Blueprint_macrophage

Blueprint Monocyte derived Macrophage culture SOP

Plastic ware
• 0.2uM sterile filter and 20ml sterile syringe • Stem cell grey magnet for 15ml large round bottom tube • 6-well SUSPENSION culture plate Gernier Bio-one 657185
Reagents
• 1x PBS no calcium chloride or magnesium chloride sigma D8537 • HSA 20% solution Gemini Products 800-121 • Sodium Citrate 13mM stock in water (powder Sigma S4641) • Buffer 1: DPBS sigma, 0.2% HSA, 13mM Sodium Citrate • Buffer 4: DPBS sigma, 0.2% HSA, 2mM EDTA • Ficoll-paque premium GE healthcare 17-5224-03 • SIP solution = 45ml percoll stock and 5ml 10X PBS. Once made mix VERY well • Stem cell monocyte negative isolation kit cat number 19059 • Invitrogen Macrophage serum free media 12065-074 • Macrophage colony stimulating factor GIBCO • LPS sigma serotype 055:B5 sigma L5418 1mg/ml • Rosiglitazone Sigma R2408, 10mM stock in 100% DMSO Antibodies for monocyte purity check:
• CD14-FITC BD 561271 (20ul/test), CD42b-PE IBGRL 9448PE NHS (1ul/test), CD64percpcy5.5 BD 561194 (5ul/test), CD45-PEcy7 MHCD4512 (5ul/test) (use the FC500 for analysis) • Small compensation beads BD Compbeads 51-90-9001229 Antibodies for Macrophage purity check:
Panel 1: CCR7-FITC, CD25-PE, CD14-PEcy5.5, CD40-PEcy7
Panel 2: CD206-PE, CD36-FITC, CD45-PEcy5.5
Panel 3: after running the unstained cell sample and panel 1 and 2 stain with PI (0.5ul of sigma stock)
Large compbeads: BD compbeads PLUS 560497 CCR7-FITC BD 561271 (5ul/test) CD25-PE MACS 120-001-311 (10ul/test) CD14-PEcy5.5 Invitrogen MHCD1418 (5ul/test) CD40-PEcy7 BD 561215 (5ul/test) CD206-PE BD 555954 (20ul/test) CD36-FITC Southern Biotech 9605-02 (10ul/test) CD45-PEcy5.5 Invitrogen MHCD4518 (5ul/test) Materials and Methods
Work in a class II tissue culture hood as the monocyte isolation needs to be a sterile process This protocol should be used for a whole unit of blood to obtained on average between 20-60 million Day 1 = the day you start the monocyte extraction 1) Aliquot 12.5ml of ficoll into 32 fifty ml tubes. 2) Allow the unit of blood to cool to room temperature, using scissors cut one of the tubes and gently squeeze the blood into the sterile 1L bottle. 3) If processing PRP for platelet isolation take 50ml of undiluted blood in two falcon tubes and spin 3) While the PRP is spinning dilute the blood 1:1 with buffer 1 4) Layer 30ml of the diluted blood onto 12.5ml of ficoll in 50ml tubes 5) Once PRP has finished spinning take two 15ml aliquots, then add the remaining blood back into the 1L bottle and wash out each 50ml tube with 50ml buffer 1, layer the remaining blood and spin at 800g for 15 minutes at room temperature up 4 down0. 6) Using a 25ml stripette carefully harvest the PBMC layer, harvest the layer from two 50ml blood tubes into one new 50ml tube. Top up the tube to 50ml with buffer 4. 7) Spin tubes at 600g for 6 minutes up9 down9 8) Carefully pour off the supernatant, re-suspend the cell pellets in a small volume of buffer 4 and 9) Remove a 10ul cell sample and dilute in 90ul of trypan blue, count using a disposable haemocytometer. While counting put the cells on to spin at 600G up 9 down 9. 10) Carefully pipette off the supernatant and re-suspend the cell pellet to 5e6 cell/ml in room 11) Make up the ‘2nd percoll solution’ = 10ml SIP and 8.64ml 1x DPBS. MIX VERY WELL! (enough for 4 12)Aliquot 4ml of SIP solution into 15ml falcon tubes and carefully layer 5ml of cell suspension on 13) Spin at 800G up 4 down 0 at RT for 15 minutes. 14) Using a sterile pastette harvest the top monocyte rich layer and the PBS layer down to 2ml. Harvest the monocyte rich layer from two 15ml tubes into one 50ml tube. Discard the bottom layer. 15) Top up the volume to 50ml with room temperature buffer 4 and spin at 600G up9 down 9 for 6 16) Pour off the supernatant and re-suspend the pellets in a small volume of buffer 4. Combine the cell suspension into one tube and make the volume up to 50ml with buffer 4. 17) Take a 10ul cell sample and dilute with 90ul trypan blue and count the cells. While counting spin the sample at 600G up 9 down 9 for 6 minutes at 4C. 18) Carefully pour off the supernatant and re-suspend the cell pellet to 50e6 cells/ml in 4C filtered 19) Follow the protocol described in the stem cell monocyte negative selection isolation kit cat#19059. Except two, 2.5 minute incubations in the magnet are carried out.
20) Add 50ul antibody cocktail per 1ml cells and incubate at 2-8C for 10 minutes 21) Add 50ul of magnetic particles per 1ml cells and incubate at 2-8C for 5 minutes 22) Top up the volume to either 5ml of 10ml depending on the initial starting cell number ,with 23) Place 5 or 10ml of cell suspension into a round bottom 15 ml tube and place the tube in the magnet without the lid for 2.5 minutes at RT. Then carefully pour off the supernatant into a fresh 15ml round bottom tube. Place the tube back into the magnet for a further 2.5 minutes and then pour off the supernatant into a fresh 50ml tube. 24) Remove a10ul cell sample for counting 25) Take 2x ~25,000 cell volume for flow cytometry analysis and 2x10,000 cell volume for cytospin 26) Spin down the remaining cells at 600G up 9 down 9 at 4C. 27) Pour off the supernatant and resuspend the cell pellet to 1e6 cells/ml in room temperature 28) Add the relevant treatment depending on the cell type you want to generate. 29) Plate 2ml of cell suspension into each well of a 6-well SUSPENSION culture plate. Swirl plates gently to distribute the cells and incubate at 37C 5% C02 = Day 1 of culture. M0 resting macrophages- stimulated with 50ng/ml M-csf for 8 days M-csf= 10ug vial from Gibco. Add 100ul of sterile water to reconstitute, then to this add 900ul of PBS 0.2% HSA. Aliquot into single use aliquots and store at -80C. Avoid multiple freeze thaw cycles. Add 10ul of this stock per 2ml of media (i.e. 10ul to each well of the 6-well plate). The media should be M1- inflammatory macrophages- stimulated with m-csf at 50ng/ml for 7 days, then LPS alone at 100ng/ml for the last 18 hours of the culture, cells harvested at day 8. M2- Anti-inflammatory macrophages- cells cultured in the presence of 15ng/ml IL-13 AND 0.1uM Rosiglitazone. Rosiglitazone must be added at the start of the culture as it is only effective on monocytes. The media should be changed at day 3 or day 4. The Rosiglitazone is made up in DMSO to a 10mM stock and stored at 4C. Use the stock solution IL-13- this is only stable for 3 months once reconstituted and should be stored at -80C. Reconstitute in sterile water 0.2% HSA. Store in single use aliquots Monocyte antibody panel (to assess purity of initial starting culture)
CD14-FITC (BD) 20ul per test)
CD42b-PE NHS (1ul per test)
CD64-percpcy5.5 (5ul per test)
CD45-PEcy7 (5ul per test)
Stain ~ 25,000 cells for 10 minutes at 4C, don’t fix but just add 600ul of 0.2uM filtered buffer 4 and
read on the FC500 using the programme MDM culture.
NB- purity of monocytes ranges form 20-70%, main contaminant is platelets.
Media changing tips:
Make sure that the media is gently swirled over the surface of the wells before aspirating the old
media, this ensures that any non-adherent cells and platelets are removed from the culture.
As the culture progresses the cells should start to appear more rounded and enlarged. The M0/M1
cultures should form at least a 50% confluent monolayer. The cells in an M2 culture maybe much
sparser.
Compound dilutions
M-csf- once reconstituted it can be used undiluted 10ul per well
LPS- diluted to 100ng/ml final concentration in the plate. Dilute 10ul of LPS into 90ul of media. Then
add 1ul of this diluted stock per 1ml of media required.
IL-13- reconstitute to 100ug/ml. Diluted stock 1 in 100 e.g. 2ul into 198ul of media. Add 30ul of this
diluted stock to 2mls of media. (i.e. 30ul per 6-well) final 15ng/ml.
RSG stock 10mM in 100% DMSO. Dilute 1 in 100 in media and then add 2ul of this to 2mls of media
(i.e. 2ul per well). final 0.1uM.
Harvesting the cells:
The cells are harvested at day 8- exactly one week after the culture was set up i.e. if you started the
culture on a Tuesday harvest the cells the following Tuesday.
Before harvesting the cells take light microscopy pictures of the plates so that the cell morphology
can be assessed.
The cells are removed from the plate by gentle scraping except for the RNA and BS samples where the cells can be lysed directly in the plate. RNA harvest- Remove the spent media from the well and discard. Rinse the well with PBS to remove any non-adherent cells and discard the PBS. Add 700ul trizol reagent directly to the empty well and pipette over the surface to insure that the cells lyse. Leave the trizol reagent in the well for 5 minutes to insure complete cell lysis. Then transfer the lysate into a 1.5ml RNase free tube and BS harvest- Remove the spent media from the well and discard. Rinse the well with PBS and discard. If not proceeding with the DNA extraction directly, then freeze the empty well at -80C for DNA extraction at a later date. DNA lysis buffer can be added to the plate and incubated for several Chromatin harvest- the cells are fixed and quenched in the plates before being scraped from the Remove the spent media from the wells and discard. Wash the wells with PBS and discard. Add 1ml of 1% formaldehyde in PBS to each well and incubate at room temperature for 10 minutes. Add 55ul of 2.5M glycine to each well, mix gently and incubate at room temperature for 5 minutes. Place the plate of cells on ice and use a cell scraper to scrape the cells from the plate. Transfer the cell suspension to a 15ml falcon tube and spin at 600G for 6 minutes up 9 down 9. Pour off the supernatant and re-suspend the cell pellet in 10ml 4°C PBS. Spin again at 600G up 9 down 9 for 6 minutes. Pour off the supernatant and add 6ml 4°C PBS. Store sample at 4°C until shipping. Dnase 1- the cells are scraped from the plates using a cell scraper and the DNAse 1 protocol is followed. NB- the cell pellet may break up during washing; just carry on as DNA is still recovered. Analyse one well of a 6-well plate for flow cytometry analysis. 1) Remove the spent media from the well and discard. 2) Gently rinse the surface of the well with 1ml PBS to remove any non-adherent cells- discard the 3) Add 1ml of sterile 0.2uM filtered buffer 4 and gently scrape the cells from the well, leaving a small portion un-scraped, for staining with rapid Romanowsky stain. Transfer the cell suspension to a 15ml tube. Keep the cell plate (stained later). 4) Spin the cells down at 300G up 9 down 9 for 6 minutes 5) Remove the supernatant and resuspend the cell pellet in 100ul of 0.2uM filtered buffer 4. 6) Divide the cells suspension equally between three 5ml BD FACS tubes 7) Add the antibodies to the tubes to create panel 1 and panel 2. 8) Stain Compbeads PLUS with a single stain of antibody 9) Place tubes in Fridge for 10 minutes to allow cells to stain 10) Add 600ul of 0.2uM filtered buffer 4 to each cell tube and the beads 11) Read the tubes on the FC500 using the protocol FB macrophage_plots (NB- as the beads are large there is no need to change the discriminator between cells and beads). Read the PI stain
LAST. There will be a large proportion of dead cells in the PI stain
12) Transfer the LMD files to the Haematology network 13) Clean the flow cytometer with TWO cleaning panels: The small proportion of cells not scraped from the flow cytometry plate are stained with rapid Romanowsky stain. Ensure that all media/PBS has been removed from the well. Allow the well to dry for at least several hours. Add 700ul of solution A to well and incubate for 40 seconds. Remove solution A and discard. Add 700ul of solution B to well and incubate for 40 seconds. Remove solution B from well and discard. Add 700ul of solution C to well and incubate for 40 seconds. Remove solution C from well and rinse the well several times with tap water until the water runs clear. Allow plate to dry and then take pictures using the microscope in Cedric’s lab. (Amanda, Kate Downes and Fizzah know how it works).

Source: http://www.blueprint-epigenome.eu/UserFiles/file/Protocols/UCAM_BluePrint_Macrophage.pdf