The development of 384-well radioligand receptor

GE Healthcare
Application Note 28-4093-72
LEADseeker Multimodality Imaging System
The development of 384-well radioligand receptorbinding LEADseeker assays using the LEADseeker
Multimodality Imaging System
Key words: LEADseeker • SPA Imaging Beads
• assay development
Materials
LEADseeker™ assays involve the use of a solid-phase bead
particle containing scintillant that is stimulated to emit light
when radioactivity is in close proximity. There are two types
of core bead suited for use with the LEADseeker Multimodality
Imaging System: yttrium oxide (YOx) and polystyrene (PS), and
these are supplied with a range of coatings such as wheat
germ agglutinin (WGA), polylysine, or polyethyleneimine (PEI)
for use in receptor-binding assays.
LEADseeker Multimodality Imaging System Radioligand receptor-binding assays using the LEADseeker
assay format are homogeneous and all of the components
are usually added at the start of the experiment. All steps
of a LEADseeker radioligand receptor-binding assay may
be automated using appropriate robotic systems for manual
and liquid handling. When performing radioligand receptorbinding assays using SPA Imaging Beads and LEADseeker
instrumentation, an entire 384-well plate can usually be imaged
in 5 min or less, which enables much higher throughput than that
achieved when counting plates using PMT based instruments.
This application note provides a step-by-step guide describing
how to successfully develop robust validated 384-well
radioligand receptor-binding assays using the LEADseeker
Multimodality Imaging System.
Products used
SPA Imaging Select-a-Bead Kit*
18-1174-91
RPNQ0291
* Contains 50 mg each of WGA PS, WGA YOx, WGA PEI Type A, WGA PEI Type B,
Polylysine PS, and PEI-PS for convenient selection of optimum imaging bead (Step 2).
Other materials used
Membrane containing receptor of choice
Radiolabeled ligand
Non-radiolabeled ligand for determination
of non-specific binding (NSB)
Assay buffer
384-well white flat bottom polystyrene
NBS™ microplates
GraphPad Prism™ software (Corning, 3652)
(GraphPad Software)
Protocol
Step 1. Selection of assay buffer
The assay buffer employed for SPA or filter-binding format
assays will usually prove suitable for LEADseeker assays and
these should always be used for Steps 1 to 4. However, if a
suitable assay window (total binding – NSB) is not achieved in
Step 4, then re-optimization of the assay buffer may be required.
Reagents such as BSA (0.1 to 0.5% w/v) or NaCl (10 to 100 mM)
may be added to reduce NSB of the ligand to the bead or
membrane. Protease inhibitors may be added to the buffer
to improve signal stability (Step 5).
Step 2. Selection of appropriate SPA Imaging Bead
The approximate capacity of SPA Imaging Beads for membrane
protein is typically in the order of 10 µg of membrane protein
per mg of bead; this ratio is therefore used for initial experiments.
Alternatively, if converting from an existing SPA, the existing
SPA bead to membrane ratio should be used. The radioligand
concentration should initially be used at approximate Kd
concentration (determined from SPA or filter-binding assays).
NSB is determined in the presence of high concentrations,
usually 10 × Kd, of unlabeled ligand. Assays are typically
incubated at room temperature and imaged for 5 min using
3 × 3 binning with quasi-coincident averaging at regular
intervals to ensure that binding equilibrium is reached (Fig 1).
The important considerations are:
• NSB of the radiolabeled ligand
• Total bound integrated optical density (IOD) units
• % ligand depletion (Appendix 2)
It should be noted that the plate can have a significant impact
on ligand depletion and therefore it is worthwhile determining
the % binding of the ligand to the assay plate by including
wells containing radioligand only. If the radiolabeled ligand
is shown to bind > 10% to the assay plate, the use of treated
plates such as 384-well white flat bottom polystyrene NBS
microplates should be considered.
In the example shown in Figure 1, the WGA-coated PS beads
exhibited the lowest NSB in both the presence and absence of
membrane. This bead type also gave the highest assay
window and was therefore selected for further assay
development despite the initially low total bound IOD.
Fig 1. Total and NSB observed using a range of bead types. Assay was set up
in a 384-well non-treated assay plate in a total volume of 40 µl. System signal
was removed for analysis (Appendix 1). Data points are mean of three replicates
with error bars ± SD.
Step 3. Optimization of membrane to bead ratio
Once the bead type has been selected, the next step is to
determine the actual capacity of the imaging bead for
receptor membrane (the membrane to bead ratio). A matrix
of the imaging bead selected in Step 1 and the receptor
membrane is performed and typical data obtained from such
an experiment are shown (Fig 2). When the specific signal
(total – NSB) is plotted, the optimum membrane to bead ratio
can be selected. In this case, 0.13 mg of bead was sufficient
to capture 1.3 µg of membrane protein. Increasing the bead
to 0.5 mg/well did not increase bound IOD indicating that
1.3 µg of membrane was bound by 1.3 mg of bead (dotted
line). The optimum membrane to bead ratio was therefore
10 µg of membrane/mg of bead.
Fig 2. Matrix of imaging bead and receptor membrane. Assay was set up in
a 384-well assay plate in a total volume of 40 μl. Assays contained 0.13, 0.25,
or 0.5 mg of bead with 1.3, 2.5, or 5 µg of membrane protein. System signal
was removed for analysis. Data points are mean of three replicates with error
bars ± SD.
Step 4. Optimization of bead and
membrane amount
Once the membrane to bead ratio has been established, the
optimum bead and membrane amount should be determined.
This is done by premixing the bead and membrane at the
ratio established in Step 3 and diluting in assay buffer to give
a series of dilutions containing varying amounts of bead and
membrane at a fixed ratio of membrane to bead.
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When the bead and membrane are premixed the resulting
suspension may occasionally aggregate causing difficulties
with pipetting; however, this may be resolved by brief sonication
(30 s) using a sonicating water bath. We have found that
premixing the bead and membrane prior to assay addition
generally improves the assay window and typical data obtained
from such an experiment are shown (Fig 3). The optimum
amount of membrane selected in this instance was 1.3 µg
(dotted line), corresponding to 0.13 mg of bead, giving a total
signal of ~ 100 IOD with a background (including system
signal) of ~ 20 IOD. The % ligand depletion (Appendix 2) at this
level of membrane was shown to be acceptable (8%).
Step 6. Determination of solvent tolerance
The tolerance of the assay to solvent, usually DMSO, is
determined by adding increasing concentrations of solvent
to the assay system. Typical data obtained from such an
experiment are shown (Fig 5). In this instance the assay was
found to be tolerant to 2.5% DMSO (v/v).
Fig 5. Bead and membrane were premixed at a ratio of 10 µg of membrane to
1 mg of bead and added to give 1.3 µg of membrane in the assay well. DMSO
was added to give final concentrations of 0 to 10% (v/v). Assays were set up in
384-well plates and incubated for 4 h at room temperature. Data points are the
mean of three replicates with error bars ± SD.
Step 7. Saturation binding analysis
Fig 3. Titration of premixed bead and membrane. Assay was set up in a
384-well assay plate. Bead and membrane were premixed at a ratio of 10 µg
of membrane to 1 mg of WGA-PS bead. Premixed bead and membrane were
serially diluted in assay buffer and added to the assay in one addition to give
5, 2.5, 1.3, 0.6, or 0.3 µg of membrane protein in the assay well. Data points
are mean of three replicates with error bars ± SD.
Step 5. Time course and stability of assay signal
Once the assay has been configured in terms of bead and
membrane additions, it is important to perform a time-course
analysis to ensure the assay is read at equilibrium and that
the assay signal is stable at equilibrium. This is done by setting
up the assay with the predetermined amounts of bead and
membrane (Step 4) and imaging at intervals for ~ 24 h. Typical
data obtained from such an experiment are shown (Fig 4); in
this case, binding equilibrium was obtained after 4 h incubation
at room temperature (dotted line) and the signal was stable
for at least 20 h. Occasionally, the assay signal may be found
to decline during incubation; we have found that this signal
decline is usually reversed by the inclusion of protease
inhibitors in the assay buffer (Step 1).
Saturation binding is readily performed with LEADseeker
assays. The assay is set up with increasing concentrations
of radiolabeled ligand in the usual manner and Figure 6
shows typical saturation binding data obtained from such an
experiment. The Kd was estimated directly from the binding
curve, in this case 0.2 nM (95% CI 0.17 to 0.23), which was in
agreement with the Kd estimated from the corresponding
filter-binding assay (0.19 nM, [95% CI 0.11 to 0.26]). Bmax
values can be estimated by sampling from the assay well
as described in Appendix 3. Using this method, the estimated
Bmax value of 3 pmol/mg was close to that estimated from
a filter-binding assay (4.5 pmol/mg).
Fig 6. Bead and membrane were premixed at a ratio of 10 µg of membrane to
1 mg of bead and added to give 1.3 µg of membrane in the assay well. Radiolabeled
ligand was added to give final concentrations of 0.035 to 4.2 nM in the assay well.
Assays were set up in 384-well plates and incubated for 4 h at room temperature.
Data points are the mean of three replicates with error bars ± SD.
Fig 4. Time course analysis. Bead and membrane were premixed at a ratio of
10 µg of membrane to 1 mg of bead and added to give 1.3 µg of membrane
in the assay well. Assays were set up in 384-well plates and imaged at regular
intervals for ~ 20 h. Data points are mean of three replicates with error bars ± SD.
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Step 8. Competition binding analysis
Step 10. Z′ analysis
The assay is further validated by competition binding studies.
These are carried out in the usual way to estimate IC50 and Ki
values from binding curves. Typical competition binding data
obtained using a range of competing ligands is shown (Fig 7).
To confirm the robustness of the assay, a Z′ analysis is
performed. The assay is set up with a number of replicate
values each for total and NSB wells. Between 50 and 100
replicates wells are typically set up to determine Z′. Typical
data obtained from 80 replicate wells is shown (Fig 9). In this
case, Z′ was determined to be 0.82 (1) which confirmed the
robustness of the assay.
Fig 7. Bead and membrane were premixed at a ratio of 10 µg of membrane to
1 mg of bead and added to give 1.3 µg of membrane in the assay well. Competing
ligands were added to give a range of concentrations in the assay well. Assays
were set up in 384-well plates and incubated for 4 h at room temperature. Data
points are the mean of three replicates with error bars ± SD.
Step 9. Determination of association and
dissociation kinetics
If required, on- and off-rate analysis is simple to perform
with LEADseeker assays. To estimate the on rate, the bead
and membrane should be mixed and incubated for 30 min,
preferably with the use of a roller mixer, before addition to
the assay. This allows the membrane to couple to the bead
ensuring that only the rate of ligand binding to the receptor is
measured. Following precoupling of the bead and membrane,
the assays are set up in the usual way. The assays are imaged
at regular intervals until binding equilibrium is reached.
Saturating levels of a competing ligand are then added and
the plate re-imaged to obtain the off rate. Typical data obtained
for this experiment are shown (Fig 8). In this instance, a onephase exponential association equation was fitted to the
specific binding curve and Kob determined to be 0.013/min.
Similarly, a one-phase exponential decay equation was fitted
to the dissociation curve and Koff determined to be 0.004/min.
Fig 8. Bead and membrane were precoupled at a ratio of 10 µg of membrane to
1 mg of bead and added to give 1.3 µg of membrane in the assay well. Assays
were imaged at 30 min intervals for 4 h before addition of competing ligand.
Following addition, the assay was re-imaged at 5 to 20 min intervals. Assays were
set up in 384-well plates. Data points are the mean of three replicates with error
bars ± SD.
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Fig 9. Z′ analysis of 80 replicates for total and NSB wells. Solid lines represent the
mean of the observations, dotted lines represent mean of the observations ± 3 SD.
Appendixes
3. Estimation of Bmax
1. System signal
Following imaging, the plate is centrifuged and a 5 μl sample
carefully removed from the supernatant of the total and nonspecific binding wells containing the highest concentration
of ligand (at saturation). The concentration of free ligand is
determined in both total and non-specific binding wells by
liquid scintillation counting.
The system signal is comprised of background phosphorescence
from predominantly the plate, and to a lesser degree, depending
on the amount present, the imaging bead. This phosphorescence
decays when the plate is stored in the dark and therefore, if
the assay is dark adapted before imaging, the contribution to
the total signal is negligible. However, if the plate is exposed
to light the contribution can be significant, particularly with
low signal assays (< 50 IOD). In these cases, the assay window
can be normalized by subtracting the system signal from both
total and non-specific binding wells prior to further analysis.
System signal wells are set up containing only bead and assay
buffer with the same total volume as the assay wells, and
imaged alongside.
A. Total and non-specific bound ligand is determined by
subtraction of free ligand from the total added ligand.
B. The specific bound ligand is determined by subtracting
non-specific bound ligand from total bound ligand.
C. Bmax is calculated from the specific bound ligand and
amount of protein in the usual way.
2. Estimation of ligand depletion
Ideally, ligand depletion should be < 10%. However, this may
be difficult to achieve in miniaturized assays with limited scope
for increasing assay volume. We have found that ligand
depletion only results in a significant shift in affinity above
30%, and therefore, whilst aiming to configure the assay at
< 10% depletion, < 30% is acceptable if it cannot be reduced
by an increase in assay volume, a decrease in binding protein,
or use of non-binding surface plates.
References
1. Zhang, J. et. al., Journal Biomolecular Screening 4(2), pp. 67–73 (1999).
Ligand depletion in LEADseeker assays is estimated as follows:
A. Following imaging, the plate is centrifuged (or left to settle
overnight) and a 5 μl sample carefully removed from the
supernatant of the total binding wells.
B. The free ligand is determined in total binding wells by liquid
scintillation counting.
C. Total bound ligand is determined by subtracting free from
total added ligand.
D. % ligand depletion is determined by dividing bound ligand
by total added ligand × 100.
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GE, imagination at work, and GE monogram are trademarks of General Electric Company.
LEADseeker is a trademark of GE Healthcare companies.
All third party trademarks are the property of their respective owners.
LEADseeker is covered under US patent number 6345115 and under US patent
numbers 6441973, 6381058, 6498690, and 6563653 and equivalent patents and
patent applications in other countries in the name of GE Healthcare Niagara Inc.
© 2007 General Electric Company—All rights reserved.
First published October 2007
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