Based on our imaging protocol 6800x is an optimal magnification ( Figure 4 ). 1 , 2 There are approximately 4–6x10 4 myelinated axons/mm 2 on average per animal under control conditions.

Quantification and statistical analysis

There are several analyses that can be performed from the images obtained using this protocol. For slide-scanned optic nerve sections, there will be only (1) one image per nerve to assess and (2) two sections each for the right and left eyes. Average demyelinated area can be calculated per animal (and separate eyes can be assessed if binocular damage is not anticipated). The average percent demyelinated area for each animal can be used to perform the appropriate statistical analysis for the number of experimental groups (i.e., t test, ANOVA, etc.). For reactive gliosis, we previously measured the average percent area of the respective glia markers. Although optical density (for DAB detection) and fluorescence intensity can be subject to variability due to exposure time and variations in quenching, these can also be used when performed appropriately to determine the extent of change in a given tissue. 13

In electron micrographs of the optic nerve, myelinated and unmyelinated axons can be counted to determine if there is loss or a change in the distribution of different axon types. In the images obtained using this protocol, myelin thickness can be measured using g ratios (ratio of axon diameter to axon plus myelin diameter). The average number of axons or g ratios can then be analyzed across experimental groups. G ratios can also be used to determine whether there is a change in myelin for different sizes of axons, or if there is a change in overall distribution of axons depending on their size. 4 Finally, qualitative ultrastructural changes can be observed such as axonal swelling and myelin compactness.

Limitations

This protocol can be used to gain a comprehensive view of changes throughout the mouse optic nerve across different treatment conditions. Though the mouse optic nerve is small and can therefore be analyzed nearly completely, it is possible that a single longitudinal section will not capture changes that occur anterior or posterior to the section analyzed. This applies to longitudinal sections, but is more likely in cross-sections, as the sample collected will be small in relationship to the entire optic nerve. Additionally, while precautions are taken to avoid tissue artifacts, there are also limitations to immunostaining and staining for EM. Particularly when measuring loss of myelin (or loss of any protein), care should be taken to avoid counting tissue holes or artifacts as a pathological structural change.

Troubleshooting

Problem 1

Blood vessels are still visible in the brain after perfusing.

Potential solution

This would suggest that the tissue was not properly perfused and therefore tissue fixation may be suboptimal. To prevent this and ensure that all tissues are adequately perfused with PBS and fixative, make sure that the needle does not puncture completely through the heart (which would allow perfusion solutions to leak out of the heart rather than perfuse systemically). Also, be sure to maintain an appropriate flow rate so that blood vessels do not rupture during perfusion. It is possible that the needle can become clogged with tissue and therefore cannot permit continuous flow—check before each perfusion that fluids are flowing and change any clogged needle for a fresh one (See Transcardiac perfusion protocols for IHC and EM).

Problem 2

Longitudinal optic nerve cryosections appear in pieces rather than the entire nerve length.

Potential solution

This issue is caused when the face of the optic nerve that was sectioned was not completely flat. This can be prevented by using forceps to gently turn the optic until it is lying flat against the bottom of the embedding mold. It also may suggest that too much freezing medium was added at first, causing the nerve to float and become uneven at the cutting face while freezing. To prevent this, use only a drop of freezing medium to cover the nerve and make sure the nerve is completely touching the bottom of the embedding mold before freezing it. Then add additional freezing medium on top (See Steps 9–11).

Problem 3

Darkness of sections after DAB detection is starkly different between samples.

Potential solution

This can be caused by incubating sections in DAB solution for different times or an issue with the efficacy of reagents. Ensure that only one slide is developed first so that the time taken is recorded. Ensure that the application of DAB is staggered so that each slide is incubated for the same amount of time and is immediately placed in distilled water once the incubation time has ended. Finally, check the expirations of all reagents and ensure that they are working properly to allow for sufficient and high-quality antibody detection (See Steps 20-22)

Problem 4

Optic nerve images from the slide scanner are partly or fully out of focus.

Potential solution

Since the optic nerve is so small, it can sometimes be difficult for the slide scanner to find and focus on the regions of interest. It is possible that the nerve section is not completely flat on the slide. To prevent and remedy the issue, ensure that tissue sections are completely flat when collected onto the slide from the cryostat. When imaging on the slide scanner, draw a region of interest as close to the individual sections as possible to increase the area of focus for detection. Also, add more focal points across the length of each nerve section to aid in the scanner’s detection of the small tissue area (See Step 26).

Problem 5

The calculated percent area of demyelination is not reflective of what is observed subjectively.

Potential solution

It is possible that the area fraction of the demyelinated area in optic nerve sections appears much higher or lower than anticipated. To fix this issue, ensure that when drawing the ROI all torn areas are excluded from the analysis, as this can cause an increase in the demyelinated area. Additionally, care should be taken to trace the meningeal tissue out of the analyzed area, as this will appear dark but is not indicative of myelinated area. When thresholding, ensure that only the white (unstained) regions are included and are completely thresholded (See Steps 27–31).

Problem 6

The myelin in electron micrographs appears very faint and/or highly disorganized.

Potential solution

Detached and disorganized-appearing myelin may be caused by improper or suboptimal tissue perfusion and fixation ( Troubleshooting Problem 1 ). To prevent this, ensure that only a flow rate of 5–10 mL/min is used, and that the tissue has been completely perfused. Use heated fixative (37°C), and performing the optional step of injection 1% lidocaine into the heart before perfusion can help to improve fixation (Preparation Step 2). Following perfusion, ensure that dissection of tissue is performed quickly so as not to let the tissue dry before post-fixation. If the myelin appears faint, also check that the osmium tetroxide reagents were fresh and prepared properly. This can also be an artifact of post-processing. Tissue must be handled with care and embedded EM tissue must be cut with a fresh blade to help maintain tissue integrity (See Step 43). Additionally, ensure that the image field on the transmission electron microscope does not contain a hole or tear in the tissue, as this typically causes a bright spot that diminishes the detection of the surrounding tissue (See Steps 44–46).

Resource availability

Lead contact

Tara M. DeSilva, desilvt@ccf.org .

Technical contact

Gabrielle M. Mey, meyga@umich.edu .

Materials availability

All materials for this protocol are commercially available.

Data and code availability

This protocol does not include fully analyzed datasets, but includes sample images and data along with anticipated data based on previously published work.

Acknowledgments

Sponsored funding was received by the National Eye Institute
R01EY025687, R01EY032342, P30EY025585, and T32 EY024236; the National Multiple Sclerosis Society
FG-1807-31882 and FG-2108-38411; and the Department of Defense
HT94252410526. Electron microscopy tissue preparation was performed by the Cleveland Clinic Electron Microscopy Core.

Author contributions

T.M.D. and G.M.M. conceptualized the protocol and the manuscript. G.M.M. wrote and revised the manuscript and generated all the figures. J.S. and M.I. generated images for figures and revised the manuscript. T.M.D. wrote, revised, and provided final approval for the manuscript. All authors approved the final submitted manuscript.

Declaration of interests

T.M.D. is on the board of directors for Coeptis Therapeutics.

References

Associated Data

Data Availability Statement

This protocol does not include fully analyzed datasets, but includes sample images and data along with anticipated data based on previously published work.