Generation of Alpha-Synuclein Preformed Fibrils from Monomers and Use In Vivo

All methods involving animals have been approved by the Michigan State University Institutional Animal Care and Use Committee (IACUC).

1. Formation of α-synuclein preformed fibrils from monomers (Figure 1)

1. Thaw α-synuclein monomers on ice, gently resuspend by flicking the tube, and centrifuge at 15,000 x g for 10 min at 4 °C.
   NOTE: The α-syn monomer must be specifically formulated for fibrilization. Recombinant monomers can be purchased from commercial sources or generated by protocols on site^4,9,14,15. If purchased from commercial sources, the product must state that the α-syn monomer is specifically for the generation of fibrils. Regardless of if the monomers are purchased or generated on site, the quality control steps outlined below should be performed with each batch to ensure fibrils have formed and will seed efficiently prior to use in experiments.
2. Transfer supernatant to a clean 1.5 mL microcentrifuge tube and record the amount transferred.
   NOTE: Be careful to avoid the pellet, which, if present, will be small.
3. Measure the protein concentration of the transferred supernatant by either a standard bicinchoninic acid assay (BCA), or measuring the absorbance at 280 nm with a microvolume spectrophotometer.
   NOTE: The BCA assay is not as accurate for the specific measurement of α-syn and can yield results overestimating protein concentration. As a result, measuring the absorbance at 280 nm is the recommended method for determining protein concentration.
   1. To measure with the BCA assay, follow standard BCA protocols and perform with three different dilutions of α-syn monomer. Suggested dilutions are 1:25, 1:50, and 1:100.
   2. To measure with the A280 method, blank the reader with sterile 1x Dulbecco’s phosphate buffered saline (dPBS) without calcium and magnesium, with a salt concentration of approximately 100 mM NaCl, and pH range 7.0-7.3 (Table of Materials).
   3. Add 2 µL of sample to the reader and read the absorbance at 280 nm.
   4. Use the Beer-Lambert law to determine concentration of the monomers.
      
      NOTE: Extinction coefficient ε for human α-syn is 5,960 M^-1cm^-1 and for mouse α-syn is 7,450 M¹cm^-1. Pathlength is measured in cm. Molecular weight of α-syn (14 kDa) is estimated assuming 1 Da = 1 g/mol.
4. Dilute the monomers with 1x dPBS to a final concentration of 5 mg/mL. Use the equation below to calculate the amount of 1x dPBS added to dilute the monomers.
   
   NOTE: All concentrations are in mg/mL, and volumes in µL. Monomer dilutions should be performed with 1x dPBS. Total volume used to generate fibrils should be between 100 and 500 µL to achieve reproducible fibrilization results.
5. Briefly vortex to mix, and centrifuge to collect all liquid at the bottom of the tube. Aliquot monomers for quality control comparison with fibrils as indicated in steps 1.8.1 - 1.8.4.
6. Use a tube lock or wax/plastic film (Table of Materials) to secure the microcentrifuge tube lid closed.
7. Place the tube in an orbital thermomixer with a lid for 7 days at 37 °C, shaking at 1,000 RPM (Table of Materials).
   NOTE: At the end of the 7 days, the contents of the tube should appear turbid. The thermomixer must have a lid to prevent condensation formation on the tube lids.
8. Gently flick the tube to resuspend the α-syn fibrils. Aliquot fibrils for the following quality control steps as indicated in steps 1.8.1 - 1.8.4.
   NOTE: Fibrils for quality control steps can be stored at RT overnight.
   1. Aliquot 6 µL for the sedimentation assay.
   2. Aliquot 5 µL for the thioflavin T binding assay.
   3. Aliquot 2 µL for the transmission electron microscopy for fibril visualization.
   4. Aliquot at least 10 µL for the endotoxin assay.
      NOTE: For endotoxin assay, a commercial Limulus Amebocyte Lysate (LAL) is used, following manufacturer’s instructions (Table of Materials). Endotoxin levels should be ≤0.5 endotoxin units/mL for the fibrils. An endotoxin removal kit can be used if this criterion is not met.
9. Aliquot the remaining fibrils and store. For long term storage (12-18 months), rapidly freeze on dry ice, and store at -80 °C.
   NOTE: The amount to aliquot depends on the desired downstream application. In vivo use typically requires more fibrils at higher concentrations than in vitro use, and aliquot volumes should be planned accordingly. Fibrils should be stored towards the back of the freezer to prevent damage from potential freeze/thaw. The protocol can be paused here.

2. Sedimentation assay

1. In a clean microcentrifuge tube, add 6 µL of PFF to 54 µL of dPBS to dilute fibrils 1:10, and pipet to mix.
   1. In a separate tube, add 6 µL of monomer to 54 µL of dPBS as an additional control.
2. Centrifuge samples at 10,000 x g for 30 min at RT and transfer the supernatant to a clean microcentrifuge tube.
3. Add 60 µL of dPBS to the remaining pellet and vortex to resuspend.
4. Add 30 µL of 3x sample buffer (140 µL of 50% glycerol/0.1% bromophenol blue, 40 µL of 10% SDS, and 20 µL of β-mercaptoethanol) to each tube and vortex to mix.
5. Incubate samples at 100 °C for 10 min and allow to cool for 5 min on ice.
6. Use a standard sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) protocol to separate protein by mass. Use a protein ladder with a range that includes 14 kDa (the size band expected for monomeric α-syn).
7. Transfer the gel into a staining dish and cover the gel with Coomassie blue stain (0.1% Coomassie brilliant blue, 20% methanol by volume, and 10% acetic acid by volume in ddH₂O). Incubate for 3 h at RT while shaking.
8. Pour off the Coomassie blue stain and add enough destain (50% methanol by volume, and 10% acetic acid by volume in ddH₂O) to cover the gel. Incubate for 30 min at RT while shaking.
9. Pour off destain and repeat the destain step until the gel is clear and bands are visible.
10. Wash the gel 3 times for 5 min each by shaking in ddH₂O and image the gel.

3. Transmission electron microscopy for fibril visualization

1. Weigh 20 mg of uranyl acetate in a microcentrifuge tube and add 1 mL of ddH₂O to make a 2% aqueous solution. Vortex until the uranyl acetate is in solution, and allow to sit overnight.
   NOTE: Uranyl acetate should be made the day before preparing samples for transmission electron microscopy. Exposure to light or agitation can cause the uranyl acetate to precipitate, as such, the tube containing the uranyl acetate solution should be covered in aluminum foil, kept in a dark place, and not shaken after the uranyl acetate is in solution.
2. Add 2 µL of PFF to 98 µL of dPBS to dilute fibrils 1:50, and then pipet to mix.
3. Prepare a clean wax/plastic film (Table of Materials) covered surface (approximately 50 mm x 50 mm) on the bench top. On the clean wax/plastic film (Table of Materials), add the following (Figure 2).
   1. Add 4 x 10 µL drops of ddH₂O.
   2. Add 1 x 10 µL drop of diluted fibrils or monomer sample.
   3. Add 2 x 10 µL drops of 2% uranyl acetate.
4. Use fine-tipped forceps to pick up a formvar/carbon coated, mesh copper transmission electron microscopy grid (Table of Materials).
   NOTE: Be sure to only pick up the grid by the edge, avoiding the mesh portion in the center (Figure 2A). If the forceps touch the mesh, the formvar/carbon coated film will be damaged, making imaging in that area of the grid impossible.
5. Float the grid formvar/carbon coated side (shiny side) down on the first drop of ddH₂O. Gently use the forceps to hold the grid and push down to ensure the entire coated surface is in contact with the ddH₂O. Let the grid float for 1 min.
6. Pick up the grid, and without touching the mesh portion of the grid, wick away the ddH₂O with filter paper.
7. Float the grid as above on the second drop of ddH₂O for 1 min and wick away the ddH₂O.
8. Float the grid on the drop of diluted fibrils for 1 min and wick away the excess as above.
9. Float the grid on the first drop of uranyl acetate for 1 min and wick away the excess.
10. Float the grid on the second drop of uranyl acetate for 1 min, wick away the excess.
11. Float the grid as above on the third drop of ddH₂O briefly and wick away the ddH₂O.
12. Float the grid as above on the fourth drop of ddH₂O briefly and wick away the ddH₂O, being sure to remove as much ddH₂O as possible.
13. Transfer to a grid box for storage until imaging.
    NOTE: Grids should dry for at least 5 min before imaging. Grids can be imaged for at least a year after preparation and should be kept in a dry environment.

4. Thioflavin T assay

1. In a clean microcentrifuge tube, add 5 µL of PFF to 245 µL of dPBS to dilute fibrils 1:50, and pipet to mix.
2. Add 250 µL of dPBS to a separate microcentrifuge tube to serve as a negative control.
3. Add 5 µL of monomer to 245 µL of dPBS to serve as the monomer control.
4. Add 250 µL of thioflavin T in glycine buffer (25 µM thioflavin T, 100 mM glycine, 1% Triton X-100, pH 8.5) to each sample and gently mix.
5. Pipet 2 replicates, 200 µL each, into a black 96 well plate. Keep plate in the dark to prevent photobleaching.
6. Incubate for 1 h at RT and read the plate using an excitation of 450 nm and emission of 510 nm.
   NOTE: Thioflavin T readings will fluctuate over time. Samples should be read at the same incubation time. If desired, multiple readings can be taken during the hour incubation and plotted over time.

5. Sonication of α-synuclein preformed fibrils

CAUTION: The sonicator and all sonication steps are performed in a culture hood to prevent exposure to fibrils that may aerosolize during sonication. The personnel performing the sonication steps should wear personal protective equipment, including gloves, clothing protection in the form of a lab coat, and a face shield while sonicating. Risk of fibril exposure can be reduced by sonicating with a cup horn sonicator, allowing the tube containing fibrils to remain closed during sonication.  

NOTE: Optimal sonication parameters of fibrils are dependent on the model of sonicator used. For this reason, some optimization will need to be performed to ensure fibrils are the correct size. The sonicator used can be found in the Table of Materials and the parameters outlined are based on previous results with this model of sonicator. The parameters below will work for 2-4 µg/µL of PFFs in 200-400 µL of solution. Test sonication with the instrument should be performed and fibrils analyzed to ensure desired results are achieved prior to use of PFFs in experiments.

1. Attach a 3.2 mm diameter probe (Table of Materials) to the converter, and set the sonicator parameters as stated below in step 5.1.1 – 5.1.3.
   1. Set amplitude to 30%.
   2. Set the pulse to 01 01 (1 s on; 1 s off).
   3. Set the time to 0:01:00.
      NOTE: 1 min of pulsing equates to 60 pulses, and this model of sonicator (Table of Materials) will stop automatically. With other sonicator models, the number of pulses will need to be counted.
2. Thaw fibrils at RT and dilute with sterile dPBS in a culture hood.
   NOTE: A clear 0.6 mL microcentrifuge tube works best for sonication and the final fibril concentration depends on intended use. Representative results shown are from a final fibril concentration of 4 µg/µL.
3. Wipe the probe of the sonicator with a lab tissue dampened with 70% ethanol to clean the probe. Submerge the tip of the probe in ddH₂O and pulse 10 times to further clean the probe, and then wipe dry with a lab tissue.
4. Place the probe tip into the tube of diluted fibrils, and position the tip at the bottom of the tube.
5. Sonicate for 60 pulses (1 s on; 1 s off). Move the probe up and down during each pulse to ensure all of the fibrils in the liquid are sonicated. and clean.
6. After 60 pulses, remove the probe from the PFFs, and add 2 µL of PFF to 98 µL of dPBS in a clean microcentrifuge tube to dilute fibrils 1:50 for sonicated fibril measurement by electron microscopy.
   NOTE: Ideally, a small subset of fibrils should be measured prior to in vivo injection and a more comprehensive measurement of fibrils performed when time permits.
7. After sonication, briefly centrifuge PFFs for 1 s at 2,000 x g to collect all liquid off the sides of the tube.
   CAUTION: If the tube becomes hot to the touch, stop sonicating after 30 pulses, wait 1 min, and sonicate for the final 30 pulses.
   NOTE: While sonicating, keep the probe tip towards the bottom of the liquid at the start of each pulse, too close to the top of the liquid will cause sample loss.
8. Submerge the probe tip in 1% SDS and pulse 10 times to clean the probe. Remove the tip from the SDS, submerge in ddH₂O and pulse 10 times.
9. Wipe the probe with a lab tissue dampened with 70% ethanol, and then wipe the probe with a dry lab tissue. Detach the probe from the converter and store.
10. Wipe down all surfaces in the hood with 1% SDS, followed by 70% ethanol.
    NOTE: The 1% SDS solution is used to dissociate fibrils and clean surfaces and equipment¹⁶.

6. Transmission electron microscopy for the measurement of sonicated fibrils

NOTE: If electron microscopy is not feasible, a thioflavin T kinetics assay, and dynamic light scattering can be used as indirect measures of seeding efficiency and fibril size^4,14.  

1. Prepare samples using the protocol from the "Electron microscopy for fibril visualization" section above.
2. Use a transmission electron microscope to take an image of fibrils from 6 to 10 different grid openings.
   NOTE: Images should be a high enough magnification to measure fibrils, but low enough to visualize multiple fibrils simultaneously. A final magnification of approximately 75,000x should be sufficient.
3. Measure the length of a small subset of at least 25 fibrils prior to using fibrils in an experiment.
   NOTE: These measurements can typically be performed using the imaging software associated with the microscope. For the quick validation, the person measuring should select representative fibrils and calculate the average size. Fibril length should average around 50 nm or less, with a more accurate measurement to follow.
4. Measure the lengths of 500+ fibrils for more comprehensive results.
   NOTE: The imaging software associated with the microscope can be used for fibril measurements. Alternatively, images can be opened and fibrils measured with image processing software, using the scale bar associated with each image as a size reference with which to compare the fibril lengths.

7. Preparation of custom glass needle syringes for stereotactic injections (Figure 3)

1. Add approximately 10 mL of siliconizing reagent (Table of Materials) to a clean 50 mL beaker.
2. Place glass capillary tubes (length of 54 mm; outer diameter: 0.86 mm; inner diameter 0.59 mm) vertically in the beaker of siliconizing reagent and allow capillary action to draw the siliconizing reagent up the tube through the lower tube opening submerged in the siliconizing reagent.
3. Pipet additional siliconizing reagent into the upper tube opening that is not submerged in siliconizing reagent to completely fill the glass capillary tube.
4. Remove the capillary tubes from the siliconizing reagent and blot the open ends of the tubes on a paper towel to remove siliconizing reagent in the tubes. Allow capillary tubes to dry for at least 8 hours.
5. Place a siliconized glass capillary tube in a glass needle puller (Table of Materials).
6. Turn on the heating element and allow the attached weights to stretch the heated glass capillary tube.
7. Cut the pulled glass capillary tube with scissors at the thinnest point in the middle and remove the glass needle from the glass needle puller.
   NOTE: The pulled needle inner and outer diameter should be approximately 80 and 100 µm, respectively. Multiple glass needles can be made at one time and stored until ready to attach to a glass syringe with attached metal needle (Table of Materials).
8. Cut a length of shrink-wrap tubing (average inner diameter 0.021" and average wall thickness 0.001") to approximately 40 mm with scissors. Slide the shrink wrap over the metal needle of a 10 µL beveled syringe (Table of Materials).
9. Use an open flame to heat and adhere the shrink wrap to the needle while rotating the needle to apply heat evenly.
10. Slide the larger end of the pulled glass needle carefully over the metal needle of the syringe.
11. Cut a length of shrink-wrap tubing (average inner diameter 0.036" and average wall thickness 0.005") to approximately 40 mm with scissors and carefully slide over glass needle to overlap the base of the glass needle and the metal needle of the syringe (Table of Materials). Use an open flame to heat the shrink-wrap to secure the glass needle to the metal needle.
12. Add an additional layer of shrink-wrap to secure the glass needle. Cut a length of shrink-wrap tubing (average inner diameter 0.044" and average wall thickness 0.005") to approximately 40 mm with scissors and carefully slide over glass needle to overlap the base of the glass needle and the metal needle of the syringe (Table of Materials). Use an open flame to heat the shrink-wrap to secure the glass needle to the metal needle.
13. Trim the glass needle with scissors so that the tip is approximately 8 mm long.
    NOTE: The needle needs to be long enough to target the desired brain regions (required length is dependent on the dorsal/ventral coordinates).
14. Use steps 7.14.1 – 7.14.3 to test the needle to insure there are no leaks and there is adequate flow both in withdrawing and dispensing liquid from the glass needle.
    1. Fill a 1 mL syringe with an attached 26 gauge needle with dH₂O.
    2. Remove the metal plunger from the custom glass needle syringe and insert the needle of the dH₂O filled syringe into the base of the syringe. Apply pressure to dispense dH₂O from the glass needle. Inspect the interface of the glass needle and the metal needle for leaks and confirm a steady dH₂O flow.
       NOTE: If needed, the glass needle can be trimmed to increase the ease of flow and additional layers of shrink-wrap added to patch any leaks from the base of the glass needle.
    3. Fill a microcentrifuge tube with dH₂O. Use the custom glass needle syringe to draw in the dH₂O. Inspect the needle to confirm liquid is being taken into the syringe and that there are no bubbles.
       NOTE: If there are bubbles or dH₂O is not being drawn into the needle, trimming the needle can help alleviate pressure.
15. Carefully store syringe with attached glass needles in the syringe boxes until needed for surgeries.
16. Use standard stereotaxic surgery methods for intrastriatal delivery of PFFs at optimized coordinates in mice (one site: AP +0.2 mm and ML +2.0 mm from bregma, DV -2.6 mm from dura) or in rats (two sites: AP +1.6 mm and ML +2.0 mm from bregma, DV -4.0 from dura; AP +0.1 mm and ML +4.2 mm from bregma, DV -5.0 from dura)^1,14,17.
    NOTE: These coordinates have been used in C57BL6/C3H mice and Fischer 344 rats. When using other strains, coordinates should be optimized.