Materials

1. Cells must be plated on a high precision #1.5H coverslip or ibidi petri dish. If you are using the 8-well one, plate cells on the center four wells for imaging.
   
2. Calibration bead slide. For detailed instructions on how to prepare bead slides, check here.
   
3. dSTORM imaging buffer, which can be procured from ONI. Please contact the CX team for getting a quote.
   
4. 4% paraformaldehyde in 1X PBS for fixation.
   
5. 5% BSA + 0.1% Triton X-100  in 1X PBS for blocking and antibody dilution.
   
6. 0.1% NaBH4 (w/v) in 1X PBS for quenching.
   
7. Triton X-100 0.3% for cell permeabilization.
   
8. Conjugated primary antibodies. These antibodies are used and recommended by ONI for staining:
   
1. For the red channel (i.e. the 640 laser line): Alexa 647 (the best dye for dSTORM).
   
2. For the green channel (i.e. the 561 laser line): Alexa 555, DyLight 555, and CF 568 (highly recommended).
   
3. For the blue channel (i.e. the 488 laser line): Alexa 488, Atto 488 (recommended), and CF 488A (recommended).
   

Instructions

1. Plate cells on a #1.5 coated glass coverslip or glass bottom ibidi chamber at a confluence of about 30~50%. Incubate for 16~24 hours, afterward wash cells briefly with 1 mL of pre-warmed PBS (37°C). If you are using coverslips to plate cells, make sure that longer incubation steps (blocking, primary and secondary antibodies) are done in humidity chambers to prevent excessive evaporation and drying of the sample.
   
2. Fix cells using 4% Paraformaldehyde for 15 minutes.
3. Treat the sample with 1 mL 0.1% (w/v) NaBH4 (freshly-prepared in 1X PBS) for 7 minutes to reduce the background fluorescence. Wash with 1X PBS three times for 10 minutes each.
   
4. Permeabilize cells with 0.3% Triton X-100 in 1X PBS for 10 minutes.
   
5. Block for 30 minutes with 5% BSA + 0.1% TritonX-100 in 1X PBS.
   
6. For primary antibody staining, dilute the primary antibodies in the blocking buffer and incubate for 1 hour, followed by three washes of 10 minutes with the blocking buffer. Recommended primary antibody dilution:
   
1. HSP60 1:50
   
2. TOM20 1:50
   
3. Phalloidin 1:100, 1:200, or 1:400
   
4. Nuclear pore 4 µg/mL final concentration
7. Incubate with the diluted secondary antibodies for 1 hour in dark,  followed by three washes of 10 minutes with the blocking buffer.
   
8. Aspirate the staining solution. Wash with 1X PBS three times for 10 minutes each.
   
9. Fix cells again with 4% Paraformaldehyde for 10 minutes.
   
10. Wash cells 3 times with 1X PBS for 10 minutes each.
    
11. Apply the dSTORM buffer for imaging or store it in 1X PBS for imaging later.

Discussions

1. We recommend the following sample configurations to better understand your data (if you see a lot of background in Sample 1 and Sample 2 then better to rectify this background before moving to Sample 3):
   

   Sample 1 (dSTROM background from the unstained sample)	Sample 2 (only secondary antibody)	Sample 3 (primary + secondary antibody)
   Cells Fixed with Paraformaldehyde	Fix cells and follow the staining protocol with secondary antibody and no primary	Fix cells and follow the staining protocol with primary and  secondary antibody
   dSTORM imaging to know the background signal with 488, 561, and 640 lasers	dSTORM imaging to know the non-specific signal for 488, 561, and 640 lasers	dSTORM imaging with required lasers

1. We recommend labeling the primary antibodies directly with fluorophores to increase the localization accuracy.