This protocol was used to produce alpha synuclein pathology in rat brains by injecting them with Type 1 mouse alpha synuclein PFFs (SPR-324) and Type 1 human alpha synculein PFFs (SPR-322). The mouse PFFs showed greater seeding capabilities.

Animals used: female Sprague Dawley rats

PFFs and monomers were sonicated prior to injection.

16 ug PFFs or monomer (negative control) were delivered over two intraparietal sites (16 ug per injection site):

• AP + 1.6, ML + 2.4, DV – 4.2 from skull
• AP – 1.4, ML + 0.2, DV – 2.8 from skull

After 30 days, rats were saline perfused and brains were cut on the coronal plane to separate midbrain from forebrain. Both were then post-fixed in 4% PFA for 48 hrs.

 

DAB Protocol

Materials

• 1X TBS (with and without TX-100)
• 30% H₂O₂
• Sodium Citrate buffer
• Donkey serum (kept at -20; stable with freeze-thaws)
• Primary antibody: SMC-600 Alpha Synuclein Antibody (pSer129)
• Secondary antibody: Biotin-SP (long spacer) AffiniPure Donkey Anti-Rabbit IgG (H+L) (Jackson Immuno; Cat No: 711-065-152)
• ABC reagent (Vector; Cat No: PK-7100)
• DAB reagents (Vector; Cat No: SK-4100)
• Well plates (size depends on how many sections you are staining)
• Paintbrush(es)
• Glass hook(s)
• Bleach
• ddH₂0
• BD Luer-lock syringe 10 mL
• Filter 0.22 uM pore size (Fisher Cat. No. SLMP025SS)
• 100% EtOH
• Histoclear
• Microscope slides and coverslips
• Permount

 

Day 1:    (quenching, antigen retrieval, blocking, primary antibody)

-Rinse sections 5x with 1X TBS (shaking, RT) to remove cryoprotectant.

-Put sections in 0.6% H₂O₂ in TBS and leave on bench for 20 minutes, covered.

*DO NOT USE MESH WELLS FOR THIS STEP*

In _______ mL TBS, _______ uL 30% H₂O₂

-Rinse sections 3x with 1X TBS (shaking, RT)

-For antigen retrieval step, pre-warm sodium citrate buffer in water bath for at least 15-30 minutes. Incubate sections without shaking in sodium citrate buffer for 1 hour in an incubator set to 37⁰.

-Rinse sections 3x with 1X TBS (shaking, RT).

-To block, put sections in 5% normal donkey serum in TBS with .25% Triton-100. Block for 1 hour in the cold room, shaking.

*Note: You can make stock solution of TBS with Triton in it (1 L TBS, 2.5 mL TX-100), and then you only need to add donkey serum to that stock solution to block.*

In _______ mL TBS,

_______ uL donkey serum, _______ uL 10% TX-100

-Rinse sections 3x with 1X TBS (shaking, RT).

-Prepare primary antibody solution at 1:5000 in 5% donkey serum in TBS or as determined by researcher. Cover sections in foil and incubate in antibody overnight in the cold room, shaking.

In _______ mL TBS,

_______  uL donkey serum, _______ uL SMC-600 Alpha Synuclein Antibody (pSer129)

 

Day 2:    (secondary antibody, ABC amplification of signal, DAB staining)

-Rinse sections 3x with 1X TBST (shaking, RT). Leave sections in foil even during rinses for the rest of the protocol

-Prepare secondary antibody solution at 1:500 in 5% donkey serum in TBS. Incubate covered sections in foil in the cold room for 2 hours, shaking.

In ______mL TBS,

 ______ uL donkey serum, _______ uL biotinylated donkey anti-rabbit IgG

-Rinse sections 3x with 1X TBST (shaking, RT).

-Incubate sections in ABC reagent for 30 minutes (shaking, cold room).

-Rinse sections 3x with 1X TBS (shaking, RT).

-Prepare DAB solution using a falcon tube and ddH₂O, adding the following per 5 mL H₂O then vortex to mix:

2 drops buffer solution

4 drops DAB solution

2 drops hydrogen peroxide solution

*Note: Everything that DAB touches needs to be used exclusively with DAB. We also have a special disposal bottle for DAB under the fume hood. Use glass hooks NOT paintbrushes to handle the sections in the DAB and rinse hooks with bleach afterwards. Plates that had DAB in them are rinsed with bleach and disposed of in a biohazard bin.*

-Filter using a syringe into the wells of a new 6-well plate. Use a new well of DAB for each brain you are staining.

-Incubate each section in DAB until it is light brown (2-3 minutes is sufficient; leaving sections in longer may result in too much background)

-Rinse 3x using ddH₂O

-Mount sections in 1X TBS and leave to dry overnight in a drawer.

 

Day 3:    (dehydration and coverslipping slides)

*Note: Before starting the dehydration process, make sure all the glass dishes you will be dipping the slides in have enough volume to cover all tissue on the slides.*

-Dehydrate sections in the following order and duration:

1. 30 seconds ddH₂0

2. 3 minutes 70% EtOH

3. 3 min. 95% EtOH

4. 3 min. 95% EtOH

5. 3 min. 100% EtOH

6. 3 min. 100% EtOH

*ON SHAKER*

7. 5 minutes Histoclear

8. 5 minutes Histoclear

9. 5 minutes Histoclear

-Remove slides from slide basket one at a time and place on paper towel.

-To dispense Permount, get a P1000 pipette and tip, cut the end of the tip off, and set it between 400 and 500 μL.

-Add a few connected drops of Permount to the middle of slide and tilt coverslip over sections. Permount will spread throughout the slide.

-Use the wooden end of a cotton tip applicator to push out any bubbles from in and around sections.

-On paper towels, allow slides to dry overnight at the benchtop.

*Note: after ~24 hrs, excess Permount should be removed with Histoclear.*