LysoTracker colocalization analysis to monitor the acidification of Mtb phagosome



  1. Subculture Raw 264.7 into two 75 cm2 tissue culture flask (1:10) and incubate for 2 day.
  2. One day before the experiment, remove old media and add 5 ml complete DMEM into each flask.
  3. Scraps off the cells into 5 ml complete DMEM and pipette up and down several times to get single cell suspension.
  4. Then merge the media of two flasks into one flask and pipet the media up and down several times to get a homogenous cell suspension so that it can be used as a single batch and to make cells to be single cells as clumping cells will affect the assays.
  5. Determine the cell density using hemocytometer. If possible stain the cell using Tryphan blue before counting with hemocytometer so that the live and dead cell can be distinguished.  Since, cells are very selective in the compounds that pass through the membrane, in a viable cell trypan blue is not absorbed; however, it traverses the membrane in a dead cell. Hence, dead cells are shown as a distinctive blue colour under a microscope.
  6. Dilute cells to 3X105 cells/ml in complete DMEM (total 25 ml for working with 6 strains of Mtb). Cells suspension is diluted so that they can grow without any nutritional stress in well plates. Because, any stress can induce autophagy. Moreover, the known number of cells per ml will be needed to calculate the Multiplicity Of Infection (MOI) for infecting with Mtb.
  7. Label the 12-well plates according to strains of Mtb and experimental condition. Use two wells for each experimental condition.
  8. Take some Methanol in a Petridis and dip the cover slip (preserved in 70% ethanol) in methanol.
  9. Then, put one cover slip in each well (Place it in vertical position so that methanol can evaporate rapidly. Methanol is toxic to cells).
  10. Observe the cover slips, if all cover slips are dry then shake the well plate in such a way that all cover slips placed horizontally in bottom of well.
  11. Dispense 1ml diluted cell suspension into each well of a 12-well plate using 10 mL pipettes.  Avoid any air bubble in tip of the pipette during pipetting.  
  12. Incubate the well-plate at 370 C, 5% CO2 overnight. Here, 370 C is used because mammalian cells require this temperature to grow. 5% CO2 is used in order to maintain the pH to be at 7.4
  13. In the date of experiment take six 50 ml conical flask and label them according strain of Mtb. Pipette 45 ml PBS in each flask. 
  14. Carry the six 50 ml conical flasks containing PBS and two 12 well plates (incubated overnight), Fixative, Lysotracker red and Alexa 488 stock solution (32 μl) into BSL-3. Wrap the Lysotracker red and Alexa 488 with Aluminum foil to avoid photobleaching by light.

In BSL-3
  1. Keep the plates as soon as possible at 370 C, 5% CO2 in incubator in BSL-3
  2. Turn on OD600 machine and Centrifuge machine to be ready them.
  3. Put DMEM and EBSS into water bath to warm them and put Lysotracker red and Alexa 488 in refrigerator at -200C.
  4. Prepare safety hood for your immediate work. {Clean the floor of hood with 8% ADBAC, Put a waste container, a waste bottle for liquid waste, a rack, 6 conical flasks containing PBS, pipette and pipette gun. For first time entry decontaminate everything with 70% ethanol, But, in case of all out  (1st out also) from hood everything  must be decontaminated with 8% ADBAC}
  5. In hood add 5ml of a log-phase Mtb culture into each conical flask according to label. Log phase culture is used because the least dead cells is present this phase. PBS is used here to wash out Tween-80 which will affect the infection. Make sure that each tube is exactly 50 mL in order to balance the tubes for the centrifugation.
  6. Centrifuge at 2500 rpm for 8 minutes and remove the supernatant. Thus the media used for growth will be washed away.
  7. Resuspend the pellet in 0.5 ml of 1X PBS by pipetting up down and transfer the cell suspension into a 1.5-ml microcentrifuge tube.  
  8. Add 5μl of Alexa 488 stock solution in each microcentrifuge tube and mix by pipetting. (Turn of the light of Biosafety cabinet to avoid photobleaching)
  9. Put six tubes in a 50 ml conical flask and wrap it with Aluminum foil. Then, incubate the flask at room temperature for 60 minute on a shaker.
  10. Then, pellet Mtb at 8000 rpm for 3 minutes at room temperature. Remove the supernatant and wash twice with1 ml of 1X PBS. Observe the pellet if any green trace is found then wash again.
  11. Pipette 5 ml complete DMEM in each 7ml Dounce homogenizer
  12. Add 1 ml complete DMEM in each microcentrifuge tube and mix by pipetting. Transfer the suspension to 7ml Dounce homogenizer and homogenize 35 times to generate single cell suspension.  
  13. Measure OD600 of the 1:10 (900 µL medium + 100 µL suspension) dilution of homogenized culture. First, put 900 µL medium in each cuvet and then measure OD600 one by one. Each time put 900 µL medium in Machine and set blank for this and then add 100 µL suspension in media, pipet up and down to mix, lid the cuvet with parafilm and finally measure OD600 and keep the record.
  14. Prepare Mycobacterial inoculum in DMEM at the concentration of 3x106 Mtb/ml (MOI=10) using this formula:
(3x106) x (total ml needed for infection / (OD600 x 109) = mL homogenate needed.
  1. Transfer Raw cells from incubator to Biosafety cabinet. 
  2. Remove media from Raw cells and add 1ml of 3x106 Mtb/ml inoculum into each well of Raw 264.7 cells. 
  3. Spin at 1200 rmp for 5 min at room temperature to settle Mtb on cells. It will help Raw cell to internalize Mtb.
  4. Incubate the plate for 15 minutes at 370 C, 5% CO2 in incubator. Within this time Mtb will be internalized by Raw cells. (Pulse period)
  5. After incubation, quickly wash each well three times with 1 ml complete DMEM to remove free Mtb which were not internalized by Raw cells. Any delay may induce autophagy in cells because of stress.  
  6. Add 1 ml DMEM in each well and incubate for 1 hours at 370 C, 5% CO2 in incubator. Within this time no new bacteria will be internalized by Raw cells, as a result all bacteria will be in phagosomes after 1 hours.
  7. Prepare 12 ml Complete DMEM and 24 ml EBSS, both containing 0.25 μM Lysotracker. (Add 3 µL of Lysotracker in 12 ml DMEM and 6 µL of Lysotracker in 24 ml EBSS). Turn of the light of Biosafety cabinet to avoid photobleaching.
  8. After the incubation, wash cells 3 times with 1 mL PBS to remove the media which will affect the starvation. Then add 2 ml of EBSS containing Lysotracker in each well labeled as starvation and add 1 ml  of DMEM containing Lysotracker in each well labeled as full.
  9. Incubate for 2 hours at 370 C, 5% CO2 in incubator.
  10. After incubation, fix the cells with 1 ml of 4% (fixative) in each well for 10 minutes at room temperature. (Be aware of fixative because any kind of direct contact of  paraformaldehyde is harmful to your body)
  11. Remove the fixative and add 1 ml of PBS in each well.
  12. Carry the well plates to BSL-2. In BSL-2 wash the cells 3 times (each time for 5 minute)
  13. Prepare microscopic slides (One slide for two cover slips, label the slides according to experimental condition). Put a drop of ProLong Gold Anti-fade Reagent for each cover slip on microscopic slide.
  14. Place the cover slips on the drops of Anti-fade Reagent on slide in such a way that the cells will face the drop of Anti-fade Reagent.
  15. Keep the slides in a dark chamber for overnight. Then preserve them in a slide box at -200C.
  16. Collect the images (10 image for each cover slip) using confocal microscope.
  17. Analyze the images and calculate the percentage of Lysotracker red and Mtb (green) colocalization.
  18. Repeat the entire assay two more times.