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The word luminescence is composed of “lumin” (Latin for light) and the suffix “-escence” (used for processes or changes). Luminescence is hence a process in which light is released. Per definition, luminescent light stems from cold sources and is differentiated from light emission from heated sources such as hot iron or a burning candle1. The light is emitted in course of energy conversion from a broad range of energy sources. The process turns invisible energy into visible radiation and is often used: by nature for a defense, for LEDs on screens, or for analysis purposes. This page focuses on the use of luminescence methods in life sciences, explains their physical background, provides information on measuring luminescence, and presents common luminescence microplate assays.
Physical background of luminescence
Luminescence is the emission of light through energy conversion. Its application in life sciences primarily uses two energy sources: chemical energy, which leads to chemiluminescence, and light energy, which leads to photoluminescence. The latter is also the basis of fluorescence.
The energy of one of the two sources is absorbed by a molecule and brings its electrons to a higher energy level (Fig. 1). As this level is unstable, the electrons fall back from the excited to the ground state. When falling back, they release energy in form of vibrational energy, heat, and/or light (photons). The latter one is what we see as luminescent emission2.
In life sciences, “fluorescence” is commonly used when talking about photoluminescence while “luminescence” typically refers to chemiluminescence. This simplification is supported by the difference in light detection: fluorescence (photoluminescence) requires excitation light whereas luminescence (chemiluminescence) does not (Fig. 2). This text adheres to this common nomenclature and concentrates on chemiluminescence, following referred to as “luminescence”.
Luminescence in life science applications
In chemiluminescence, a substrate reacts in an electronically excited state. The excited product or intermediate falls into its ground state by emitting a photon, hence being luminescent. Prime example of a chemiluminescent reaction is the reaction of luminol in the presence of hydrogen peroxide (Fig. 3). In an alkaline environment, luminol exists in a form (Dianion) that reacts with molecular oxygen O2. The oxidized intermediate then reacts to an electronically excited 3-Aminophthalic acid (3-APA). The molecule then drops to its ground state and emits light. This simple principle of producing an excited state molecule by a chemical reaction that then falls into its ground state by emitting light is also the basis for enhanced chemiluminescence and bioluminescence. Thus, both are chemiluminescent processes as well.
b) Enhanced chemiluminescence (ECL)
As the name indicates, ECL uses enhancers in their chemiluminescent reaction. ECL mainly employs luminol in conjunction with hydrogen peroxide. However, the oxidation of luminol is catalyzed by an enzyme: horseradish peroxidase (HRP). Additionally, ECL reactions contain chemicals that enhance the light emission such as p-coumaric acid or 4-iodophenylboronic acid. The use of an enzyme to catalyze the chemical reaction enables ECL to be used in enzyme-linked reactions. Its major application is for immunoblots: proteins are separated by size, transferred onto a membrane, and detected by ECL. The proteins of interest immobilized on the membrane are bound by a protein-specific antibody. A second, horseradish peroxidase coupled antibody is used to link the protein-antibody with the enzyme. Thanks to enzymes and enhancers, a bright luminescent signal is produced only where the protein is found.
The same principle is employed for microplate-based quantification of biomolecules. ELISA assays with luminescent readout have a higher sensitivity and are based on enhanced chemiluminescence. The protein of interest is immobilized onto the microplate well and specific antibodies, HRP-coupled secondary antibody, substrate, and enhancer produce light which increases along with protein concentration.
Bioluminescence is a form of chemiluminescence that occurs in living organisms. The term also covers luminescent reactions using enzymes and substrates originally derived from living organisms, even though being used outside the organism or bio-engineered to provide improved characteristics. Bioluminescence is used by various organisms for different biological purposes. Fireflies use their ability to glow to attract mates. Jellyfishes such as Aequorea victoria are thought to use light production to ward off biological enemies. Bacteria such as the marine Vibrio fischeri start to emit light as they become dense. The process is called quorum sensing and allows the population to communicate and coordinate.
How do these organisms manage to emit luminescence? They produce an enzyme called luciferase. It catalyzes the oxidation of luciferins, a substrate that is expressed along with the luciferase. During enzyme-mediated substrate conversion, light is emitted. Different organisms produce different enzymes and substrates. Furthermore, the light-emitting reactions require different co-factors and result in varying emission wavelengths. Figure 4 shows three luciferase reactions of widely used luciferases in life science assays.
d) Bioluminescence resonance energy transfer (BRET)
Resonance energy transfer describes the transfer of energy from an electronically excited donor molecule to an acceptor fluorophore. The process excites the acceptor fluorophore which in turn emits light. In case, the donor energy is generated by bioluminescence (luciferin+luciferase), the process is called bioluminescence resonance energy transfer (BRET). Various conditions must be met to allow energy transfer: the emission spectrum of the donor molecule needs to overlap with the excitation spectrum of the acceptor fluorophore. Additionally, donor and acceptor molecule need to be in proximity (typically 1-10 nm), as the transfer decreases with distance. Consequently, BRET is commonly used to measure the binding of two biomolecules. The output of BRET is the intensity of the acceptor fluorophore related to the intensity of the donor luminescence, known as BRET ratio. BRET1, BRET2 and NanoBRET use different combinations of luciferases and acceptor fluorophores as shown in table 1.
Table 1 – Types of BRET
|Name||Donor luciferase||Substrate||Donor emission||Acceptor fluorophore||Acceptor emission|
|BRET 2||Renilla luciferase||DeepBlueC||400-450 nm||GFP||500-540 nm|
|NanoBRET||NanoLuciferase||Furimazine||420-500 nm||NanoBRET 618||550-675 nm|
|AlexaFluor 633||600-700 nm|
e) Flash luminescence
Luminescence-based assays for life science applications often state whether they use flash or glow luminescence. The difference between the two is the duration of signal emission. Flash luminescence assays emit light for a maximum of a couple of seconds. The light intensity of the light flash produced by these assays is typically recorded directly from the start to the end of the reaction. This implies, flash luminescence reactions need to be started by the addition of a reaction starter. On microplate readers, reagent injectors are needed to automatically start the reaction and record the signal simultaneously. Popular flash luminescence assays are the Dual-Luciferase Reporter™ technology or SPARCL assays.
f) Glow luminescence
In contrast to flash luminescence, glow luminescence emits stable signals for up to hours. The assays do not require automatic dispensing and can be read over a longer time span. Researchers, as well as kit providers, tend to prefer glow luminescence assays as they are easier to handle and detect. One of the most prominent representatives of glow-luminescence is the viability test CellTiterGlo®.
Apart from ECL-detection of western blots, luminescence tests typically employ microplates. These contain between 6 or 1536 wells in each of which reactions take place that are quantified using a microplate reader. This section explains how luminescence in microplates is measured.
Components for luminescence detection
Luminescence is easier to detect than fluorescent or colorimetric assays as no excitation is required. The minimum components needed to measure luminescence are a lens to collect the light and a detector (Fig. 5). A wavelength selection tool is required to measure BRET and light guides may be necessary for specific detection constructions.
a) Photomultiplier tube (PMT)
Photomultiplier tubes (PMTs) serve as detectors in luminescence detection. PMTs differ in their sensitivity (the lowest detectable signal), their noise, and their capability to measure other detection modes. Many luminescent measurement devices come with a universal PMT that reads luminescence as well as other detection modes. Their advantages are low space and cost requirements despite providing high sensitivity. The possibility to measure very low signals is provided by optimized optical systems. Such an optimized system is the free-air optical path combined with a luminescence plus optic module found in PHERAstar FSX microplate readers.
Dedicated PMTs for luminescence measurements provide either a lower noise or the use of a photon-counting principle. While both dedicated PMTs can detect low signals as well, a photon counting system for luminescence measurements is limited in detecting higher signals (Fig. 6).b) Light guides
Ideally, the detector is placed directly above the well. If this is not possible, the light needs to be guided from the well to the detector. This task can either be performed by lenses and mirrors or by a light guide. The guide absorbs some of the luminescence emission which decreases sensitivity compared to a free air optical path. The latter can be found on VANTAstar, CLARIOstar Plus, and PHERAstar FSX microplate readers.
c) Wavelength selection
Some applications require to detect only the light at a specific wavelength. This is accomplished by optical filters or monochromators placed in the light path. BRET measurements need wavelength selection as two signals come from one and the same sample. One filter is transmissive for the light coming from the donor luciferase, and a second filter for the light coming from the acceptor fluorophore. This allows distinguishing between light coming from the luciferase and light coming from the fluorophore.
Conventional grating-based monochromators play a subordinate role in filtered luminescence and BRET measurement because of limited sensitivity. This is due to scattering effects and narrow bandwidths. However, a monochromator technology using Linear Variable Filters (LVF) provides the sensitivity needed for luminescence detection. An LVF-based monochromator exhibits filter-like light transmission and provides bandwidths up to 100 nm. This assures enough luminescent signal reaches the detector for filtered luminescence such as BRET (Fig. 7).
Measurement time interval, integration time, or acquisition time
A luminescent signal lasts a second or longer and is therefore clearly different from the fluorescence that decays in a nanosecond range. Accordingly, the signal is recorded over a time period. Typical times for measuring a single well range between 0.1 - 1 s. This time has different names: measurement time, integration time, and measurement interval time. The chosen integration time depends on several aspects that have to be balanced against each other such as signal intensity and total time to read a plate.
Luminescence measurements are comparatively easy as they typically have fewer settings to modify in microplate readers. However, there are instrument-related and general variables that impact luminescence measurements and data quality.
As stated above, the acquisition time depends on several factors. The following aspects should be considered when choosing the integration time:
a) Signal intensity
Most assays emit enough light to record the signal for half a second or even down to 0.02 seconds. Only very rarely it is necessary to increase the integration time up to a few seconds in order to detect differences in luminescence signals.
b) Flash and glow luminescence
Glow luminescence signals are typically acquired for 0.1 - 1 s as they emit a stable signal. Therefore, the total read time, as well as the timing and duration of signal measurement, play minor roles. This is different in flash luminescence assays. For these, it is important to collect the complete signal emission from start to end. This can be accomplished by integration times of several seconds, depending on how long signal emission occurs. If it is important to know what the signal curve looks like and what the maximum emission of a flash luminescence reaction is, multiple measurements at very low integration times are necessary. For instance, if a reaction takes approximately one second, 50 measurements of 0.02 s can be performed to cover one second and monitor how the signal develops and decays.
c) Temporal resolution
This point is of relevance only for kinetic measurements. In case a cell-based or biochemical reaction is monitored with a luminescent assay, the time course of the reaction dictates the integration time. An example illustrates this: in calcium assays responses occur in a range of 20 seconds after stimulation and can be displayed by 10 measurements every 2 seconds. If only one well is measured, an integration time of 1 or 2 s can be chosen and results in an appropriate temporal resolution. However, if more wells need to be read with the same temporal resolution (measurement point for each well every 2 seconds), the integration time needs to be reduced in order to measure the remaining wells and avoid loss of data.
d) Total read time
As the integration time is spent on each well, it contributes largely to the total read time of a plate. Increasing the integration time on each well by only 0.2 s, increases the overall read time of a 96-well plate by 20 s, of a 384 well plate by more than a minute. Therefore, it is important to use low integration times in high-density plates (384 or 1536 well plates) and in high throughput applications where thousands of plates need to be measured in a day.
The gain in luminescence measurements can be regarded as an amplification factor that moves a fixed dynamic range window along the concentration curve of the sample. Low signal intensities require higher gains, whereas intense luminescent signals require lower gains. Usually, the gain is set to have a maximum measurement output on the sample with the expected highest intensity. This is done in order to have the largest possible dynamic window between the highest and the lowest measurement values. Hence, if positive controls with a maximum signal are used along with unknown samples, the gain can be adjusted on them.
The possibility to measure with different gains resulting in a large dynamic range allows for measuring very low luminescent signals and bright light emissions with only one instrument.
Gain settings are not required for all luminescent microplate readers, depending on the detector and automation of the gain adjustment process.
Modern luminescence plate readers automatically accomplish gain adjustments. This not only takes away this responsibility from the researcher but also assures to measurement a broad dynamic range. Such instruments are the CLARIOstar® Plus and the VANTAstarTM with its Enhanced Dynamic Range technology.
Plate color for luminescence measurements
White plates are best suited for luminescence measurements as they reflect the signal instead of absorbing it. Detailed information on the plate choice is found in our blog post: Which is the best microplate for my assay.
Importantly, there is one exception to the rule: if a luminescent assay is combined in the same well with a fluorescent assay, a black plate is preferred. The reason is that fluorescence measured in a white plate leads to very unstable values and very high background signal because of the reflection of excitation light, making fluorescence detection very weak in white plates. Measuring luminescence in black plates also reduces the assay window, but as the background is highly reduced and measurements exhibit lower deviations, the measurements still give satisfying results.
What is cross-talk and how to circumvent it?
Cross-talk is a phenomenon only affecting luminescence measurements. Cross-talk is the light from any but the measured well that is unspecifically measured by the detector and modifies the signal of the actual measured well.
Since the light produced in a luminescent reaction is diffuse, it may not only shine directly above the well but also to neighboring wells and directly to the site of detection, even though another well is measured. This leads to biased signals, higher signal variations, and to a lower overall sensitivity. The problem can be solved by different means, depending on the reaction type and on the diffusion of the luminescent signal.
For flash luminescence assays that only shine for a few seconds, it is sufficient to measure the plate in a different order. As the signal decays quickly, the only wells affected by cross-talk are the adjacent ones when measured directly after each other. If a distant well instead of an adjacent one is measured the cross-talk is drastically diminished as the decaying signal is not detectable at the distant site. For this purpose, BMG LABTECH microplate readers offer an interlaced reading mode. The interlaced mode measures the first well, skips one well, and jumps to the third. Every other well is measured until the end of the plate is reached and then the omitted zones are measured. This allows the first measured signals to decay until their direct neighbors are measured.
In contrast, glow luminescence tests emit stable signals for hours and require other strategies to eliminate cross-talk. Undesired signals can reach the detection site in two ways: above the plate and through the wall of the wells (Fig. 8). Both ways need to be approached differently.
a) Physical blocking
Apertures physically block the unwanted light that shines over the wells into the detector. An aperture is a black spoon-shaped accessory with a hole in it that is placed above the microplate (Fig. 8). Through the hole, the light of the well of interest reaches the detector, while all light coming from its surrounding is physically blocked. BMG LABTECHs PHERAstar® FSX, as well as VANTAstarTM and CLARIOstar® Plus, come with apertures for improved luminescence detection.
Light can shine through the plastic wall of a well, even in white microplates. The type of microplate significantly affects cross-talk shining through the walls of a plate. As a rule of thumb, higher density plates (e.g. 1536 well) display higher light leakage through the walls than lower density plates, as the walls are thinner. A second aspect is a well geometry: square wells have a higher through-the-wall-cross-talk because adjacent wells share a wall. Round wells do not share the wall and, hence, display lower cross-talk through the wall. Furthermore, the color of the plate influences cross-talk through microplate walls: the darker the plate the lower the cross-talk. Grey plates offer a compromise between cross-talk reduction and signal reflection.
If it is not possible to further optimize the plate, a mathematical cross-talk reduction can be applied to the acquired data. To this end, the bleed-through of signal to neighboring wells is determined and an algorithm corrects the luminescent data. An automatic determination and cross-talk correction is found in the BMG LABTECH plate readers PHERAstar® FSX, VANTAstarTM, and CLARIOstar® Plus.
Light emission from the plate
White plates have an intrinsic phosphorescence. This means that light is emitted from the plate itself after it was exposed to light. The light emitted by the plate is recorded along with the light emission from the reaction and therefore alters the measurement data. As the plate light emission also increases the blank and reduces the assay window, it is recommended to either prepare the plate in the dark or to leave the plate in the dark approximately 15 min before measuring.
Filters are mainly needed for BRET measurements. As luminescence often exhibits low light outputs, it is recommended to use filters with a broad bandwidth of 80 - 100 nm. Broader bandwidths result in more light coming through to the detector, which increases the sensitivity of the measurement.
Reporter assays use gene regulatory sequences combined with the genetic information for a reporter molecule to study either change in gene expression or modifications in the regulatory region itself. Luminescent reporter assays employ a luciferase as a reporter molecule. The genetic information for regulation and luciferase (reporter gene) needs to be introduced into cells which are then measured with and without experimental perturbation. If the reporter gene is active, the enzyme is transcribed and translated. In presence of a substrate, it is converted, and light is produced. This way, light emission reports on the activity of regulatory sequences.
Dual-luciferase reporter assay
The dual luciferase reporter assays (DLR™) differ from a “single” reporter assay as it adds a second reporter to the system. Next to the regulatory sequence of interest and an according to luciferase, a second luciferase controlled by a known control promoter serves as internal control.
The most popular luminescence-based viability assay employs firefly luciferases’ dependence on ATP. The activity of the enzyme increases with rising ATP levels and so does the light emission. Viable cells produce ATP, which after cell lysis fuels the luciferase. Accordingly, light emission correlates with cell number and viability. In addition to ATP-based endpoint viability assays, luminescent assays may measure cell viability in real-time. To this end, a pro-substrate, as well as a luciferase, are added to the cell culture. The pro-substrate is reduced only by viable cells and is then processed by the luciferase in a light-emitting reaction. This way, luminescence reports on viability changes in real-time. You can find out more about these assays in our scientific talk “Real-time cell health assays deliver better data with less effort”.
Receptors are common drug targets and their ligands are possible therapeutics. The binding of ligands to their receptors can be studied in cell-based assays using the BRET principle. The luciferase is expressed at the extracellular part of the receptor and a ligand is labelled with a suitable acceptor fluorophore. If the ligand binds its receptor, donor luciferase and acceptor fluorophore are close enough for energy transfer and for the BRET ratio to increase. The method further allows for the studying receptor binding of unlabeled compounds. Unlabeled ligands can compete for binding with the known ligand, labelled with a fluorophore. If the BRET ratio decreases in such a competitive setup, an unlabeled compound displaced the acceptor molecule leading to a loss of energy transfer (Fig. 9).
How these assays help to study receptor pharmacology is explained in our scientific talk “Real-time profiling of receptor pharmacology”.
The principle of BRET is also used to study interaction of two proteins. To this end, one of the interacting proteins is coupled to the luciferase, the other to an acceptor fluorophore. Upon interaction of both, energy transfer occurs leading to fluorophore emission and an increase in BRET ratio.
Luminescence microplate assays are often more sensitive than their counterparts based on other mechanisms. For instance, luminescent ELISAs are more sensitive than colorimetric ones and luminescent viability assays are more sensitive than absorbance-based assays. Furthermore, they offer solutions for a wide variety of biological questions. Due to its sensitive nature, it is highly used for high-throughput applications which typically work with low volumes and hence low amounts of analyte. It is however necessary to detect it with instrumentation that is just as sensitive as the method itself. An advantage for non-HTS use of luminescence is the availability of homogenous assays based on an add-and-measure principle which makes the assays easy and fast to process. Furthermore, a variety of real-time luinescent assays allow the quantification of cellular processes as they occur.