Section10:Method Development and Optimization

From Assay Guidance Wiki

Optimization Experiments for GPCR Targets Coupled to Ca+2 Mobilization

Early method development should include the following experiments to demonstrate the validity of the assay concept:

1. Gq coupling (or promiscuous G-protein coupling) of the cells expressing the GPCR should be demonstrated. Load selected cell clones with Fluo-3AM or other suitable dye, trigger Ca+2 flux with a known agonist, and measure fluorescence signal. Select the clones with the most robust response.
2. Determine whether cells need to be constantly maintained in culture or whether they can be prepared as frozen aliquots to be thawed and plated the day prior to the assay. The use of frozen cell stocks is a convenient and efficient alternative if it can be shown that the FLIPR™ signal is sufficiently robust and stable.
3. Conduct dye-loading experiments. Select the combination of cell line, agonist and dye concentrations that produces the most significant signal window. Use a control cell line without receptor expression to establish signal base line. Choose between use of cells in culture and frozen cell stocks.
4. Conduct preliminary experiments to establish a reasonable cell density that could be further optimized in subsequent experiments as described below.
5. Using a known antagonist or potentiator, demonstrate that the Ca+2 mobilization induced by the agonist can be blocked or enhanced, respectively.
6. Test poly-D-lysine coated plates with selected cell lines and conditions demonstrated in preliminary experiments. Select the plate with a stable and acceptable signal window.
7. Establish preliminary growth conditions and DMSO tolerance for the selected cell line.

Statistical experimental design can be employed to optimize these conditions and the following factors should be included:

• Cell clones
• Cell seeding density/well
• Type of dye (wash vs. no-wash)
• Dye loading concentration
• Dye loading temperature
• Dye loading duration
• Coated plate type
• Buffer additives: eg: probenecid, concanavalin A, etc.
• Height, speed and mixing of FLIPR pipettor
• Volume of addition

Notes on Optimization Experiments for GPCR Targets Coupled to Ca+2 Mobilization

Some general points regarding a FLIPR™ assay for GPCRs need to be noted:

• Some receptors contain trypsin-sensitive sites in their extracellular domain that results in a loss of response if the cells are harvested by trypsinization. In these instances, cells should be harvested by either scraping or using enzyme-free dissociation buffer.
• Care should be taken when removing media and dye from the cell plate. It is common for mechanical aspiration to disrupt the cell monolayer, resulting in a deterioration of the assay performance. It is recommended to manually invert the plate and shake or “flick” the liquid out of the plate and blot onto paper towels if you are using a dye that requires washing.
• Several no-wash dyes are commercially available. Testing of multiple dyes is strongly recommended, as signals differ widely. Depending on the receptor studied, media may interfere with the no-wash dyes, so testing both with and without media may be required. An example of the difference between the signal obtained from the traditional Fluo-3 dye and the new Calcium 4 no-wash dye is shown in Figure 5.
  Figure 5: Comparison of different Ca+2 dyes on maximum response of a GPCR. In this example, a no-wash dye produced a significantly larger signal window than the traditional Fluo-3 dye. Signal windows are specific to receptors and cell lines, so it is recommended that testing be done during the initial optimization to ensure the appropriate choice of dye.
• Probenecid should be included in the dye and the buffer following dye loading whenever using CHO cells (5mM probenecid is sufficient). This prevents the release of dye from the cells back into the medium. AV12 and HEK293 cells do not require probenecid.
• CHO cells are dye-loaded at 37°C, whereas AV12 and HEK293 cells can be dye- loaded at 25°C.
• Poly-D lysine coated plates can improve the results obtained from some cell lines.
• Variability in the signal obtained on the FLIPR™ can sometimes be improved by adjusting the tip height or dispense speed on the FLIPR™.
• The standard assay buffer used in FLIPR™ experiments is HBSS with 20mM HEPES.
• The most common fluid addition volumes for a FLIPR assay are:

  	Volume per Well
  	96-Well Format	384-Well Format
  Dye	50ul	20ul
  Buffer	50ul	20ul
  1st addition in FLIPR	50ul	20ul
  2nd addition in FLIPR	100ul	20ul

The development of a FLIPR assay generally requires the following experiments:

1. Cell density determination and incubation time:
   This is typically the first parameter that is examined. The best way to assess cell density requirements is to seed an entire assay plate at a single density; therefore, several plates are required to examine multiple cell seeding densities. The cells should be examined on the FLIPR™ using buffer in the first addition and a maximal concentration of agonist in the second addition. This will allow one to assess the extent of variability within the plate and detect any patterns in variability. The most common variability pattern we have observed is an edge effect which can usually be resolved by increasing the cell density or the humidity during incubation. We recommend examining the following cell densities for the indicated cell types:

   	Seeding Densities (cells/wells)
   Cell Line	96-Well Format	384-Well Format
   AV12	30K, 40K, 50K, 60K	20K, 30K, 40K, 50K, 60K
   CHO	10K, 20K, 30K, 40K	5K, 10K, 15K, 20K, 30K
   HEK293	30K, 40K, 50K, 60K	20K, 30K, 40K, 50K, 60K

   
   

   Some assays will perform best with a 24-hour incubation time prior to assay, while others may need a 48-hour incubation time.
2. Dye loading time, dye concentration and temperature:
   The optimal dye loading can range from 30 minutes to 2 hours depending on the cell line and the dye used. The concentration of Fluo-3 used in the majority of FLIPR assays is 8µM. Lower concentrations can be examined in order to reduce the cost of the assay. The no-wash dyes have been shown to be effective at lower concentrations as well. CHO cells are dye loaded at 37°C, whereas AV12 and HEK293 cells can be dye loaded at 25°C.
3. DMSO tolerance:
   DMSO can alter the response of the cells as well as shift the dose response curve for agonist. It is recommended to perform an agonist dose response curve in the presence of different concentrations of DMSO in order to assess the DMSO tolerance of the assay. Extreme care should be taken if a DMSO concentration >0.1% is required.
4. Agonist/antagonist dose response curves:
   The reproducibility of the assay can be examined by performing two independent days of agonist/antagonist/or potentiator dose-response curves. The EC50/IC50 values should remain relatively constant over the course of the two experiments.
5. Full plate variability and Z’ factor determination:
   The variability of the assay is determined by running triplicate max/mid/min plates on three days and then calculating the Z’factor.

Considerations When Performing 384-well FLIPR™ Assays

384-well FLIPR assays have a number of challenges that are not apparent in the 96-well format. The first is mixing in the well. Most 96-well experiments are designed to allow a larger volume to be added to a larger space where mixing is not a concern. In a typical 96-well assay, 50µl of test compound are added to 50µl of buffer in the cell plate at a height of approximately 80 to 95µl. The height is the liquid height in the well at which the tips dispense. The 384-well plate is limited to a maximum volume of a 30-µl addition in a much smaller diameter well, and using the 96-well technique will result in variable response. When adding to a 384-well plate, the tips are typically in the buffer solution of the cell plate when the dispense takes place. In a number of cases, the speed of dispense has to be increased as well. These heights and speeds should be tested with buffer to check for unwanted “pre-firing” of the cells. Another issue that arises with the 384-well format is the limited amount of diluent that can be added to the compound plate. This limitation can result in having to create intermediate dilution plates off-line, thereby slowing throughtput and adding costly consumables. This has been eliminated by using an in-tip dilution on the FLIPR™ (Figure 6). Although the final DMSO concentration is the same, the bolus of DMSO in the bottom of the tip can have an effect on the cells (Figures 7a and b). In our hands, a ratio of 15µl buffer/5µl compound was found to have the least DMSO effect. However since this result can be variable, different combinations should be tested during development. This in-tip dilution method can be used in both the two- and three-addition FLIPR™ methods.

Notes on Tip Washing

The FLIPR™-2 and FLIPR™-3 have tip wash stations that can be incorporated into the assay to eliminate the need to change tips. This allows one to use reservoirs without fear of cross contamination among the test compounds. In addition, a DMSO pre-wash can be performed at the tip load station with the proper adapter. When running a single-point screen of more than 100K compounds, tip washing should be tested first to minimize cost and maximize throughput. Occasionally, the compound used for the EC90 addition cannot be washed off the tips, resulting in significant carry-over of active compounds in to the subsequent plate (example in Figure 8); in these cases, the tips will have to be changed. This typically happens when peptides are added as the EC90 dose.

Optimization Experiments for Ion Channel Targets with Ca+2 Permeability

Some ion channels (e.g. ionotropic glutamate receptors) differ from GPCRs in that they desensitize very quickly to agonist exposure, and in most cases, it is not possible to see a response in FLIPR™ with agonist alone. Such targets require the use of agents that decrease the rate of desensitization, which are called channel modulators or “clamps”. The choice of which channel modulator to use is dependent upon the receptor. The following is a brief summary of modulators that we have used:

Since ion channel modulators are needed to decrease the rate of desensitization of the channel to agonist, the assay design is somewhat different than for GPCRs. Like for GPCRs, the ability of the FLIPR™ to make two fluid additions to the cells enables the detection of agonists, antagonists, and allosteric modulators in one assay. Representative kinetic profiles for iGluR1 flip and flop are shown in Figure 9A. Test compounds are added in the first addition along with a 90% dose of the known agonist, in this case glutamate, which normally does not generate a measurable Ca+2 response because the rate at which the receptor desensitizes is too fast to be detected on the FLIPR™. A response in the first read will indicate that the test compound is either a non-desensitizing agonist or a positive allosteric modulator (Figure 9B). The second addition consists of an optimal concentration (~90%) of a known allosteric modulator which results in maximal response by clamping the channel open and decreasing receptor desensitization. A reduced response in the second read will indicate that the compound is an antagonist (Figure 9C). The question of whether the compound is a non-desensitizing agonist or an allosteric modulator will be answered in the secondary assay in which the compound is added in the absence of any glutamate in the first read. If the compound alone elicits a response, it is a non-desensitizing agonist. Alternatively, if the compound only gives a response in the presence of glutamate (read 2), then it is a potentiator.

In the case of the Kainate receptor iGluR6, the allosteric modulator ConA needs to be incubated on the cells for a minimum of 5 minutes prior to adding agonist. ConA takes longer to bind and has an effect on receptor desensitization.

Optimization Experiments for Ion Channel Targets with Ion Permeability that Significantly Impacts Cell Membrane Potential

Changes in membrane potential associated with ion channel activity may be measured on the FLIPR™ instrument using a voltage-sensitive dye available from Molecular Devices. The following are some of the parameters that need to be considered in developing a FLIPR™-based membrane potential assay:

• Cell Density: Optimal cell conditions for the FLIPR membrane potential assay require the creation of a confluent cell monolayer. The cell seeding density depends on the cell type and the time in culture following the plating of the cells. Receptor expression levels can change with the cell passage number or as a result of the drug-selection conditions used for cell maintenance. Thus, it is critical to monitor changes in functional activity over time. Refer to the previous in this chapter for optimizing the cell seeding density.
• Assay Buffer: HBSS + 20mM HEPES + added CaCl2 (5mM final concentration).
• Preparation of Membrane Potential Dye: We recommend dissolving the dye in assay buffer. After formulation, the loading buffer can be stored frozen in aliquots for several months without loss of activity.
• Method of Dye Loading Cells: Dilute the loading buffer 1:1 with assay buffer. Aspirate the media from the cells and add 100µl of diluted buffer per well for 96-well plates. (Note: We have not had success following the Molecular Devices recommendation of adding the dye directly to the media with the iGluR targets.) The dye:buffer ratio can be optimized to reduce cost of the assay. Dye-loading the cells should be tested at 37C and at ambient temperature. The optimal dye loading time, on average, for HEK293 cells is 60 minutes, but the range can be wide (5-60min).
• Antagonist Assays – Results Export Range: The kinetic profile of the calcium response to ion channel activation is prolonged when compared to the typical profiles generated by GPCR activation. As a result, agonists introduced in the first addition, read frame I, will lead to a baseline shift which will not return to baseline prior to the second addition, read frame II (see figure 9B). This baseline shift within read frame II is due to the prolonged activation of receptor when agonists are introduced. Because the EC90 challenge dose for antagonists assays is added within the initial portion of read frame II, the read frame I baseline shift due to agonists will lead to antagonist assay interference if exporting data from read frame II only (Max-Min). For this reason, one should consider exporting both read frames I and II for ion channel antagonist assays, which includes the pre-compound addition portion of read frame I, to capture the pre-compound addition or actual assay baseline (Figure 9B, time 0-350 seconds). By utilizing the pre-compound addition baseline of read frame I, false positive agonist interference in antagonist ion channel targets can be avoided.
• Clamp: Clamping agents such as Concanavalin A may be required to prevent rapid desensitization of ion channels. Depending on the incubation time required for the clamp, it could either be added with the loading buffer or it could be added with the compound.
• FLIPR Setting: Choose filter #2 in the experiment setup of the FLIPR™ software to measure membrane potential. Set the background reading ~ 20000 RFU. The following are some of the recommended setup parameters for the compound (1st addition) and agonist (2nd addition) additions to a 96-well plate.

  	Volume	Addition Speed	Pippettor Height
  1st Addition	50ul	50ul/sec	100ul
  2nd Addition	50ul	50ul/sec	150ul




• Control: We recommend running a KCl dose curve as a positive control to measure changes in membrane potential independent of the ion channel activity. The following is an example of time course tracings observed with the iGluR6 assay (Figure 10).

Performing FLIPR™ Using a Non-Adherent Cell Line

So far, we have been describing methods appropriate for adherent cells cultures. In these cases, dye can be loaded directly onto cells grown to confluency in microtiter plates. In contrast, when the transfected cell line is weakly adherent or grows in suspension culture the following procedures should be followed:

1. Remove growth media from cell culture flask.
2. Add 10ml PBS to each flask to rinse.
3. Remove PBS and repeat rinse step.
4. Add 10ml cell dissociation buffer to each flask.
5. Rock flask gently.
6. Add 10ml Alpha-MEM and discard the rinse.
7. Transfer cells to 50ml centrifuge tube.
8. Add 30 ml buffer.
9. Pellet cells for 5 min at 2000 rpm.
10. Remove supernatant.
11. Add 30ml buffer with 30 µL Fluo-3 AM (1:1000 dilution) and 30 µl pluronic acid.
12. Cover tube with foil and shake gently.
13. Place on shaker for 60 min at 180 rpm at room temperature.
14. Fill up tube with buffer and spin for 5 min at 2000 rpm and remove supernatant.
15. Repeat step #14.
16. Resuspend cells at 1 x106 cells/ml.
17. Plate 50 µl/well of Poly-D-Lysine pre-coated plates.
18. Wait 20 min and centrifuge plates for 3 min at 1500 rpm.
19. Place plates in FLIPR until ready for use.

Notes:

• If cells are weakly adherent, start at step #1.
• If cells are in suspension, start at step #7.
• If using a no-wash dye, skip steps #14-15.