Maximize Your Microplate Assay Results
eBook
Published: October 8, 2024
Credit: BMG LabTech
Today’s research heavily relies on microplate readers for a variety of applications, from cell-based assays to molecular biology studies. However, obtaining reliable data depends not only on the technology but also on the choices you make for your assay.
Selecting the appropriate microplate for specific experiments is critical to ensure accurate results and to minimize variability.
This eBook explores best practices for choosing and optimizing microplate readers, considering several factors such as materials, detection modes and well formats to help researchers maximize their output.
Download this eBook to discover:
- How to select the right microplate for various assays
- How well shapes and materials affect data accuracy
- Practical tips for reducing signal interference and improving data consistency
MICROPLATES
IN ACTION
Recommendations for use
2
Today’s research often involves microplate readers as fast
and reliable tools to push experiments forward. Whether
it’s cell-based assays, high-throughput screening, or a host
of molecular biology applications, reliable and meaningful
data are crucial to support experiments.
This fi rst part of the eBook “Microplates in action” explores
ways to get the best out of your microplate. Why should you
choose a clear microplate versus a black one? What is the
best well shape for specifi c assays? Research requires the
right choices. But what are the crucial plate parameters
for different detection modes? And how do you measure
success?
The information presented here describes some of the best
practices gleaned from experiences in different research
settings. The aim is to help accelerate discovery and
support research goals.
FOREWORD
Dr Barry Whyte
Application Scientist
3
CONTENTS
CHAPTER 1
Use the right microplate ............... Page 4
CHAPTER 2
Sources of variance ............... Page 10
CHAPTER 3
Plate reader parameters ............... Page 15
CHAPTER 4
Performance indicators ............... Page 27
4
USE THE RIGHT MICROPLATE
Microplates consist of a plate with multiple
cavities or wells used as small sample tubes.
As a much-used standard in the life sciences,
they come in various colors, and offer different
well formats and shapes. Choosing the right
microplate is an important step in ensuring
the success of your assay. Here we look at
some of the options and considerations for
the selection and use of the right microplate.
A. CLEAR MICROPLATES
All ultraviolet (UV)/visible absorbance and
colorimetric measurements need a clear
(transparent) plate since detection requires
light to pass through the sample. A plate
with a clear bottom (but not a clear plate)
is essential for bottom reading assays. For
bottom fl uorescence or luminescence assays,
a clear plate is detrimental as black or white
walls are required. Assays with adherent cells
where you may want to measure from beneath
the plate require clear bottoms. Moreover, if
you wish to avoid interference from bubbles
or other confounding factors present at the
surface of the well, you need transparent
bottoms on your plates to read signals from
underneath.
Standard polystyrene plates are transparent in
the range of visible light. However, polystyrene
does not transmit UV light (< 320 nm) and
is hence inappropriate for the detection of
nucleic acids or other macromolecules that
absorb at or below 320 nm. For this purpose,
UV-clear or UV-transparent microplates
must be used. These are made of cyclic olefi n
copolymer (COC) and allow for an improved
ultraviolet light transmission in the range
200 – 400 nm (Fig. 1).
CHAPTER 1
Fig. 1: Transmission spectra of clear and UV-transparent microplates
5
B. BLACK MICROPLATES
Black microplates partially quench the signal
of the sample as the black color partially
absorbs the light signal coming from the
sample. These plates are well suited for
fl uorescence intensity measurements
including Förster’s Resonance Energy
Transfer (FRET), and for fl uorescence
polarization assays. These are usually
detection modes with a strong signal yield
and the use of the black color helps to reduce
background, auto-fl uorescence and well-towell cross-talk, which in turn provide better
signal-to-blank ratios.
Black microplates are generally not
recommended for luminescence, timeresolved fl uorescence (TRF) or Time-Resolved
(TR)-FRET assays. These detection modes
usually have a comparably low signal yield
that could be further quenched by the black
color. However, in some circumstances,
extremely bright luminescence assays yield
better results in black plates. Commonly
used white-walled luminescence plates
look opaque to the human eye but are semitranslucent. It is easy to see a strong light
source through most white-walled plates.
These white-walled plates are suitable
for blocking the well-to-well cross-talk
transmitted through the plastic wall of a
plate but are not very helpful in the case of
an extremely bright chemiluminescent assay.
If a very bright luminescence test is being
performed, the detection of dimmer lower
intensity samples ceases to be a concern and
assay quality depends mostly on preventing
light spillover from the brightest to the less
bright wells nearby.
Manufacturers of TRF and TR-FRET tests
like Dissociation-Enhanced Lanthanide
Fluorescence Immunoassay (DELFIA®),
Lanthascreen™, Homogeneous TimeResolved Fluorescence (HTRF®), THUNDER™
and others generally recommend white
plates as these will improve the signal
yield. However, the trade-off is a reduced
assay window. Black plates can be used
in combination with high-sensitivity plate
readers to maximize data output. A sensitive
TR-FRET instrument gives the best results
and the highest dynamic range when
background fl uorescence is low.
Fig. 2: Photograph of selected microplates
6
C. WHITE MICROPLATES
White plates typically are recommended for
luminescence, TRF and TR-FRET assays.
The white color of the well partially refl ects
the sample signal helping to maximize it.
The drawback is that white microplates also
increase the background signal. However,
this increase in background signal is usually
quite low and rarely an issue in luminescent
assays. In TRF and TR-FRET, the delayed
measurement window eliminates the
infl uence of the background. It is therefore
benefi cial to collect as much as possible every
photon emitted from your sample.
If a photon originates from a well in a
luminescence test, something you want to
measure has emitted it. In a clear plate, most
of the photons emitted from the sample might
be traveling away from the detection optics
and pass through the walls of the plate where
they cannot be measured. If a black plate is
used, these photons would likely be absorbed.
In general, about six times more light is
collected in a white plate compared to a clear
plate, which signifi cantly improves the limits
of detection and reduces assay noise.
In addition, the solid white walls prevent
most signal from leaking from a bright well
to an adjacent well or even neighboring
wells two or three rows over. Once signal
is detectable by a luminometer, the most
important factor in assay quality is often
keeping cross-talk between nearby wells to
a minimum. For luminescence, minimizing
cross-talk is critical. A well that serves as
a negative control may have less than or
around 10 counts, while the brightest well
on the plate may approach 106
– 108
counts
Without
aperture
Detector
With
aperture
Detector
Fig. 3: Minimizing cross-talk
(please note that count numbers are relative
and may vary among plate reader models and
manufacturers). If even 0.5% of that signal
leaked from positive to nearby negative wells,
a count of 10 might display as 1,000,000. Highquality microplate readers have cross-talkpreventing apertures between the well and the
detection optics, but this feature is of little use
if light has a free path through the sides of a
well into the neighboring wells. White plates
with solid white walls help keep cross-talk
to a minimum and allow other cross-talkprevention features to be used for further
refi nement (Fig. 3).
7
Opaque sides also improve luminescence assays by improving uniformity. A well in the corner
of the plate has three other wells as neighbors. A well in the middle of the plate has eight. In a
clear microplate, the amount of stray light and refl ecting and refracting plastic varies across
the plate due to this unavoidable geometry. With opaque walls, the impact of this difference is
minimized and the uniformity of the replicates and linearity of standards and dose-response
curves are improved.
Even moderate cross-talk can have a negative impact on detection measurements. It can for
example make blanks or negatives highly variable reducing Z’ values.
D. GREY OR SILVER PLATES
Grey microplates are an intermediate solution between black and white ones. These are
specifi cally recommended for AlphaScreen® and AlphaLISA® (ALPHA for Amplifi ed Luminescent
Proximity Homogeneous Assay) as they reduce cross-talk and background while still providing
good signals. AlphaScreen® and AlphaLISA® produce intense light where cross-talk is a concern
but also benefi t from refl ective, signal-strengthening well walls. Like a white plate, grey or
silver plates are designed to have a low optical transparency but enough refl ectivity that they redirect signal toward an upper optic.
E. WELL AND BOTTOM SHAPE
Wells can be either round or square. Square wells contain a larger sample volume and increase
the light transmission area compared to round wells. Round wells instead have a smaller
total area and are better suited for shaking. Moreover, as round wells normally do not share a
common wall with adjacent wells, they are less affected than square ones by signal cross-talk
through the well wall. The different shapes of microplate well bottoms (F-, V-, U- and C-bottom)
are shown in fi gure 4. For most measurements, fl at bottomed (F-bottom) wells provide the
best results and light transmission. They are ideal for adherent cell cultures and are suited for
bottom-reading assays.
Fig. 4: Different well bottom shapes
F U V C
8
In a round bottom well (U-well), every position
in the well has a slightly different fl uid depth
and local environment. Therefore, even a
small shift in the measurement position
from well to well can mean that the depth
of fl uid directly under the optic is different.
Furthermore, the angle that excitation or
other light from an optic encounters at the
bottom of the plate varies considerably
depending upon where it interacts with the
slanted surface of a round bottom well. These
plates can still be measured, but fl at bottom
wells are preferred in most cases. Round
bottom wells can be useful when good mixing
and washing are critical to the performance
of the assay. The round shape of the well
allows plate shaking to increase the mobility
of the fl uid. If the fl uid volume in a 96-well
plate is necessarily low or variable, round
bottom wells help center any fl uid that has
been added to the well. On the one hand, this
makes detection more consistent compared
with the situation where each small drop has
a variable position in a fl at bottom well. On
the other hand, it enables easy and residuefree pipetting and is benefi cial for cells in
suspension and spheroids.
V-bottom (conical) wells enable maximal
volume retrieval of small and precious
samples because of their shape. However,
the conical shape is disadvantageous for
spectrophotometric applications. Curved
bottom wells (C-bottom) are a compromise
between F-bottom and U-bottom wells. The
fl at bottom makes them suitable for optical
measurements, while the rounded edges
facilitate mixing and washing.
F. PCR PLATES
The wells of plates used for polymerase chain
reaction (PCR) are often conical to allow low
sample volumes to rest and mix in the bottom
of each well Fig. 5. Un-skirted or half-skirted
PCR plates must have a skirt or adapter added
before the plate carrier of an instrument can
accept it and measurements can be made
in the reader. It is also useful for focusing
optics to be beneath the plate. Detection at
the bottom of the plate can put light onto a
sample that might be missed by reading at the
top of a PCR plate.
Fig. 5: PCR plates with conical wells
9
G. HIGH-BINDING, MEDIUM-BINDING OR CELL-CULTURE PLATES?
Typically, “standard” microplates are called medium-binding and are suitable for most work in
the laboratory. However, depending on the application, microplates with higher or lower binding
properties are benefi cial. For example, enzyme-linked immunosorbent assays (ELISA) benefi t
from high-binding while fl uorescence polarization assays benefi t from low-binding properties
since protein sticking to the side of the well could cause assay complications. High-binding and
low-binding plates each have coatings that make them suitable for their respective applications.
.
H. CELL CULTURE PLATES
Plates used for cell culture must always be sterile. When working with adherent cells, they must
also be treated to render them suitable for the attachment of cells. Classically, tissue-culturetreated polystyrene is used, which has been for example oxidized by plasma. The OH and COOH
groups introduced in this way result in a hydrophilic surface that enables cell adhesion (Fig. 6).
Furthermore, poly-lysine-coated plates or collagen-coated plates are often used. Cells adhere
more readily to the hydrophilic surface of poly-lysine-coated plates due to the electrostatic
interaction between the negative charge of the cell membrane with the poly-lysine cations. Both
poly-D-lysine and poly-L-lysine are used but poly-D-lysine is preferred since it is more stable
to degradation by cellular proteases. Collagen, an extracellular matrix protein, provides an
attachment framework for the adhesion and growth of cells. In each case, the goal of treatment
of the plates with poly-lysine or collagen is to promote cellular adherence to the bottom of the
well.
H H H
C C C
H H
OH HO
C
H
C
H
C
H
H
C
O O
C
H
C
H
H
C
H
H
C
H
C
H
H
C C
H
C
H
C
H
OH
OH
H
H C C
H
O
Surface treatment
Fig. 6: Polystyrene plate
surface with tissue-culture
treatment
10
SOURCES OF VARIANCE
A. REPLICATES. HOW MANY ARE ENOUGH?
The purpose of replicates is to provide confi dence in the ability to draw
a conclusion from a data set (Fig. 7) . For the most part, replicates in
microplate reader experiments are considered technical replicates
of the same sample. These replicates ensure that the method and
instrumentation are sound and not imparting variability. Since most
assays are based on well-established methods, the number of
replicates is often kept low (duplicates, triplicates) so that information
on a wider variety of treatments can be assessed.
When are more replicates benefi cial? Having triplicates or a higher
number of replicates helps determine if there are any outliers in your
replicates. Once identifi ed, these outliers can be removed before
additional evaluation is performed.
In the early stages of assay development and as a working rule
for measurements with cells, a higher number of replicates is
advantageous. The determination of the cumulative mean specifi es
when the mean converges to a particular value. This allows a
better appreciation for the variability imparted by the method and
instrumentation. Ideally this variation is as low as possible.
96
A
B
C
D
E
F
G
H
1
S1
S2
S3
S4
S5
S6
2
S1
S2
S3
S4
S5
S6
3
S1
S2
S3
S4
S5
S6
4
S1
S2
S3
S4
S5
S6
5
S1
S2
S3
S4
S5
S6
6
S1
S2
S3
S4
S5
S6
7 8
B
B
B
B
B
B
9
B
B
B
B
B
B
Blank Standard 10 11 12
Check timing Use enhanced dynamic range Start measurement OK Cancel Help
Sample
Content:
Basic Parameters Layout Concentrations & Volumes Shaking
Control Pos Ctrl Neg Ctrl
Empty
Groups
Index
Start value:
Constant Increase
On
0
Replicates
Number:
Horizontal
Reading direction:
Vertical
1
Fig. 7: Replicates in microplate layout
CHAPTER 2
11
B. WHAT VOLUME DO I NEED?
In general, the minimum amount of fl uid needed is the amount that suffi ciently covers the
bottom of the well to a uniform depth (Table 1). As a rule, it is typically recommended to use at
least one-third of the maximum fi ll volume of a well. In a standard 96-well plate, this is usually
around 100 µl. In most circumstances, larger volumes are preferred. At very low volumes,
surface tension dominates the interaction between the fl uid and the plastic surface it is in
contact with. This can cause undesirable meniscus effects that make detection less consistent.
Larger volumes in a well mean longer pathlengths. This provides more opportunities for
excitation or UV/visible illumination to interact with the column of fl uid under the optics. Low
volumes should be used in a plate designed to handle them. The same amount of fl uid in a 384-
well plate or a half-area 96-well plate creates a deeper column of fl uid. This column of fl uid has
more chance to interact with the detection optics directly above and below the microplate.
C. MENISCUS
A meniscus, which is defi ned as the curve in the upper surface of a liquid produced by surface
tension, may form any time a liquid is in contact with a solid vessel (Fig. 8). In a microplate,
each well represents a unique instance where this type of interaction may occur. Variability in
the shape and location of the formed menisci may arise from well to well. At the location where
a particular reading is taken this can lead to variability due to liquid level variation. This can
impact how consistent the results are for the reading of replicates. This is especially true in
absorbance assays.
The issues of variable meniscus can be largely addressed by plate choice. Round wells are
preferable to square wells. Plates that are free from imperfections are less likely to exhibit
variable meniscus effects. A concentration gradient of detergent or protein across wells —
something that reduces the surface tension of a liquid in which it is dissolved — can also
sometimes lead to a variation in meniscus that correlates with the concentration of surfactant.
Well number Recommended volume
6 2 - 5 ml
12 2 - 4 ml
24 0.5 - 3 ml
48 0.5 - 1.5 ml
96 100 - 300 µl
96 half area 50 - 170 µl
384 30 - 100 µl
384 low/small volume 5 - 25 µl
1536 5 - 15 µl
3456 1 - 5 µl
Table 1: Recommended volumes for different microplates.
12
In certain cases, variable meniscus effects in an absorbance assay can be addressed by
selecting a correction to the pathlength. This correction normalizes absorbance values
to a 1-cm pathlength using a water-peak-based approach. This correction is performed
independently in each well and accounts for pathlength changes due to a variable meniscus.
D. COMMON PIPETTING MISTAKES
Good experiments require careful planning. This includes ensuring appropriate (and calibrated)
pipettes are used. The correct tips for each type of pipette must be on hand. But even more is
possible during the pipetting process:
Make sure liquids and equipment are equilibrated at ambient temperature before you start. Air
pressure, relative humidity, and vapor pressure are all temperature-dependent properties that
can affect the volume of a sample delivered. Similarly, pipettes should be handled as little as
possible to reduce heat transfer to equipment.
A)
Water - no
meniscus
Typical
meniscus
Low detergent High detergent
Low DNA Medium DNA High DNA
B)
C)
Fig. 8: Different types of menisci and the effect of detergent
13
Consistency is key. Each time you
perform a pipetting action consider four
recommendations:
1) Avoid touching the side of the sample
container;
2) Immerse the pipette tip to the proper depth
below the meniscus (too little immersion
may lead to the risk of air aspiration; too
much immersion may lead to liquid
clinging to the tip);
3) Pause after aspiration: briefl y leave the tip
in the solution to ensure fi lling is complete;
4) Use the same pressure and speed for
pressing and releasing the pipette plunger.
Check and check again throughout the
pipetting process that you are following these
recommendations. Examining the pipette
tip before and after dispensing ensures that
no extra liquid is present on the tip from the
aspiration step. This allows confi dence that all
volumes aspirated have been dispensed.
E. AVOIDING AND REMOVING
BUBBLES
Bubbles in samples can cause light scattering
and lead to variability. If possible, degassing
solutions helps to reduce the air that is added
with the samples. Careful pipetting is also
helpful. For challenging situations, consider
using the reverse pipetting technique for
small volumes of reagents that tend to foam.
Here the tip is overfi lled by pushing the
plunger to the second stop position during
fi lling. Next, only push the plunger to the fi rst
stop during the dispensing step.
If bubbles persist, briefl y centrifuge the
plates. Another way to remove bubbles is to
pop each bubble with a needle or tip. Ethanol
can be misted over the plate with a spray
bottle. Alternatively, a small volume of ethanol
can be added to each well to remove bubbles.
F. EDGE EFFECTS AND EVAPORATION
Edge effects and evaporation are frequently
encountered when using microplate readers.
These effects occur when some of the
medium in wells around the perimeter of the
microplate evaporates during the incubation
(Fig. 9). The change in volume of the sample
can signifi cantly impact readings and the
quality of the results.
Fig. 9: Different volumes in
central and edge wells after
prolonged reading/incubation times
14
In some cases, researchers have avoided edge effects by leaving the outer wells of the plate
empty or by fi lling them with assay buffer. However, this practice considerably reduces
throughput and effi ciency. Other options are available to prevent evaporation of media from the
outer wells. Solutions include use of an evaporation lid, sealing fi lms or tape, or different ways
to regulate temperature. The selection of specialized microplates to help reduce evaporation is
also an option. Such microplates may include condensation rings to further minimize losses.
Others include an additional channel around the outer wells of a plate that can be fi lled with
water to reduce evaporation and enable higher homogeneity between the outer and inner wells of
a plate. Evaporation can also be reduced by maintaining high levels of humidity and limiting the
number of times incubator doors are opened. Microplate readers can offer different options for
temperature control. The Advanced Assay Stability (AAS) system from BMG LABTECH provides a
steady temperature that is unaffected by external environmental changes while the plate reader
is operating - a prerequisite for better assay stability and more reliable data.
G. CROSS-TALK. WHEN IT MATTERS AND HOW TO AVOID IT.
Ways to reduce the impact of cross-talk when using white microplates are outlined in Chapter 1,
section C, page 6. In addition, different layout options are often used for cross-talk reduction
(Fig.10).
Empty
Blank/Negative control
Positive control
Samples
1 2 3 4 5 6 7 8 9 10 11 12
A
B
C
D
E
F
G
H
High cross-talk
1 2 3 4 5 6 7 8 9 10 11 12
A
B
C
D
E
F
G
H
Low cross-talk
Fig. 10: Different layout options on a microplate to reduce cross-talk.
15
PLATE READER
PARAMETERS
A. NUMBER OF FLASHES
Most detection methods in microplate readers require an excitation
light source as the fi rst step in detection. The exception is
luminescence. Nearly all microplate readers utilize a light source that
fl ashes or pulses rather than one that emits constant illumination.
The primary light source on all BMG LABTECH readers is a xenon
fl ash lamp that is capable of very consistent output from fl ash to fl ash.
Typically, the user can select the number of fl ashes for a specifi c
detection protocol from a minimum of 1 to a maximum of around 200
fl ashes (depends on manufacturer and model). When using multiple
fl ashes, an average result is generated from the combined fl ashes to
produce raw data values. This helps to reduce variability. However,
a higher number of fl ashes will increase detection time. The only
exception to the fl ash or pulse requirement is nephelometry, which
employs a specifi c laser setup. For most detection options the question
is therefore how many fl ashes are needed?
ABSORBANCE
If microplate read times are the priority, one or two fl ashes will likely
yield quality results. However, increasing the number of fl ashes to fi ve
only changes the read time for an entire plate by a few seconds since
the fl ash lamp operates at a very high frequency. While a minimum of
fi ve fl ashes works in most circumstances, 20 fl ashes or higher can be
benefi cial for certain applications.
More fl ashes can be used when speed is not a primary concern
during microplate reading. Here, the use of 22 fl ashes is a typical
recommendation since there are diminishing returns for improvement.
The one exception is when high-resolution spectra are needed. In this
case, 40 or even 50 fl ashes are useful when the absorbance of the
peaks is in the 3–4 OD range.
CHAPTER 3
16
FLUORESCENCE
In fl uorescence-based detection, users should take full advantage of the consistent output of the
xenon fl ash lamp. If speed is critical, the one-fl ash ‘fl ying mode’ can help read many plates very
quickly. For more routine endpoint assays where speed is not critical, the number of fl ashes
should be increased to minimize variation (Fig. 11) . The number of fl ashes needed for an assay
depend in part on the microplate reader used and different manufacturers will have different
recommendations. BMG LABTECH typically provides advice on the number of fl ashes needed for
different assays in its Application Notes.
1 5 10 20 50 100 200
8
7
6
5
4
3
2
1
0
Number of Flashes
%CV
1
0.1
0.01
0.001
Blank
Fig. 11: Infl uence of used number of fl ashes on signal stability
You can read more about how the number of fl ashes infl uence measurements in the following HowTo Note:
How does the number of fl ashes infl uence measurement results
KINETIC ASSAYS
It is useful to monitor the change in absorbance or fl uorescence signal over time. There are two
main categories of kinetic assays: fast (or fl ash) and slow. For both modes, it may be useful to
reduce the number of fl ashes for very distinct reasons.
In a fl ash kinetic assay, changes in signal can occur rapidly. By using fewer fl ashes per data
point, the number of collectable data points per second increases. For slow kinetic assays, there
is no specifi c limitation. However, fewer fl ashes may preserve the long-term performance of
the microplate reader. The fl ash lamp and relevant controlling systems are very robust. They
have the capacity to perform hundreds of millions of fl ashes without appreciable decrease in
performance. However, it may be prudent to reduce the number of fl ashes since the performance
is fi nite. This is especially true when there is very little time between plate read cycles.
17
B. GAIN AND PERFORMANCE
Any fl uorescence or luminescence detection
device has a sensor that turns light emitted by
a sample into current that can be measured
electronically and reported as a value. A
bright signal from a well invokes more current
output from a detector like a photomultiplier
tube (PMT) than a signal from a dim well. The
proper detection setting – called gain – is
related to how much current is produced by
each photon striking the sensor. Inappropriate
gain values negatively affect data quality,
assay window, and sensitivity.
A high gain setting results in a greater
amplifi cation of the light signal coming from
a well and thus often improves sensitivity.
A high gain setting and a very high signal
might reach a maximum value or saturate
the detector. Once the maximum amount
of current output from the PMT detector is
reached, it becomes impossible to know how
bright a signal might be. You want the gain
to be high enough to appropriately record all
data without reaching signal saturation.
Time
Fluorescence (RFU)
Enhanced Dynamic Range
Fig. 12: Gain graph: fl uorophore concentration versus signal
In some cases, having a gain higher than
is necessary to discriminate between your
samples, controls and blanks might also
increase background and noise with no
real benefi t. Conversely, a gain setting that
is too low might limit the dynamic range
of the assay, and it may not be possible to
discriminate between dim samples or they
may become indistinguishable from the
background.
In most cases, the best way to set the gain is
to perform a gain adjustment on the brightest
well in your plate, typically a high standard
or positive control. If that bright well is not
saturating or reaching a maximum, then all
other wells are likely to be within range as
well.
For kinetic tests where it cannot be known
where the maximum signal will be at the end
of the measurement, preparing a sample well
that is allowed to go to completion before your
test begins may be an appropriate way to set
the gain. This is often the case for a kinetic
test that will be measured over seconds,
minutes, or hours. Another method is to set
the “target value” for the gain setting to a low
percentage of the maximum. For instance, an
adjustment method with a target value of 10%
could be performed on blanks.
Some microplate readers have an advanced
gain feature like Enhanced Dynamic Range
(EDR). EDR or similar technologies offer
advantages for both assay performance and
usability (Fig. 12). Methods to extend the
dynamic range usually work by reading wells
at a range of high and low gains and using
the best gain for each well. By very quickly
18
measuring a well and automatically adjusting the gain during a measurement, users do not
need to regulate the gain manually beforehand, which makes the operation easier for beginner
and intermediate level users. By measuring dim wells at a high sensitivity setting and bright
wells at a low sensitivity setting within the same test, the dynamic range of the assay can be
signifi cantly increased while improving sensitivity and decreasing noise. You can read more
about how the use of an appropriate gain setting impacts results in the following HowTo Note:
How to optimize the gain setting of my microplate reader
C. BOTTOM VS TOP READING
Instances where you might need bottom or top reading are discussed in Chapter 1, part a, page
4 under clear microplates.
D. FOCUS HEIGHT
Except for absorbance and nephelometry, all measurement used in a microplate reader
are based on the amount of light produced in a well. The z-height focus feature available on
advanced readers allows you to identify where the maximum intensity of light production can
be detected for your assay. This location is dependent on the volume of sample in the well, the
nature of the sample itself as well as the type of plate used.
The fi rst time you run a test with new conditions, you will want to perform a focus adjustment.
An intensity curve is displayed upon completion of this adjustment (Fig. 13). From this curve the
effect of using an off-peak focus on your result should be clear. For some assays, as much as a
25% decrease in intensity can be observed in as little as 0.2 mm deviation from the peak.
A 0.1-mm resolution of the focal height ensures that the highest intensity possible is read for
your experiment. This is true whether you select top or bottom reading.
0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15
3.600
3.400
3.200
3.000
2.800
2.600
2.400
2.200
2.000
1.800
1.600
1.400
1.200
1.000
800
600
400
200
0
Focal height [nm]
Measurement Value
z=11.6 mm
Fig. 13: Signal intensity
measured at different focal
heights
You can read more about
how the optimal focal
height impacts results in
the following HowTo Note:
How to optimize the focal
height of your microplate
reader measurements
19
E. POSITIONING DELAYS AND SETTLING TIME
For the most part, samples in a microplate reader are liquids.
Therefore, the movement of the plate within the reader can cause
liquids within a well to be in motion after the movement of the plate
between two wells is complete. Positioning delays help to ensure that
liquid movement in the well has ceased. The delay comprises a brief
pause upon completion of plate movement before a reading of each
well is performed.
How long to wait? The waiting time depends mainly on the type of
plate. In particular, the number of wells and the volume of sample.
Microplates can have as few as 6 or as many as 3456 wells. This
means widely different surface areas can be encompassed by each
well. More surface area allows for more freedom of movement of the
liquid. In addition, larger wells have higher volumes. The greater the
surface area and volume, the longer the waiting time needed before
reading the well. BMG LABTECH’s software takes this into account.
Each time you change your plate format the settling time is updated.
For example, a 6-well plate will have a suggested settling time of 1
second if precision settings are used; this value is 0.1 seconds for
384-well plates.
F. SHAKING AND INJECTION
Many assays require other actions from a microplate reader beyond
detection of a signal. The ability to shake and add liquids to your
samples are two functions that have become standard on microplate
readers. How these capabilities are implemented depends on what you
are trying to achieve with your assay.
SHAKING
Microplates can have from 6 up to 3456 wells. The typical volume
used decreases as the well number increases. In general, a higher
intensity of shaking is needed to move the liquid in a plate with many
wells and a low volume. Typically, very low volumes in small wells
require high speed shaking for any mixing to occur at all. The forces
experienced by ten microliters in a low-volume 384-well plate are
dominated by surface tension. Shaking speeds of more than 600 rpm
QUICK TIP
How long should you wait
after the movement of a
microplate in a reader
before making a
measurement? The
waiting time depends
mainly on the type of
microplate. A 6-well plate
may have a suggested
settling time of 1 second.
This value is 0.1 seconds
for a 384-well plate. An
advanced microplate
reader will automatically
update the settling time
each time the plate format
is changed.
20
may be important to ensure aeration, mixing
or suspension. Similarly, cell-culture plates
with volumes over 400 µl are likely to receive
suffi cient mixing at low shaking speeds around
200 rpm.
Well shape also affects mixing. Square wells
have corners that defl ect the movement
of liquid and can be useful in assays
where turbulence is needed. The greatest
displacement of fl uid and the most traditional
mixing situation is achieved in a round well
using round or orbital shake orientation.
Round bottom wells may sometimes be used
when the volumes are low or variable and
linear shaking can sometimes be useful to
take advantage of the “sloshing” effect that the
round bottoms permit.
Quality plate readers will always have
multiple shaking options that permit users to
implement clearly defi ned recommendations
for these parameters. If you are considering
adding shaking to your assay, make sure it
is necessary. Shaking a sample that is well
mixed and does not benefi t from additional
shaking may disturb the fl uid surface, produce
air bubbles, or add variability to the meniscus
that increases assay variability with little
benefi t. Next, decide if you want a relatively
aggressive force imparted by linear shaking or
the gentler mixing that results from orbital or
double orbital shaking. This is a good starting
point to test whether you are shaking in a way
that leads to reliable results for your assay.
21
SHAKING FOR SIMPLE MIXING
Most assays contain different components, and the assays work best when they are equally
distributed within the well. Similarly, some assays contain components that could precipitate
with time. In these cases, brief shaking before measurement is suffi cient. If the purpose is to
redistribute a precipitate that occurs over time, a shaking step is needed before each plate reading.
SHAKING FOR MICROBIAL GROWTH
Microbes like bacteria are typically grown in an incubator with an orbital shaker. This provides
effective mixing of the increasing number of bacteria and maintenance of aerobic conditions.
Growth of bacteria can be measured with an absorbance plate reader through detection of
absorbance at 600 nm. If shaking is performed between measurement time points, oxygen
penetration within the medium remains uniform while cell numbers increase.
The measurement of light scattering by particles using the absorbance mode may give different
results from one instrument to another. The reason is that different instruments use different
light beams and the detector is positioned at different distances from the sample. For a light beam
being scattered by a microorganism, a nearby detector still captures the light, while a detector
positioned further away does not (Fig. 14). Therefore, choosing one instrument
series for microbial growth measurements by OD600 is recommended.
SHAKING FOR MONITORING PROTEIN AGGREGATION
In most cases this refers to assays that use thiofl avin T, a fl uorophore whose fl uorescent signal
increases when it associates with beta sheet-rich structures like in amyloid fi brils. Such amyloid
aggregates are observed in several neurodegenerative disorders. Samples suspected of containing
misfolded prions are added to wells containing normally folded protein, which leads to misfolding
and aggregation along with an increase in fl uorescence in wells that contain prions. Optimization
of shaking speed and temperature is important for performance in tests like these. The rate at
which cyclical amplifi cation of misfolded protein proceeds is higher at higher temperatures but
may result in more false positives. Shaking at moderate to high speeds — up to 1100 rpm —
followed by a brief rest interval is important and impacts the rate of misfolding. Adjusting the
shaking speed and temperature parameters are important optimization steps to ensure that
aggregation is proceeding at a relevant rate.
Detection distant from the sample
Detection close to the sample
Microorganism scatters light
Light beam
Fig. 14: Different detection instruments give different results. Illustrated is the
infl uence of detector position: detectors of the same size placed closer to the
sample detect scattering while no signal is detected further away.
22
G. INJECTION
Many assays require the addition of a reagent before detection is possible. These types of assay
benefi t from and sometimes require the use of on-board reagent injectors (Fig. 15). These
injectors provide consistent, reproducible dispensing that outperforms most hand pipetting.
In some cases, the addition of a reagent causes a rapid response. Here it is an advantage to
read at the time of injection. This feature ensures that you will not miss any part of a fast kinetic
response, such as that observed in some calcium-release, membrane potential, protein-protein
interaction, or enzyme assays.
Two reagent injectors can be used when study of the effects of activation and subsequent
inhibition of a signal are important. Alternatively, one injector can add varying volumes of
reagents while the second injection adds a complementary volume of buffer so that the fi nal
volume is identical in each well. One can also monitor the cumulative effect of two compounds.
Another frequently used example for two reagent injections is a dual luciferase assay to study
gene expression. Here the fi rst injection initiates a response, and the second injection quenches
the fi rst activity and begins a second activity.
In addition to injection of two reagents, there are a wide variety of injection options. Different
wells can receive different volumes of reagents at different times. Reagents can be added at
multiple time points throughout an experiment.
Fig. 15: Injectors enable
reagent delivery to any plate
format
INJECTION SPEED
Typically, the default injection speed set by the manufacturer is suitable for most purposes. The
main exception is when you are injecting into a well that contains adherent cells. In this case,
you may need to decrease injection speed to leave the cells undisturbed. An injection speed that
is too fast may cause detachment of the cells in the center of the well, where it is most likely to
23
affect measurement. A similar scenario is the
use of reagents that tend to form bubbles. In
this case, the speed should be decreased to a
minimum of 50 μl/s.
SHAKING FOLLOWING INJECTION
In many cases, the act of injecting a
reagent is suffi cient to fully mix the
injected reagent and the sample in the well. A
shake after addition of reagents is most useful
after an injection step if your assay requires
a slow injection speed and if only a small
volume of reagent was added to the well. If
there is a signifi cant difference in viscosity
between the reagent being injected and the
sample in the well, volumes less than 20 µl
may not mix properly without a shaking step.
H. KINETICS: PLATE VERSUS WELL
MODE
The type of reaction you are measuring will
determine the way reagent injections and
measurements are timed. A reaction that is
over quickly cannot be initiated outside of the
plate reader unless it can be measured as an
endpoint test. If the signal is transient or if
the information about rate of progress of the
reaction or initiation is important, an injection
must occur close to the time of measurement.
Even if an injection is performed inside a plate
reader, it may still require a rapid well-bywell detection program to see good results.
If the half-life of an enzyme reaction is a few
seconds, most interesting rate kinetics will
be missed if a well is injected and immediate
detection does not ensue.
Well mode is used when you want to measure
one well of the microplate multiple times
(up to 100 per second) and subsequently
move on to the next well and repeat the
same steps. This is usually accompanied
by an injection to initiate the reaction
before measurement begins. This mode is
suitable for fast kinetic experiments, such
as calcium fl ux measurements, enzyme
kinetics and ion channel activity — tests
where data would perhaps be completely
missed if measurement did not immediately
follow the dispensing of reagents. Another
benefi t of well-mode injection protocols is
that injection and measurement occur at
the exact same time relative to each other in
every well. If a whole plate is dispensed and
then the measurement begins, there may be
a differential in the time between injection
and measurement that varies from the fi rst
well to the fi nal well. Well-mode tests may
also be benefi cial when extremely dense data
collection rates are important. If it takes
30 s to read a whole plate, it means each well
will only be measured about four times in
two minutes. Compared to plate mode, well
mode enables the acquisition of up to 100 data
points in every 2 minutes. Conversely, using
well-mode kinetic detection for slow kinetics
is not recommended as it will artifi cially
prolong the detection time for the whole plate.
Plate mode is used when you want to measure
each well of the microplate once (like endpoint
mode) and then repeat the process. This mode
is suitable for slow kinetic experiments, such
as measuring bacteria and yeast growth, cell
migration and invasion, and slower enzyme
kinetics. If a reaction proceeds for several
minutes, most quality plate readers can
24
read many data points in that time frame. By
injecting the whole plate and then measuring
the whole plate multiple times, it is possible
to collect the largest amount of data most
effi ciently. If a reaction takes fi ve minutes
to go to completion, it will take about eight
hours to take 96 individual fi ve-minute
measurements in well mode. Therefore,
reading in plate mode is appropriate for most
assays that do not require extremely rapid
data collection.
The choice of well mode or plate mode
therefore depends on the type of assay and
the speed of reaction you want to monitor.
I. WELL SCAN OPTIONS
Several variables can lead to a nonhomogenous signal distribution across the
well surface. Notably, this includes the use
of adherent cells with intracellular signals or
the use of samples that form a precipitate or
clots. Microbial assays are non-homogenous.
Likewise fl occulating bacteria or yeast
as well as biofi lm-forming bacteria are
unevenly distributed. In these situations, a
more complete picture of what is occurring
throughout the well is needed as a regular
measurement in the center of the well may
lead to misleading results or to no signal at
all. Different well-scanning options (Fig, 16)
are useful in this context as they cover a wider
surface of the well and can compensate for
the heterogeneous signal distribution. These
options should be considered for use with
fl uorescence, luminescence, and absorbance
measurements. You can read more about
how to capture the signal from all cells in
microplate reader measurements in the
following HowTo Note: How to reduce data
variability in heterogenous cell samples
MATRIX SCANNING
In matrix scanning, the microplate reader can
take multiple measurements in each well. The
software displays each scan point graphically
and creates a map for each well. The user can
easily remove individual scan points or entire
sections of each map.
ORBITAL AND SPIRAL SCANS
Another possible way to measure the contents
of a well is using orbital and spiral averaging
features (Fig.16). Using these modes, the
plate reader takes several measurements for
each well on a defi ned orbit, collects the data,
and calculates an average.
Fig. 16: Orbital averaging, spiral averaging and matrix scanning
25
J. PATH LENGTH CORRECTION IN ABSORBANCE ASSAYS
Most absorbance assays were fi rst done in a cuvette with a 1 cm
pathlength. Performing this type of assay one at a time is tedious. As
these assays have moved into microplates, the effective pathlength
has become smaller and is determined by the amount of liquid in the
well. In an absorbance assay, the relationship between absorbance and
concentration is defi ned by the Beer-Lambert law. The Beer-Lambert
law states that the amount of light absorbed is proportional to the
concentration of the light-absorbing molecule and is expressed by
A = ε · c · l. It is the pathlength l that is different in a microplate versus
a conventional cuvette.
A pathlength correction is used any time you want to perform a
concentration determination of a substance based on the BeerLambert law. The default approach now is to use the water peak in the
infrared part of the spectrum to perform this correction. This feature
of aqueous solutions can provide a precise pathlength for each well,
reducing the potential effects of meniscus on the fi nal calculations.
A pathlength correction is not always needed if, for example, your
analysis includes the use of a standard curve. In some cases, the use
of the water peak-based correction is not appropriate. This is the case
for the OD600 assay. If using the water peak is not suitable, users can
manually input the fi ll volume of the samples on their microplate and
the pathlength can be estimated for all samples based on that volume
and well geometry.
It is important to be aware of the implications of pathlength correction
and be able to make necessary adjustments. Several factors can
contribute to misleading changes to pathlength correction including
differential evaporation for samples and controls and effects due to
bubbles. If a blank has a different meniscus compared to the sample,
for example due to the presence of sample protein or nucleic acid
in one well but not in the control, then this can also pose problems
for pathlength correction (Fig. 17). In this case, the menisci can be
adjusted by the addition of other molecules including detergents that
do not interfere with subsequent measurements. Interference from
other compounds may also be problematic for pathlength correction.
QUICK TIP
A blank may have a
different meniscus
compared to the sample.
This may be due to the
presence of sample
protein or nucleic acid, for
example, in one well but
not in the control and can
pose problems for
pathlength correction.
If this is the case, the
menisci can be adjusted by
the addition of detergents
or other molecules that
do not interfere with
subsequent
measurements.
26
Interference may arise for example due to the presence of nonaqueous substances, the presence of compounds that give differential
absorbance at the crucial wavelengths of 1000 and 900 nm (for example
from chlorophylls or carotenoids), or to turbidity effects. In some cases,
the appropriate dilution will minimize the impact of interference or
turbidity on pathlength correction. In other cases, selection of a more
appropriate reference may alleviate any discrepancies. 0.4 cm
0.5 cm
Fig. 17: The meniscus of a liquid surface changes the path length in microplate wells.
27
PERFORMANCE INDICATORS
A. LIMIT OF DETECTION
The limit of detection (LOD) is the lowest
signal or the smallest measurement of a
concentration that can be determined with a
specifi ed precision or reproducibility. Limit
of detection and sensitivity are often used
interchangeably but are not the same. The
LOD is a function of both sensitivity (signal
strength) and signal stability (the amount
of signal noise). Detection limits are direct
indicators of the detection performance of a
microplate reader. Sensitivity alone cannot be
related to the detection performance of the
system.
As there is no standardized way to determine
LODs on plate readers, LOD calculation
may vary among manufacturers. A typical
0 1 2
330
320
310
300
290
280
270
260
250
68,27%
95,45%
Blank + 3xSD
Gaussian distribution
IUPAC calculation:
Blank:
SD:
3xSD:
M (slope):
3xSD/m(slope)=23.1/23.22E12
LOD = 0.99 pM
260 RFU
7.7
23.1
23.22E12
Blank + 2xSD
99,73%
Blank + 1xSD
Blank
LOD calculation
Fig. 18: Calculation of LOD based on the blank and 3 x SD
approach involves measuring replicates of a
blank sample to estimate the detection limit,
determining the mean value and standard
deviation (SD), and calculating LOD as the
mean at a defi ned confi dence level (typically
at a SD of 3, see Fig. 18). This is a simple and
quick method. If a sample is present, it is
assumed that it will produce a signal greater
than the noise in the absence of sample. The
second approach is more direct and involves
analysis of small but known concentrations
of the sample. The data are used to compare
the response of the blank sample and the
low concentration samples. This comparison
helps determine what amount of sample is
needed to distinguish its presence from its
absence.
CHAPTER 4
28
B. UNDERSTANDING SIGNAL-TO-BLANK, SIGNAL-TO-NOISE AND DYNAMIC
RANGE
Signal-to-blank and signal-to-noise are two assay parameters used to evaluate assay
performance. They are often used to determine the dynamic range of an assay, namely the
largest and smallest value that a certain quantity can assume.
The signal-to-blank ratio measures the signal of replicate samples and expresses them relative
to the values of replicates of the blank (see equation 1). The signal-to-noise ratio provides a
measure of the signal quality (see equation 2). It quantifi es and compares the level of a signal
relative to the average level in the background noise, which includes any signal from sources
other than the target signal source. The signal-to-noise ratio can vary from measurement to
measurement.
S/B = µsignal/µbackground Equation 1
µsignal is the average of the values over the replicates for which the S/B value is calculated
µbackground is the average of the values over the replicates of the blank
S/N = |µsignal - µbackground|/(σsignal2 + σbackground2) Equation 2
µsignal is the average of the values over the replicates for which the S/N value is calculated
σsignal is the standard deviation over the replicates for which the S/N value is calculated
µbackground is the average of the values over the replicates of the noise (blank)
σbackground is the standard deviation over the replicates of the noise (blank)
A researcher might encounter a wide range of signal intensities when using a microplate reader.
The lowest sample concentration might for example be 1 µM and the highest concentration
might be 1,000 µM. In this case, the dynamic range spans three decades of concentrations.
In some cases, the upper limit might be a million times higher than the lowest value, a
dynamic range of six decades. Signal intensities can range from very dim to bright over such a
concentration range. The dynamic range is related to the gain setting. An optimal gain setting
will ensure the largest possible dynamic range for an assay.
The signal-to-noise ratio and the dynamic range are criteria commonly cited as indicators of
detection performance.
The CLARIOstar® Plus and VANTAstar® plate readers have been specifi cally designed to offer
the largest possible dynamic range. The Extended Dynamic Range (EDR) can be applied to
any wavelength in fl uorescence intensity and luminescence measurements. Results can be
measured over eight concentration decades with no manual intervention.
29
C. Z-PRIME (Z´)
In some cases, signal-to-blank and signal-to-noise ratios are not
suffi cient to determine the quality of detection for an assay. When this is
the case, the Z-prime statistic, which considers four parameters, can be
used as a measure of assay quality.
The Z-prime statistic is distinct from the Z-factor. The Z-factor, which
is suitable for studying the quality or performance of high-throughput
assays, is itself defi ned as follows:
Z = 1- (3σs + 3σc)/ |µs- µc| Equation 3
where µs and µc are the means of the signals for the sample and
control, respectively, and σs and σc are the standard deviations of the
signals of the sample and control, respectively. The Z-factor considers
the dynamic range of the assay signal, the data variation due to the
sample measurement, and the data variation due to the reference
control measurement. The Z-factor thus defi nes a characteristic
parameter of the capability of hit identifi cation for each given assay at
the defi ned screening conditions. In contrast, the Z-prime statistic is
a characteristic parameter for the quality of the assay itself without
the use of test compounds. The Z-prime statistic is appropriate for
evaluating overall assay quality whereas the Z-factor reveals the quality
of a confi gured assay for a particular high-throughput screening. The
Z-prime statistic is therefore relevant to any assay and is not restricted
to high-throughput screening.
The Z-prime statistic considers the means (µ) and the standard
deviations (σ) of the signals for the positive and negative controls (see
equation 4).
Z’ = 1- (3σc+ + 3σc-)/ |µc+ - µc-| Equation 4
In practice, the assay in question should fi rst be optimized according
to the Z-prime statistic for conditions such as reagents, procedure,
kinetics, instrument, and other factors other than those related to the
test compounds. This ensures the assay format will have suffi cient
dynamic range and signal variability and that it will provide useful data.
If the Z-prime statistic is small (negative or close to zero), it usually
indicates that the assay conditions have not been optimized or that the
QUICK TIP
Z-prime (Z’) and Z-factor
are two distinct ways to
determine the quality of
an assay. Use Z-prime
if you want to evaluate
the overall quality of any
assay. Use the Z-factor to
determine the quality of a
specifi c confi gured
assay for a particular
high-throughput
screening. The Z-prime
statistic is relevant to any
assay and is not restricted
to high-throughput
screening. The Z-prime
statistic is a characteristic
parameter for the
quality of the assay itself
without the use of test
compounds.
30
assay format is not feasible for generating useful data. After this step,
the properties of the compound library used for the screen should be
considered and the Z-values determined. Z-values between 0.5 and
1.0 indicate excellent performance. Values between 0 and 0.5 may be
acceptable. Values less than zero indicate the assay is not useable.
The Z-factor and Z-prime statistic have other uses beyond assay
development and the validation of high-throughput assays on a
specifi c instrument. They can also be used for example to compare the
performance of different instruments for data quality under the same
conditions.
BMG LABTECH offers the MARS data analysis software for automated
data reduction. MARS can easily be used to determine Z-factor and
Z-prime statistics as well as many other quality control parameters
including signal-to-noise and signal-to-blank ratios.
31
FURTHER READING
Armbruster DA, Pry T. Limit of blank, limit of detection and limit of quantitation. Clinical
Biochemistry Reviews 2008 Aug;29 Suppl 1(Suppl 1): S49-52.
Auld DS et al. Microplate Selection and Recommended Practices in High-throughput Screening
and Quantitative Biology. 2020 Jun 1. In: Markossian S et al., editors. Assay Guidance Manual
[Internet]. Bethesda (MD): Eli Lilly & Company and the National Center for Advancing
Translational Sciences; 2004–.
Auld DS, Inglese J. Interferences with Luciferase Reporter Enzymes. 2016 Jul 1 [updated 2018
Jul 1]. In: Markossian S et al., editors. Assay Guidance Manual [Internet]. Bethesda (MD): Eli Lilly
& Company and the National Center for Advancing Translational Sciences; 2004–.
Simeonov A, Davis MI. Interference with Fluorescence and Absorbance. 2015 Dec 7 [updated 2018
Jul 1]. In: Markossian S et al., editors. Assay Guidance Manual [Internet]. Bethesda (MD): Eli Lilly
& Company and the National Center for Advancing Translational Sciences; 2004–.
Zhang JH, et al. A Simple Statistical Parameter for Use in Evaluation and Validation of High
Throughput Screening Assays. Journal of Biomolecular Screening. 1999;4(2):67-73. doi:
10.1177/108705719900400206.
You can learn more about the optimal reader settings for your measurements in our HowTo
Notes
You can also learn more about the detection methods mentioned here
AlphaScreen®, AlphaLISA®, DELFIA® and Homogeneous Time-Resolved Fluorescence (HTRF®)
are registered trademarks of PerkinElmer, Inc; THUNDER™ is a registered trademark of
BioAuxilium
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