Staining
Primary antibody (dilution)
Clone
Antigen retrieval
Blocking solution
Secondary antibody (dilution)
T cells (mouse)
Rat anti-human CD3e (1:50)
CD3-12
Citrate pH = 6
10 % FCS + 0.1 % Triton X-100
Biotin anti-rat (1:500)
Activated macrophages (mouse)
Rat anti-mouse CD107b (Mac3) (1:100)
M3/84
Citrate pH = 6
10 % FCS
Biotin-anti-rat (1:500)
T cells (rat)
Mouse anti-rat CD3 (1:50)
1F4
Citrate pH = 6
10 % FCS
Biotin-anti-mouse (1:200)
Macrophages (rat)
Mouse anti-rat CD86 (1:500)
ED1
Citrate pH = 6
10 % FCS
Biotin-anti-mouse (1:200)
Axonal damage (mouse, rat, and human)
Mouse anti-APP A4 a.a. 66-81 (1:2,000)
22C11
Citrate pH = 6
10 % FCS
Biotin-anti-mouse (1:200)
5.
Primary antibodies: Rat anti-human CD3e (clone CD3-12; cross-reaction with mouse CD3, AbD Serotec, Kidlington, UK), Rat anti-mouse CD107b (Mac3) (clone M3/84; Biolegend, San Diego, CA, USA), Mouse anti-rat CD3 (clone 1F4, AbD Serotec, Kidlington, UK), Mouse anti-rat CD68 (clone ED1, AbD Serotec, Kidlington, UK), Mouse anti-APP A4 a.a. 66-81 (clone 22C11, EMD Millipore, Billerica, MA, USA).
6.
Secondary antibodies: Biotin anti-mouse IgG (GE Healthcare, Little Chalfont, UK), Biotin anti-rat IgG (DCS Diagnostics, Hamburg, Germany).
7.
ExtrAvidin Peroxidase (Sigma-Aldrich, St. Louis, MO, USA).
8.
DAB developer solution: Dissolve 2 g 3,3′-Diaminobenzidine (DAB) tetrahydrochloride hydrate (Sigma-Aldrich, St. Louis, MO, USA) in 80 ml PBS, and store 1 ml aliquots at −20 °C. For the developer solution, dilute 1 ml of DAB in 49 ml PBS and add 20 μl of 30 % H2O2. Prepare developer solution always fresh.
9.
Optional: Vector® M.O.M.™ Immunodetection Kit BASIC (Vector laboratories, Burlingame, CA, USA):
(a)
M.O.M.™ Mouse Ig Blocking Reagent: add 2 drops of stock solution to 2.5 ml of PBS.
(b)
M.O.M.™ Diluent: add 600 μl of protein concentrate stock solution to 7.5 ml of PBS.
(c)
M.O.M.™ Biotinylated Anti-Mouse IgG Reagent: add 10 μl of stock solution to 2.5 ml of M.O.M.™ Diluent prepared above.
3 Methods
All procedures should be carried out at room temperature (RT) unless specified otherwise.
3.1 Perfusion
All animal experiments and interventions have to be performed after permission and in accordance with local authorities. Therefore, recommended anesthesia protocols may differ depending on the local regulations.
Paraformaldehyde (PFA) is toxic. Work under a ventilated hood and avoid inhaling vapors during perfusion and preparation of tissues.
1.
Prepare one bottle with 4 % PFA and one bottle with PBS (50 ml/mouse—200 ml/rat). Place a Styrofoam plate in a collection receptacle. Install a plastic tube attached to a syringe with a needle or a butterfly syringe in the peristaltic pump and fill it with PBS. Important: Avoid air bubbles. Adjust pump velocity to approximately 2 ml/min and turn off. Place the tube into 4 % PFA.
2.
Anesthetize the animal and verify anesthesia efficacy by squeezing the foot pads with tweezers.
3.
Place animal on the Styrofoam plate and attach limbs to it with an adhesive tape. Open the animal’s chest with small scissors. Insert the syringe needle into the left ventricle. Start the pump, and at the same time open the right atrium using small scissors. Perfuse the animal with approximately 30 ml/mouse or 150 ml/rat of 4 % PFA (see Note 3 ).
4.
Prepare a piece of spleen and place it into a 15 ml falcon tube filled with 8 ml of 4 % PFA. Prepare the cord and the head (attached to each other), remove all fur, and place it into the same 15 ml falcon tube.
5.
Postfix overnight at 4 °C. Then change to PBS for at least 3 h before tissue preparation.
3.2 Tissue Preparation and Processing
2.
Following the scheme provided in Fig. 1, cut the cerebrum carefully in four slices, the cerebellum in two pieces and the spinal cord in eight to ten pieces of approximately 3–5 mm length (see Note 5 ).
Fig. 1
Tissue processing and block arrangement. (a) The spinal cord is separated from the brain. By positioning the brain upside down, the crossing of the optical nerve is clearly visible. There you perform the first sagittal cut. Then, perform three additional posterior cuts 2 mm away from each other. (b) Cut the cerebellum into two pieces at an angle of approximately 45°. (c) HE stained paraffin section of an assembled tissue block
3.
Place all tissue pieces from one animal including spleen as internal staining control into an embedding cassette labeled with a block or identification number using a graphite pencil and close it. Place the embedding cassettes into a beaker containing PBS.
4.
Perform dehydration and paraffination procedure according to the manufacturer’s instruction of your paraffination and embedding system.
5.
Preheat paraffin and embedding molds and the heating plate to approximately 70 °C. Place the embedding mold on the heating plate and fill it with liquid paraffin. Open the embedding cassette containing the paraffinized tissue pieces on the heating plate. Discard the lid, but keep the labeled cassette.
6.
Following the scheme provided in Fig. 1 place cerebral and cerebellar slices, liver, and spleen as well as the spinal cord cone into the embedding mold. Take care about their orientation. Leave enough space for other spinal cord samples.
7.
Put all other spinal cord pieces upright on the heating plate, in the same orientation as they have to be placed into the embedding mold. Carefully remove the embedding mold now containing all tissue pieces except the spinal cords from the heating plate. When the paraffin in the embedding molds starts to solidify from the bottom, place spinal cord pieces upright into it using preheated tweezers.
8.
Place the labeled cassette on top, potentially add a bit more of the liquid paraffin on the top, and move the embedding mold to the cooling plate.
9.
10.
Using a microtome, cut 2 μm sections and let them outstretch on water. Mount sections on Superfrost® Plus Glass Slides and let dry.
3.3 Deparaffination
1.
Place slides with mounted paraffin sections (from here on referred to as slides) in a slide holder at 70 °C in the oven (20 min—overnight).
2.
From now on work under a ventilated hood. Prepare glass receptacles containing the following solvents: 3× Xylol/Ultraclear, 2× 100 % EtOH, 95 % EtOH, 90 % EtOH, 70 % EtOH, 50 % EtOH and ddH2O.
3.
For deparaffination, place the slide holder with the preheated slides sequentially into the three glass receptacles containing Xylol/Ultraclear. Incubate for 10 min in each receptacle (see Note 6 ).
4.
For rehydration, place slides sequentially into the glass receptacles containing decreasing concentrations of EtOH. At last place slide holder into the glass receptacle containing ddH2O. Depending on the staining to be performed, rehydration might have to be stopped before complete rehydration (specified in protocols, see below).
3.4 Histological Stainings
3.4.1 Hematoxylin/Eosin (HE)
1.
Deparaffinize and rehydrate sections until ddH2O.
2.
Filter Mayer’s hemalum working solution. Incubate slides in Mayer’s hemalum working solution for 30 s, and subsequently wash slides in ddH2O.
4.
Rinse slides in running tap water for approximately 10 min to “blue” staining. Verify nuclear staining under a light-optical microscope (see Note 8 ).
5.
Incubate slides in 1 % Eosin for 5 min, and then rinse slides extensively in ddH2O.
6.
Rehydrate slides by incubating in increasing concentrations of EtOH (50 % EtOH, 70 % EtOH, 90 % EtOH, 95 % EtOH, 2× 100 % EtOH) followed by 2× Xylol/Ultraclear for 30 s each.
7.
Using a xylol-based mounting medium coverslip slides, remove excess mounting medium by firmly pressing the coverslip on the slide.
8.
Let dry at RT overnight or at 37 °C for 1 h.
9.
Refer to Fig. 2 for anticipated staining results.
Fig. 2
Anticipated staining results. Representative staining of paraffin sections of mice during EAE (peak of disease). (a) Left panel: sagittal mouse spinal cord sections stained with HE (top), LFB/PAS (middle), and Bielschowsky’s silver staining (bottom). Please note the presence of inflammatory foci in the HE staining and the discrimination of white and grey matter in the LFB/PAS staining. Right panel: Visualization of an inflammatory EAE lesion (left: overview, right: higher magnification 200×) by HE (top), LFB/PAS (middle), and Bielschowsky’s silver stain (bottom). Please note the presence of compact small nucleated lymphocytes and macrophages visible in the HE staining, the demyelinated area visible in the LFB/PAS staining, and the profound axonal loss in the lesion visible in the Bielschowsky’s silver staining. (b) Left panel: Anti-CD3 staining in an inflammatory spinal cord EAE lesion developed with DAB and counterstained with hemalum. T cells are identified as brown-colored nucleated cells (magnification: 200×). The arrow indicates a CD3+ T cell as example. Right panel: Anti-Mac3 staining in an inflammatory spinal cord EAE lesion developed with DAB and counterstained with hemalum. Activated macrophages are identified as brown-colored nucleated cells (magnification: 200×). Arrow indicates a Mac3+ activated microglia/macrophage as example. (c) Anti-APP staining in an inflammatory spinal cord EAE lesion developed with DAB and counterstained with hemalum. The punctuated Anti-APP staining identifies axonal APP accumulation due to impaired axonal transport as a surrogate for axonal damage (left: overview, right: magnification 630×). Please note that APP+ axons do not show hemalum counterstaining!
3.4.2 Luxol Fast Blue (LFB): Periodic Acid Schiff (PAS)
1.
Deparaffinize and rehydrate sections until 90 % EtOH.
2.
Filter LFB solution and add a couple of drops of glacial acetic acid.
3.
Incubate sections in LFB solution in a cuvette at 60 °C between 1 and 2 days. Prevent evaporation of LFB solution by covering the cuvette with parafilm and lid.
4.
Prepare a cuvette with 90 % EtOH and three big petri glass dishes (a–c) with ddH2O (a), 0.05 % Li2CO3 (b), 70 % EtOH (c). Prepare an additional cuvette with ddH2O to store slides after differentiation (see Note 9 ).
5.
Transfer portions of four slides from the LFB solution to 90 % EtOH. Differentiate each slide individually by sequentially washing the slides in solutions (a)–(c): (a) Submerge slide with ddH2O. (b) Wash in Li2CO3 until slide does not emit blue color anymore. (c) Wash slide in 70 % EtOH. Repeat steps (a)–(c) until the staining turns light blue. Perform microscopic control to verify myelin staining (e.g., strongly stained corpus callosum, only faintly stained cortex, and strong blue staining of spinal cord white matter, in contrast to adjacent gray matter) (see Note 10 ).
6.
Transfer slides to a fresh cuvette with ddH2O (prepared in 4).
7.
Incubate slides for 5 min in 1 % periodic acid under a ventilated hood.
8.
Rinse slides for 5 min in running tap water.
9.
Rinse slides in three different cuvettes with ddH2O to remove all tap water.
10.
Prepare Schiff’s reagent in a fresh dry cuvette (fume hood). Incubate slides for 20 min in Schiff’s reagent. Verify under a light-optical microscope: faintly LFB-stained areas such as the cortex and spinal cord grey matter should have turned pink (see again Note 9 ).
11.
Rinse slides with running tap water for 20 min. Transfer slides into ddH2O.
12.
Costain slides with Mayer’s hemalum by dipping the slides five to ten times (depending on intensity of staining desired). Enhance nuclear staining by rinsing slides in running tap water for 5–15 min. Verify nuclear staining under a light-optical microscope.
13.
Rehydrate slides in increasing concentrations of EtOH (50 % EtOH, 70 % EtOH, 90 % EtOH, 95 % EtOH, 2x 100 % EtOH), incubate slides for 30 s each in 2× Xylol/Ultraclear.
14.
Using a xylol-based mounting medium coverslip slides, remove excess mounting medium by firmly pressing the coverslip on the slide.
15.
Acquire images of spinal cord sections using a light microscope equipped with a camera or a slide scanner using at least at a ocular of 20×.
16.
Analyze degree of demyelination as % demyelinated area/white matter (see below).
3.4.3 Bielschowsky Silver Impregnation Stain
Work under a ventilated hood.
1.
Deparaffinize and rehydrate sections. Store slides in ddH2O until staining.
2.
Prepare fresh 20 % AgNO3 solution.
3.
Prepare three cuvettes: (a) 20 % AgNO3, (b) ddH2O, (c) diluted ammonia (50 ml ddH2O + 1 ml NH3 conc.).
4.
Incubate sections in 20 % AgNO3 (a) for 20 min. Sections turn light to dark brown depending on their thickness.
5.
Rinse sections in ddH2O.
6.
Meanwhile add conc. NH3 to 20 % AgNO3 drop by drop (a) until the precipitate formed in step 4 has just disappeared (a*). Do not add excess ammonia, since it will inhibit impregnation.
7.
Incubate sections in 20 % AgNO3 + NH3 (a*) for 15 min protected from light.
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