Protocol for Flow cytometry

Flow cytometry

Prepare Cells (day 1)

1. Subculture Raw 264.7 into 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 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 pipet the media up and down several times to get a homogenous cell suspension 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.
7. Label the 12-well plates according to experimental condition.
8. 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.  
9. Incubate the well-plate at 37⁰ C, 5% CO₂ overnight.

Stain Bacteria and prepare inoculum (day 2)

10. In hood add 5ml of a log-phase bacteria culture into 45 ml of PBS in 50 ml conical flask. Log phase culture is used because the least dead cells is present this phase. Make sure that each tube is exactly 50 mL in order to balance the tubes for the centrifugation.
11. Centrifuge at 2500 rpm for 8 minutes and remove the supernatant. Thus the media used for growth will be washed away.
12. 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.  
13. Add 5μl of Alexa 488 stock solution in microcentrifuge tube and mix by pipetting. (Turn of the light of Biosafety cabinet to avoid photobleaching)
14. Wrap the microcentrifuge tube with Aluminum foil. Then, incubate the flask at room temperature for 60 minute on a shaker.
15. Then, pellet bacteria 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.
16. Pipette 5 ml complete DMEM in a 7ml Dounce homogenizer
17. 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. 
18. Measure OD₆₀₀ of the 1:10 (900 µL medium + 100 µL suspension) dilution of homogenized culture. First, put 900 µL medium in a cuvet and then measure OD₆₀₀. 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.
19. Prepare bacterial inoculum in DMEM at the concentration of 3x10⁶ bacteria/ml (MOI=1:10) using this formula:

(3x10⁶) x (total ml needed for infection / (OD₆₀₀ x 10⁹) = mL homogenate needed.

Phagocytosis

20. Transfer Raw cells from incubator to Biosafety cabinet. 
21. Remove media from Raw cells and add 1ml of 3x10⁶ bacteria/ml inoculum in a well and DMEM medium without bacteria (Control) into anther well. 
22. Spin at 1200 rpm for 5 min at room temperature to settle bacteria on cells. It will help Raw cell to internalize bacteria.
23. Incubate the plate for 30 minutes at 37⁰ C, 5% CO₂ in incubator. Within this time bacteria will be internalized by Raw cells.
24. After incubation, terminate phagocytosis by adding 1 ml of ml of ice-cold PBS in each tube
25. Quickly wash each well two times with 1 ml of ice-cold PBS to remove free bacteria which were not internalized by Raw cells.
26. Then add 1 ml of ice-cold PBS in each well and detach the adherent RAW264.7 cells by GENTLY scrapping well surface using a cell scrapper and transfer the cell suspension from a well to a polypropylene tube. 
27. Centrifuge at 1200 rpm for 5 minute and then remove supernatant
28. Add 500 uL ice cold PBS in each polypropylene tube
29. Keep on ice until flow analysis. 

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  (1^st 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.

29. Transfer Raw cells from incubator to Biosafety cabinet. 
30. Remove media from Raw cells and add 1ml of 3x10⁶ Mtb/ml inoculum into each well of Raw 264.7 cells. 
31. Spin at 1200 rmp for 5 min at room temperature to settle Mtb on cells. It will help Raw cell to internalize Mtb.
32. 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)
33. 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.  
34. 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.
35. 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.
36. 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.
37. Incubate for 2 hours at 37⁰ C, 5% CO₂ in incubator.
38. 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)
39. Remove the fixative and add 1 ml of PBS in each well.
40. Carry the well plates to BSL-2. In BSL-2 wash the cells 3 times (each time for 5 minute)
41. 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.
42. 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.
43. Keep the slides in a dark chamber for overnight. Then preserve them in a slide box at -20⁰C.
44. Collect the images (10 image for each cover slip) using confocal microscope.
45. Analyze the images and calculate the percentage of Lysotracker red and Mtb (green) colocalization.
46. Repeat the entire assay two more times.

Protocol for Western Blot

Sample preparation for Immunobloting

1.     Seed cells to 6 well-plate (1x10⁶ cells per well) and incubate overnight at 37⁰C, 5% CO2

2.     Treat cells for …. Minute / Hours according to experimental condition

3.     Wash the cell twice with 1x PBS (1mL)

4.     Add 1mL PBS and pipette up and down vigorously to detach the cells from floor of the plate

5.     Collect the cells in a 1.5 mL eppendorff tube preserved in an ice box

6.     Spin at 10,000 RPM for 1 min and then remove supernatant

7.     Add 150 µl lysis Buffer and mix well by pipetting up and down.

8.     Keep eppendorff tube in ice box

9.     Sonicate sample in each eppendorff  tube for 30  time (one second pulse  with 50% duty or power level 3.0)

10.     keep in -20⁰c for next day work

Protocol for SDS-PAGE and immunoblotting

Day-1

1.     Heat sample for 5 minute at 100⁰c

2.     Centrifuge at 10,000 rpm for 5 minutes at 4⁰C

3.     Set mini gel apparatus for SDS-PAGE

Ø Clean glass plates and alumina plates with ethanol.

Ø Assemble the glass plates and spacers.

Ø Mark a line 5.5cm from the bottom of a long side of the glass

Ø Attach the glass plate assembly to the holder.

Ø To prevent leak use parafilm at the bottom

4.     Make 5 ml of separating gel (Tris-HCl ph8.8) depending on MW of protein (15% Acrylamyde is for LC3, 10% for Beclin1)

5.     Add the separating gel with a pasteur pipette.

6.     Gentlely add 3A water to the top of this until the level reaches the top of the plates.

7.     The separating gel should polymerize in 30 minutes.

8.     Make the stacking gel reagent (Tris-HCl ph6.8)

9.     Pour off 3A water. Dry the watery interplate surface with a piece of Whatmann paper.

10.            Insert the comb straight on down, then pour the stacking gel on top of the polymerized separating gel to fully seal the comb. Remove any bubbles from underneath the comb, if possible, by moving the comb gently from side to side so the bubbles get into the space in between and float up.

11.            The stacking gel should polymerize in 20 to 30 minutes.

12.            Add some sds page buffer in SDS-PAGE apparatus

13.            Remove the glass-gel set from holder and set it in SDS-PAGE apparatus

14.            Remove the comb from glass-gel set

15.            Fill the space with SDS-PAGE buffer

16.            Load 2µl marker into first well of the SDS-PAGE gel

17.            Load 10 µl of sample into the well of the SDS-PAGE gel

18.            Run gel for at  25 mA, run until the blue marker goes to the end of the gel (about 1 hr)

19.            While the gel is running, prepare the transfer buffer, and cut out a piece of nitrocellulose membrane (9x6 cm) and four pieces of Whatmann filter paper for the transfer. Biorad tank takes about 2.5 liters of transfer buffer.

20.            Cut out the Whatmann filter papers so they each are slightly larger than the gel in each dimension--about 1/4 to 1/2 cm larger. Cut out the nitrocellose so that it is slightly smaller than the Whatmann filter papers but still covers the entire surface of the gel.

21.            Prewet the nitrocellulose membrane in TOWBIN’s buffer for at least five minutes before blotting step.

22.            Start cooling machine and put magnetic stirrer in transfer tank to cool the it at 4⁰C

23.            Remove the plates from the apparatus. Remove the spacers and pry off one of the plates by bending the spacer. The gel should now be stuck to one of the plates.

24.            Carefully float the gel off the plate into a tupperware container containing transfer buffer. .

25.            When the gel has equilibrated, construct a "sandwich" for the transfer as follows:

a) Place some transfer buffer in a large glass or tupperware container and place the plastic frame inside so the white side is lying in the buffer and the black side is folded out.

b) Place a sponge on top of the white side, then stack two sheets of the Whatmann paper you cut out on top. These things should be submerged in the buffer so that they become completely wet.

c) Place the nitrocellulose filter, labeled appropriately, on top, then carefully position the gel on top of this.

d) Add two more pieces of Whatmann filter paper, then a second sponge. Then press everything down into the buffer and squeeze out any air bubbles that might be trapped in the sandwich using a pipette as a roller.

e) Close the sandwich by folding the black side over onto the white and locking it in place. Use rubber band to lock it. Then immerse the sandwich in transfer tank to which transfer buffer has been added. Since the proteins will migrate toward the positive end, make sure that the nitrocellulose side is closest to the positive electrode. Make sure that the lid is put on the right side. That is, the white side (membrane) of the sandwich should be facing the red side and the black side (gel) facing the black side

26.            Electrotransfer to nitrocellulose membrane at 90 V for 1.5 hr. Make sure to have the cooling unit on at 4 C so that cold water can circulate through the tank while the transfer is taking place. Otherwise, if the buffer gets hot, bubbles of air may come out of solution and become trapped in the sandwich.

1.     After transfer, stain the membrane with 1x ponceau S for 5 sec. and destain with ddH₂0

2.     Wash membrane with 1x PBS for 5 min

3.     Block the membrane for 1h with 5ml of 5% blocking reagent (Roche) in PBS on the shaker

4.     Incubate with primary antibody (antibody in 5ml of 2.5% blocking reagent in PBS) for overnight, Incubate at 4⁰C on the shaker

Day 2

5.     Wash membrane 3 times (10 min. each time) with 10 ml of 0.1% Tween 20 in PBS at RT

6.     Incubate with secondary antibody (antibody in 5ml of 2.5% blocking reagent in PBS) for 1hr at RT

7.     Wash membrane 4 times (15 min. each time) with 10 ml of 0.1% Tween 20 in PBS at RT

8.     Cut and set plastic bag in X-ray box during last wash

9.     Detect protein with Enhanced Chemi-Luminescence (ECL). Incubate membrane in ECL solution (prepared fresh by mixing 20μL of ECL 1 in 2mL of ECL 2) for 2 min. Drain the ECL and wrap the membrane in plastic bag and expose to film.