TITLE: Lysosome analysis with confocal microscopy
AUTHORS: Sjors Maassen

[DESCRIPTION]
This protocol provides an overview of lysosomal analysis using confocal microscopy and Fiji

[STEPS]
SECTION: Sterilize glass cover slips
1. Put the glass coverslips in a 24-well plate, submerge the coverslips in 70% ethanol, and then expose them to UV light in a tissue culture hood for between 20 and 30 minutes.

SECTION: Sterilize glass cover slips
2. Remove the ethanol, wash the wells containing coverslips three times with sterile PBS, and remove the PBS. Add 450 µL of cell-culture media to the well. 
SECTION: Seed, treat and fixate cells
3. Make a suspension of your cells (1.200.000 cells/mL) and seed these at 60.000 cells per well - add 50 µL. 

Note: This cell number is an empirically observed amount with minimal contact between relatively large cells (Fibroblast/Macrophages). However, optimal seeding densities might vary per cell type.   

Alternatively, you could use confluence and the area of a cell-culture dish and aim for 50% confluency. This might require adding extra volume to make the total volume 500 µL. 
Example: you have a 1mL suspension of a 100% confluency T75-flask. 
The area of a 24-well plate well =  ∼1.9cm2
(1.9/75)*0.5 = 0.0126 mL = 12.6 µL cell suspension 

Incubate the cells overnight at 5%CO2 and 37 ºC, to let them adhere. 
SECTION: Seed, treat and fixate cells
4. The next day, refresh the media to remove any dead cells and treat the attached cells. Wash the cells with 500 µL 1X sterile PBS 3 times and fixate the cells 15 min at 4C with 2-4% PFA 
SECTION: Seed, treat and fixate cells
5. After fixation, wash the wells 4 times with 500 µL 1X sterile PBS. Add 500 µL 1X sterile PBS to the well and store them at 4ºC until use.  
SECTION: Stain glass coverslips
6. Remove PBS and add Abs-staining buffer supplemented with 0.1 % Saponin and incubate at room temperature for 30 min.   
SECTION: Stain glass coverslips
7. Dilute your LAMP1 (D2D11) mAb rabbit (1:200) in Abs-staining buffer supplemented with 0.1 % Saponin for 30 µL per sample. put on ice
SECTION: Stain glass coverslips
8. Put parafilm in a petridish. Add 30 µL droplets of Abs-staining buffer containing the primary Ab 
SECTION: Stain glass coverslips
9. Invert the glass coverslips onto the droplets, facing cells down into the droplet
 

SECTION: Stain glass coverslips
10. Add a damp lint-free tissue to the Petri dish and store the sample overnight at 4ºC.

 

SECTION: Stain glass coverslips
11. The next day, invert the glass coverslips back into a 24-well plate and wash two times with cold 1X PBS
SECTION: Stain glass coverslips
12. Repeat steps 7, 8, and 9, but now instead of the primary Ab, use the secondary against the primary Ab and other markers like DAPI (5 µg/mL) or phalloidin. Make sure to incubate the sample in the dark at room temperature for 45-60 min   
SECTION: Stain glass coverslips
13. Invert the glass coverslips back into a 24-well plate and wash three times with cold 1X PBS
SECTION: Stain glass coverslips
14. Add 5µl of Trolox media to a microscope glass slide. Take out the glass coverslip from the 24-well plate and briefly hold the side of the coverslip against a dry lint-free tissue to absorb as much liquid as possible before inverting the coverslip onto the Trolox media. 
SECTION: Stain glass coverslips
15. Let the sample cure for an hour or two at room temperature in the dark and seal the sample with nail polish once the sample has dried to some extent and is not moving when touched gently. After sealing the sample, store it away at room temperature in the dark until image acquisition.  

 

SECTION: Stain glass coverslips
16. Before image acquisition, you should see a lot of dried salts when looking at your sample. Make sure to add some milliQ to the top of your sample and gently add a fibre-free cloth onto the water droplets to remove the dissolved salts. repeat this step a few times and wipe the samples dry afterwards, now we can go to the microscope for imaging 
SECTION: Imaging
17. Since not every lab has the same microscope, I feel it does not make sense to specify a microscope here. However, I recommend performing confocal microscopy with high magnification (63X, NA 1.4) to look at lysosomes, as these organelles are small and require some resolution to make sure lysosomes are distinguished enough for identification.

Overall I try to have a sufficient amount of images for analysis (n=10-20 per condition times 3 technical repeats)

When an intensity quantification is desired to indicate protein levels or other stainings, I recommend resorting to flow cytometry or western blot techniques. If you still want to use microscopy as a quantification tool, at least make sure not to overexpose your images and saturate pixels. 
SECTION: Image analysis
18. Depending on the question that you might require different image analysis strategies are suitable for looking at lysosomes. In this section, I will explain the basal procedure that I use to analyse lysosomes.  This protocol assumes that you stored your samples as a '.lif' file.

This tutorial is for users that run: 
- Windows 11 version 23H2 (OS Build 22631.3007) 
- ImageJ 1.54f

SECTION: Image analysis
19. Open Fiji
SECTION: Image analysis
20. Drag your '.lif' file into the program and select all samples of interest
SECTION: Image analysis
21. Run this code in the ImageJ2 Script editor (ImageJ Macro), to get all images into a TIFF:

 

After you run this code, consider putting all TIFFs into a folder per condition 
SECTION: Image analysis
22. Now we want to get all images from a folder/condition, create a LAMP1 lysosome mask and analyse the structures: 




 


Now you should have all your data stored separately in a CSV, a mask for reference, and a summary of all CSV's in your folder for further analysis
SECTION: Image analysis
23.