A Cellular Assay for the Identification of CXCR4-Interacting Agents

NOTE: All steps described under sections 1 and 2 are carried out under sterile conditions in a laminar flow cabinet.

1. Maintenance of U87.CD4.hCXCR4 Cells

1. Grow the cells in T75 culture flasks at 37°C and 5% CO₂ in a humidified incubator.
   NOTE: The in vitro cell line used in this protocol is a U87 glioblastoma cell line stably expressing cluster of differentiation 4 (CD4) and human CXC chemokine receptor 4 (CXCR4) and has been previously described. Cell surface expression of CD4 and CXCR4 is continuously monitored by flow cytometry and expression levels remain constant over time (~ 100% CD4⁺ and ~ 100% CXCR4⁺ cells). A detailed description of the generation of the used cell line and of the flow cytometry procedure to investigate receptor expression levels is not within the scope of this protocol.
   1. Subculture cells at 80-85% confluency. Allow all reagents to reach room temperature (RT) before cell culturing.
   2. Remove the conditioned culture medium from the cells and wash the cell monolayer once with 5 mL phosphate-buffered saline (PBS).
   3. Add 3 mL of 0.25% trypsin-ethylenediamine tetraacetic acid (trypsin-EDTA) and distribute it evenly over the cell monolayer. Then remove the excess trypsin-EDTA and incubate up to 5 min at 37 °C until cells start to detach.
   4. Add 10 mL of fresh complete growth medium (Dulbecco's Modified Eagle's Medium (DMEM) + 10% fetal bovine serum (FBS) + 0.01 M 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) + appropriate selection agents). Resuspend the cells by gently pipetting them up and down. Transfer the cell suspension to a sterile 50 mL tube.
   5. Count the number of viable cells using a method of choice. In the case of U87 glioblastoma cells, viability generally reaches ~ 100%.
      NOTE: Cell numbers can be determined in several ways. We routinely use an automated cell analysis system based on trypan blue staining (see Table of Materials) according to its standard handling procedures, but other manners should work equally well.
   6. Add 2 x 10⁶ or 3 x 10⁶ (viable) cells to a final volume of 25 mL fresh growth medium in a T75 culture flask and incubate at 37 °C and 5% carbon dioxide (CO₂).
      NOTE: If 3 x 10⁶ cells are used the cells will reach 80-90% confluency after 2 days. If 2 x 10⁶ cells are used, they will reach the same confluency after 3 days. The growth rate of the cells should first be determined empirically if other cell lines are used.

2. Seeding of the Cells for the Ca²⁺ Mobilization Assay

1. On the day of cell passaging (day 0), coat black-walled polystyrene 96-well plates with clear bottoms (see Table of Materials) with a 0.1% gelatin solution to facilitate cell attachment. 
   NOTE: Coating of the 96-well plates with gelatin might be omitted if pre-coated plates, which are commercially available (see Table of Materials), are used. It is recommended to evaluate the use of pre-coated plates instead of manual coating for each cell line under investigation before continuing with the protocol.
   1. Prepare gelatin solution by adding 1 g of gelatin to 100 mL PBS to obtain a 1% solution. Dilute by 10x with PBS before further use. To improve the solubility of gelatin, heat the solution to 37 °C.
   2. Add 100 µL of 0.1% gelatin solution per well of the 96-well plate using a multichannel pipette. Incubate for 2 h at RT.
2. In the meantime, prepare the cells for seeding in the coated 96-well plate.
   1. Detach the cells from the culture flask, resuspend them in the fresh growth medium, and count them using the trypan blue staining method.
   2. Spin down the cells in a 50 mL tube for 5 min at 400 x g at RT.
   3. Resuspend the cells in a fresh growth medium (DMEM + 10% FBS + 1% HEPES, no antibiotics) to obtain a cell density of 0.1 x 10⁶ (viable) cells/mL.
3. Remove the gelatin solution from the black-walled plates with a clear bottom by flipping over the plate and drying it on a tissue. Add 200 µL/well PBS to remove excess gelatin and flip over the plate again.
4. Dispense 200 µL of the cell suspension (corresponding to 0.2 x 10⁵ cells) per well in the gelatin-coated 96-well plate (from now on this plate will be further referred to as the "measurement plate").
5. Incubate the plate overnight at 37 °C and 5% carbon dioxide (CO₂).

3. Loading of the Cells with a Fluorescent Ca²⁺ -sensitive Dye

1. On day 1 perform the actual Ca²⁺ mobilization assay, starting with loading of the seeded cells with the fluorescent Ca²⁺-binding dye fluo-2 acetoxymethyl (fluo-2 AM).
   1. Prepare assay buffer by adding 40 mL HEPES (1 M) to 200 mL Hank's Balanced Salt Solution (HBSS, 10x, no phenol red, no sodium bicarbonate). Add ultrapure water to obtain a final volume of 2 L and add 4 g bovine serum albumin (BSA). Dissolve the BSA via magnetic stirring. Adjust the pH to 7.4 (with sodium hydroxide, NaOH) and filter the solution.    
      NOTE: During all following steps the same buffer, referred to as "assay buffer", is used.
   2. Prepare a stock solution (4 mM in dimethyl sulfoxide (DMSO)) of the fluorescent Ca²⁺-sensitive dye fluo-2 AM. Avoid excessive exposure to light. Dissolve 1 mg fluo-2 AM (molecular weight: 1,061 g/mol) in 235.6 µL DMSO.
   3. Prepare a working solution of fluo-2 AM. For one 96-well assay plate mix 12.5 µL of fluo-2 AM stock solution (4 mM) and 12.5 µL nonionic surfactant polyol (e.g., pluronic f-127) solution (20% weight/volume in DMSO, see Table of Materials). Add 22 µL of this mixture to 11 mL assay buffer in a 15 mL tube. The final concentration of fluo-2 AM is 4 µM. 
      NOTE: The nonionic surfactant is used as a dispersion agent to improve the aqueous solubility of fluo-2 AM and, in consequence, to enhance cell loading. To solubilize the surfactant in DMSO, heat the sample at 37 °C and mix regularly to obtain a homogenous solution.
   4. Remove the growth medium from the previously seeded cells in the 96-well measurement plate by flipping over the plate and drying it on a paper towel.
   5. Add 100 µL of loading dye solution per well using a multichannel pipette and incubate for 45 min at RT in the dark.

4. Preparation of 96-well Polypropylene Plates Containing the Chemokine Ligand CXC chemokine ligand 12 (CXCL12) or the Compounds under Investigation

1. Obtain round-bottom polypropylene (PP) 96-well plates (see Table of Materials).
   1. Allow the stock solution of CXCL12 (1 mg/mL) and the stock solution of the compounds under investigation to equilibrate at RT.    
      NOTE: The stock solution of CXCL12 is prepared in ultrapure water supplemented with 0.01% Tween20 and stored at -20 °C as single-use aliquots.
   2. Prepare a 5x concentrated solution of CXCL12 in assay buffer (250 ng/mL which equals 31.25 nM) and a 5x concentrated dilution of each compound (in assay buffer).
   3. Dispense the stock solutions into the appropriate 96-well plate according to a predefined plate layout. Dispense 75 µL/well in the plate containing CXCL12 (the "chemokine plate"), and dispense 50 µL/well in the plate containing the compounds (the "compound plate").
      NOTE: Both compounds and CXCL12 will finally become diluted 5x upon dispensing in the measurement plate. It is also important to include negative control wells containing only assay buffer in both the compound plate as well as the chemokine plate. Also, positive control wells need to be included (i.e., wells with CXCL12 in the chemokine plate, but with assay buffer in the corresponding wells of the compound plate).

5. Protocol Settings on the Fluorescence Microplate Reader

NOTE: The fluorescence microplate reader used in this protocol is referred to in the Table of Materials.

1. Switch on the cooler unit of the fluorescence plate reader first, and then switch on the fluorescence reader itself. Let initiate for a few minutes and open the system's software.
2. Create the assay's protocol using the drag-and-drop menu that includes the following steps:
   1. In the "Settings" box, select 'Read_Mode' with excitation wavelength: 470-495 nM; and emission wavelength: 515-575 nM.       
      NOTE: The selected wavelengths are appropriate for use with the Ca²⁺-sensitive dye fluo2.
   2. Include the "Mix with TF" (transfer fluid) box for the automatic mixing of the compounds in the compound plate, which will be put at the source 2 positions of the device (see step 6.4). Select 15 µL of solution in each well to be automatically aspirated and mixed three times. Adjust the height of the pipette tips to 20 µL below the liquid surface. Set the speed of aspiration and dispensing at 50 µL/s.   
      NOTE: The height of the pipette tips refers to the volume that is left underneath the pipette tips. For instance, if the wells of a compound plate contain 50 µL, a height of 30 µL corresponds to 20 µL below the liquid surface. The volume of compound solution taken to mix, the speed of mixing, the position of the pipette tips during mixing, and the number of mixing cycles can all be adjusted using the software.
   3. In the "Transfer Fluid" box, define that 20 µL of each well from the compound plate will be transferred into the measurement plate. Position the pipette tips at 20 µL below the liquid surface i.e., at a height of 30 µL during aspiration of the compounds and at a height of 60 µL during dispensing in the measurement plate. Set the speed of aspiration at 50 µL/s and the speed of dispensing at 25 µL/s.      
      NOTE: The compound plate contains 50 µL of solution per well (see step 4.1.3), thus a height of 30 µL corresponds to a position 20 µL below the liquid surface. The measurement plate will contain 80 µL of assay buffer per well (see step 6.3), thus a height of 60 µL corresponds to a similar position of the tips.
   4. Select the "Read with TF" button. During the first interval, select 60 reads (fluorescent measurements) with a read interval time of 1 s. Define that 10 reads are recorded before dispensing the compounds into the measurement plate, and 50 reads afterward. Define the second interval with a read interval time of 30 seconds and 18 reads. 
      NOTE: During this step, fluorescence will be measured kinetically (at the defined time intervals) for ~ 10 min in total.
   5. Include the "Wash Tips" button in the protocol three times. Within each washing step, select the fluid type: fluid A (ultrapure water) or fluid B (70% ethanol), the number of wash cycles (1), the pump speed (Fast), and the number of strokes (5) during each wash step.      
      NOTE: During the first and last wash steps fluid A is used, during the second wash step fluid B is used.
   6. Include the "Pause Pipettor" function. Set the pipettor to pause for 300 s.
      NOTE: By including this step in the protocol, the pipettor head, which is integrated in the fluorescence plate reader device, will pause for 5 min before continuing the protocol.
   7. Include the "Mix with TF" box to allow automatic mixing of the CXCL12 solution in the chemokine plate, which will be positioned at the source 3 position of the device (see step 6.4). Select 15 µL of solution in each well to be automatically aspirated and mixed three times. During aspiration and mixing, position the pipette tips 20 µL below the liquid surface. The speed of aspiration and dispensing is set at 50 µL/s.        
      NOTE: Similar to step 5.2.2, these parameters can be adjusted.
   8. In the subsequent "Transfer Fluid" box, define that 25 µL from each well of the chemokine plate will be transferred into the measurement plate. Position tips 20 µL below the liquid surface i.e., at a height of 55 µL during aspiration from the chemokine plate that contains 75 µL/well (see step 4.1.3), and at a height of 80 µL during dispensing in the measurement plate. Set the speed of aspiration at 50 µL/s and the speed of dispensing at 25 µL/s.
   9. Include the "Read with TF" button. During the first interval, select 145 reads with a read interval time of 1 s. Define that 5 reads are recorded before dispensing CXCL12 into the measurement plate, and 140 reads afterwards. Define the second interval with a read interval time of 6 s and select 20 reads.
      NOTE: Taken together, throughout the entire protocol 243 reads (fluorescent measurements) are recorded: 78 at step 5.2.4 and 165 during step 5.2.9.
   10. Similar to step 5.2.5, include the "Wash tips" button three times in the protocol.
       NOTE: Including this step at the end of the protocol allows that the tips can be re-used to perform another assay without needing to change the tips.
3. Set the temperature of the device at 37 °C by clicking the "Set stage temperature" button and selecting 37 °C.

6. Running the Fluorescence Assay

1. After the measurement plate has been incubated with loading buffer for 45 min (see step 3.1.5), remove the buffer by flipping over the plate and dry it on a tissue.
2. Wash the seeded cells by adding 150 µL/well of assay buffer and incubate for 2 min.
3. Remove the buffer again by flipping over the plate. Add 80 µL/well of assay buffer with a multichannel pipette.
4. Put all the plates into the device at their appropriate position: the compound plate at the source 2 position, the chemokine plate at the source 3 position, and the measurement plate at the read position. Put a box of black tips at the source 1 tips position. Shut the device's door and incubate for 5 min before continuing the protocol.
5. Select the "Protocol signal test" button to determine the background relative light units (RLUs) and fluorescence variance over the plate. A new window will pop up. Select "Test signal".
   NOTE: During this step, background RLUs are determined by the intensified charge-coupled device  (ICCD) camera, which is integrated into the optics compartment of the fluorescence reader device. These RLUs result from the excitation of the Ca²⁺-sensitive dye (fluo-2) by light-emitting diodes (LEDs) (LED output wavelength 470-495 nm). Values of 8,000-10,000 RLUs are well suited for this application. RLUs can, if necessary, be adapted by changing the excitation intensity (Select Exc. Intensity), or the camera gate (Select Gate Open). The variance over the plate should ideally be less than 7.5%.
6. Select "Update" if background values of 8,000-10,000 RLUs are obtained. Save the main protocol by clicking the Save button.
   NOTE: By doing so, the settings from the protocol signal test will be carried over to the main protocol.
7. Run the assay by pushing the "RUN" button.