We are searching data for your request:
Upon completion, a link will appear to access the found materials.
We're considering organizing some interlaboratory work on calibrating luminescence reporters (e.g., luciferase), and one of the key questions I don't know the answer to is whether most plate readers can measure luminescence or not.
From a first principles perspective, any fluorescence plate reader ought to be able to measure luminescence as well - just don't turn on the excitation light source. Thus, I would expect that a typical fluorescence reader should also be able to measure luminescence.
On the other hand, searching online, I am swamped by manufacturers eager to sell me specialized luminescence readers that emphasize their sensitivity, which leads me to think that maybe typical fluorescence plate readers can't measure luminescence.
My question, then, is this: should I expect that a typical current fluorescence plate reader will also be able to measure luminescence, or is this a much more specialized capability?
As the other answers here say, technically plate readers which are capable of fluorescent measurements can also make luminescent readings, but the sensitivity may be low (https://www.biotek.com/resources/technical-notes/use-of-the-fluorescence-optics-to-measure-luminescent-reactions-in-high-well-density-microplates-measuring-luminescence-in-1536-well-microplates/).
What I haven't seen mentioned is that there is potentially another hardware limitation depending on what type of luminescence you're using. Broadly there are two types: glow and flash luminescence.
Glow luminescence is probably most similar to fluorescence in that it builds gradually and can remain stable over an extended period (think of a lightbulb being turned on and warming up). Flash luminescence, however, is much more transient as the name might suggest. Reactions which cause flash luminescence produce a short burst of light (which could last seconds to minutes) which is typically brighter than that produced by glow luminescence. (https://www.bmglabtech.com/what-is-the-difference-between-flash-and-glow-luminescence-assays/)
As some flash luminescence assays will only produce a signal for a few seconds after the reagent is added, it is important to have immediate measurement, which can be provided by using auto-injectors within the plate reader. (https://www.biotek.com/products/detection-flash-luminescence-technology.html) These may or may not be included in a plate reader dedicated to fluorescent readings.
So to summarise, while luminescence can be measured by plate readers which can measure fluorescence, the sensitivity may be too low. In addition, extra hardware requirements, such as auto-injectors, may be required for certain luminescence assays.
I am René Inckemann and I am PhD student at the Max Planck Institute in Marburg, Germany. In my PhD I am using luminescence on a regular basis and I was also responsible to organize some of our Platereaders.
So therefore I can tell you that you need a Platereader which is able to measure luminescence and you can not simply take a fluorescence platereader. The hardware for measuring luminescence and fluorescence is different.
However many Platereader models in these days already come with both options for the hardware, which means that you can measure luminescence and fluorescence in the same device. You just have to check your platereader model if it is capable of measuring luminescence.So you don't need a seperate platereader just for luminescence, but you need one, which has the luminescence addon installed.
Regarding your idea for normalizing luminescence, I also have strong interest in that and we have already started working on that. Maybe we can exchange ideas at some point?
Anecdotally, all of the fluorescence capable plate readers that I have used have also been luminescence capable (BMG and Tecan models).
If access to luminescence capable plate readers is restrictive, we have also been able to measure luminescence using a GelDoc designed for imaging electrophoresis gels. This may be something to explore since I imagine gel imagers are more widespread that plate readers.
A priori what you say about the fluorescence and luminescence readers is correct, even wikipedia states this directly:
Luminescence detection is simpler optically than fluorescence detection because luminescence does not require a light source for excitation or optics for selecting discrete excitation wavelengths.
However, it is worth comparing the fluorescence and the luminescence devices sensitivity and saturation threshold. It might well be that a luminescence device, designed to detect very weak levels of luminescence is blinded (i.e., saturated) by stronger light. What is more relevant to your case, the detectors in a fluorescence device might be insufficiently sensitive for practical purposes of luminescence study.
Still, it is better to consult a real expert before spending money on a new piece of equipment.
Top reading usually provides better signal-to-noise ratios for solution-based assays such as DNA quantification or protein quantification . In general, reading fluorescence from the top is more sensitive than reading from the bottom. This is a result of the light being attenuated and scattered by the plastics of the well-bottom which can increase the background and decrease the efficiency of the measurement. Thus, the quality and thickness of the bottom clear plastic and the type of fluorophore are important considerations when designing your assay.
Fluorescent Plate Reader
The Biomolecular Analysis Facility has a Molecular Devices SPECTRAmax Gemini EM fluorescent plate reader. The instrument is located in Pinn Hall room 1074, in the Biomolecular Analysis Facility. Users need to contact Dr. John Shannon of the Biomolecular Analysis Facility at 434.243.9399 or by email at [email protected] to establish an account on the computer which controls the instrument. Dr. Shannon will offer basic training for the operation of the instrument. Instructions to get started are available. Most users need only a short time to take readings, so it is unusual to experience a wait to use the instrument, hence reservations are not required for standard use.
Some applications for the instrument are
- measuring live and dead cells in the same well, using Promega MultiTox reagents to measure proteinases specific for live and dead cells. This assay takes advantage of the instrument’s ability to measure two fluorophores in the same assay and to scan wells to look for clumps of cells.
- measuring glycogen content of cultured cells
- quantitation of DNA and RNA with PicoGreen or other dyes
- cathepsin assay
- measuring relative cell numbers with Alamar Blue
- intracellular calcium
- sensitive protein estimation
- cholesterol assay
- one user is quantitating DNA at 5 ng/mL.
Features of the SPECTRAmax instrument are
- dual monochromators (bandwidth 9 nm) allowing any fluorophore to be read within the range of 250 to 850 nm
- fluorescent spectra capability
- top or bottom reading of plates for adherent cells
- well scanning
- kinetic measurements
- 384, 96, 48, 24, 12, 6 well plates
- heating of plates
- shaking of plate
- SoftMax Pro 4.7 software for data analysis.
- claimed sensitivity of 3 fmol/well of FITC with top reading
This instrument does not measure absorbance.
It can perform some time resolved fluorescence measurements but not polarized fluorescence.
The instrument can be used as a luminometer for some applications. It does not have the sensitivity of a dedicated luminometer, nor does it have injectors required for flash luminescence assays e.g. Promega Dual Glo Luciferase assays.
Detection limit: 10 amol/well alkaline phosphatase 200 μL/well (obtained with Emerald II TM reagent from Applied Biosystems).
The instrument is user operated. The current charges for use are $15 for the first 10 minutes, $15/hr after the first 10 minutes.
For top read fluorescence, all black plates are recommended For luminescence, white plates which should be kept out of light to reduce light emission from the plastic.
An Introduction to Plate Readers
Many biological assays use bioluminescent, fluorescent or colorimetric reactions to generate a signal, which must be quantified to interpret the experimental result.
Luminescence and fluorescence signals are detected and quantified using luminometers and fluorometers, respectively, and colorimetric and absorbance measurements are detected by light absorption at specific wavelengths. If the assay is conducted in a microwell plate, the light output from these assays is measured using a microplate reader (or plate reader).
Luminometers and fluorometers range from simple, handheld devices to sophisticated microplate readers that can be integrated with automated systems for high-throughput sample analysis.
Key features to evaluate when choosing a luminescence, fluorescence or absorbance detection system include throughput needs (how many assays will you run?), flexibility (what types of assays will you be running?), portability (where will you be using the instrument?) and ease of use (how much set-up and training is required?).
Considering the number and the types of assays you plan to run will determine the detection modalities and throughput capacity that you need, and whether a dedicated luminometer, a multimode reader, or a simple fluorometer will meet your needs. If you anticipate that your needs will change over time, plate readers that offer modular formats, allowing extensibility to a broader range of assays in the future, are a good choice.
If you are looking for specific plates to use, suggestions for 96-well plates are listed below:
White Plates, Solid Bottom
White Plates, Clear Bottom
Black Plates, Solid Bottom
- Corning® Costar plate (Cat.# 3916)
- Greiner Bio-One CELLSTAR plate (Cat.# 655079)
- Nunc™ F96 MicroWell™ Plate (Cat.# 137101)
Black Plates, Clear Bottom
Need a Microplate Reader?
Costar is a registered trademark of Corning, Inc. MicroWell and Nunc are trademarks of Nalge Nunc International. Optilux is a trademark of Becton, Dickinson and Company.
Products may be covered by pending or issued patents or may have certain limitations. Please visit our web site for more information.
Using a BioTek Fluorescence Microplate Reader for Chemiluminescence Detection
Chemiluminescence is a highly sensitive technique that has been employed in a wide variety of applications primarily in the biological sciences. Here we describe the use of the FL600 Fluorescence Microplate Reader for "glow" chemiluminescent determinations.
Molecules that release light do so as a result of being in a state of high energy, often referred to as an "excited state", and then returning to a lower energy or ground state with a subsequent release of energy (Figure 1). The production of light from excited molecules, as a means to release excess energy, is unusual. In the case of fluorescence the energy necessary to achieve the excited state is provided by illumination with a specific wavelength of light. Chemiluminescence, on the other hand, does not require the input of exogenous light, but rather utilizes the energy contained within specific chemical reactions to provide the necessary energy. With very few exceptions, the efficient chemiluminescence of organic compounds is the result of oxidation (1), but specialized "high energy" oxygenated-compounds have also been developed that will lumninesce when fragmented (2).
The enzyme alkaline phosphatase catalyses the hydrolysis of phosphate moieties from a variety of different molecules. Quantitation of alkaline phosphatase enzyme activity is generally required when alkaline phosphatase has been conjugated to an antibody as part of an immunodetection system. In regards to measurement of alkaline phosphatase several different substrates have been developed that allow direct measurement of the product. For example, the hydrolysis of p-nitrophenyl phosphate (pnpp) by alkaline phosphatase results in the formation of a colored compound that absorbs light maximally at 405 nm (3). Likewise, the action of alkaline phosphatase on methylumbelliferone phosphate (MUP) results in the formation of methylumbelliferone, which can be detected by its fluorescence (4). In order to achieve greater sensitivity substrates have been developed that emit light (luminesce) after being acted upon by alkaline phosphatase.
Figure 1. Schematic diagram depicting the generation of light from luminescent molecules in the excited state. Note that destabilization or oxidation of the ground state molecule can accompany the change from ground state to excited state.
One group of compounds that emit light are the 1,2-dioxetanes. As seen in Figure 2, when the 1,2-dioxetane compound CSPD is dephosphorylated by alkaline phosphatase an unstable intermediate anion is formed.
Figure 2. Chemical reaction of substrate (AMPPD) and alkaline phosphatase yielding products that break down and eventually emit light. 1,2 Dioxetane structure that can act as a substrate to several different enzymes depending on the side groups attached for specificity. Regardless of enzyme, after removal of stabilizing side group, compound breaks down into unstable intermediates and after a series of steps eventually emits light.
This intermediate subsequently decomposes, to a carbonyl compound that eventually releases its energy as 466 nm light. Because the intermediate anion decomposes slowly and at a fixed rate, the two step process allows for steady-state chemiluminescence that is dependent on the concentration of alkaline phosphatase.
Other enzymatic activities can be measured with the same basic 1,2 dioxetane structure. For example, a substrate used for ß-galactosidase determinations has the same 1,2 dioxetane compound structure present, but has a galactose moiety substituted for the phosphate group seen in Figure 2. Hydrolysis of the galactose group from the molecule leads to the same decomposition, resulting in the emission of light.
The chemiluminescent signal can be further increased by the addition of macromolecular enhancers. Aqueous environments reduce the chemiluminescent signal by water-induced quenching. Macro-molecular enhancers exclude water from the site of chemiluminescence production, thus increasing emission efficiency. Interestingly, many enhancers will alter the emission characteristics of the generated light.
In this application note we describe the use of the FL600 fluorescence microplate reader for the determination of glow-type chemiluminescence with 1,2 dioxetane substrates in conjunction with enhancers.
Materials and Methods
Alkaline phosphatase (catalogue number P-5521) and ß-galactosidase (catalogue number G-3153) were from Sigma Chemical Company (St. Louis MO). CSPD TM , Galacton-Star TM , and Sapphire II TM enhancer were purchased from Tropix (Bedford, Mass.). Opaque white microplates (catalogue number 3912) were obtained from Costar (Cambridge, Mass.).
Alkaline phosphatase assays were performed as follows. A series of dilutions ranging from 0.0 to 2000 ng/ml of calf intestinal alkaline phosphatase were made using alkaline phosphatase assay buffer as the diluent. Alkaline phosphatase assay buffer consisted of 1 mM MgCl2, 100 mM DEA pH 9.0 in deionized water. After dilution, 10 µl aliquots of samples and standards were pipetted into microplate wells in replicates of four.
Substrate-enhancer mix was prepared fresh by diluting Concentrated Sapphire II TM enhancer 1:10 with assay buffer then adding 17 µl of concentrated CSPD substrate solution to each 1.0 ml of diluted enhancer mix resulting in a final substrate concentration of 0.4 mM. The reactions were then initiated by the addition of 200 µl of enhancer-substrate mixture. The reactions were then incubated at ambient temperature for various times dependent upon the initial enzyme concentration. Luminescence determinations were made using a FL600F fluorescence microplate reader. The sensitivity setting was at 150 and the data collected from the top with a 5 mm probe using static sampling with a 0.35 second delay, 50 reads per well. The lamp was turned off and the emission filter was removed.
ß-Galactosidase assays were performed as follows. A series of dilutions ranging from 0.0 to 500 µg/ml of ß-galactosidase enzyme were made using ß-gal assay buffer as the diluent. ß-Gal assay buffer consisted of 100 mM sodium phosphate pH 7.5 in deionized water. After dilution, 10 µl aliquots of samples and standards were pipetted into microplate wells in replicates of four. Enhancer-substrate mixture was made by diluting concentrated Sapphire II tm enhancer 1:10 with B-gal assay buffer. Concentrated Galacton-star tm substrate solution was then added to a final substrate concentration of 0.1 mM. The reactions were then initiated by the addition of 200 µl of enhancer-substrate mixture. The reactions were then incubated at 37°C for various times dependent upon the initial enzyme concentration. Luminescence determinations were made using a FL600FA fluorescence microplate reader. The sensitivity setting was at 100 or 150 and the data collected from the top with a 3 mm probe using static sampling with a 0.35 second delay, 50 reads per well. The lamp was turned off and the emission filter was removed.
The importance of appropriate incubation intervals is demonstrated in Figure 3. When the luminescence of an alkaline phosphatase reaction was measured kinetically, an initial increase in luminescence, which peaks and eventually drops off. This increase is most evident and quite rapid with high enzyme concentrations suggesting that the drop in luminescence is the result of substrate consumption by the enzyme. Incubation times that allow determinations prior to the peak are necessary in order to achieve linearity.
Figure 3. Change in luminescence over time. Luminescence determinations of alkaline phosphatase reactions (5 x 10 -2 DEA units/well) were made kinetically every 2 minutes for 30 minutes. Reactions were prepared as described in materials and methods.
Figure 4 demonstrates the importance of incubation time in obtaining a useful calibration curve. When unknown concentrations are suspected to be quite low, then a longer incubation time of 30-40 minutes is suggested to allow for enzyme activity to generate an amount of luminescent product that can be measured, while very high enzyme concentrations require an incubation time of 2 minutes or less. In our hands an incubation time of 10-15 minutes provided the broadest range of linearity. With a 10 minute incubation, enzyme dilutions over 5 orders of magnitude were found to be linear. As stated previously, increased sensitivity was obtained with longer incubations, but at the expense of being able to determine higher concentrations.
In terms of sensitivity, a 30 minute incubation time allowed for the detection of 25 pg/ml (p=0.005) of alkaline phosphatase. Taking into consideration the sample volume in the well (10 µl), the detection limit represents the determination of as little as 2.5 attomoles of alkaline phosphatase per well.
ß-Galactosidase enzyme activity was determined using Galacton-Star tm substrate. Similar to the CSPD tm substrate used for alkaline phosphatase determinations, luminescence increases in a linear fashion with increasing amounts of enzyme (Figure 5). Using a 5-minute incubation, a linear range of four orders of magnitude was observed. When the reaction was incubated for 15 minutes a linear relationship is observed for ß-galactosidase concentrations from 0 to 62.5 µg/ml (Figure 6). As with alkaline phosphatase determinations, high enzyme concentrations generally require shorter incubation times, while very low enzyme concentration naturally require longer incubation times (data not shown). In terms of sensitivity, ß-galactosidase concentrations as low as 0.01 µg/ml (p=0.03) can be detected using a 30 minute incubation. Taking into account the volume placed in the well, this represents the ability to detect 100 pg of ß-galactosidase enzyme.
This application note demonstrates the capability of the FL600 fluorescence microplate reader to perform glow type chemiluminescent determinations. While not specifically designed for this application, the FL600 with the excitation lamp turned off and the emission filter removed provides the same instrument platform as a specific luminescence microplate reader.
As with any experiment, optimization of the reaction provides the best results. In the case of 1,2 dioxetane chemiluminescence, the correct incubation time is important for linear results. High enzyme concentrations, which tend to consume substrate quickly, require very short incubation times, while low enzyme concentrations have a better signal to blank ratio if a longer incubation time is chosen.
Unlike fluorescence, which requires the input of light at specific wavelengths for emission, chemiluminescence does not require a light source for the sample to emit light. Therefore, wells adjacent to the location of the detector are capable of emitting light and potentially influencing the determined luminescence of the well the detector is measuring. This phenomenon, often referred to as crosstalk, can be a problem with luminescence determinations if very intense chemiluminescence reactions are adjacent to reactions with considerably less chemiluminescence.
When using 1,2 dioxetanes substrates for chemiluminescence, the concurrent use of enhancers is important to provide adequate signal. Macromolecular enhancers provide a non-aqueous environment, which increases the emission efficiency of light production by preventing water-induced quenching. These enhancers also alter peak wavelength of emission. For example Sapphire-II tm enhancer will shift the emission maximum of 1,2-dioxetanes from 461nm to 475 nm, while the enhancer Emerald-II tm shifts the maximum to 542 nm. Depending on the application, different enhancers are more appropriate than others. Because Emerald-II TM provides a greater absolute intensity than Sapphire-II TM at its peak emission wavelength, it is the optimum choice if very low concentrations or maximal signal intensity is required. On the other hand, Sapphire-II tm provides a greater dynamic range since the photodetector is less likely to be saturated.
In this application note we have described the use of the FL600 for chemiluminescence determinations. Although the 3 mm detector probe provided very good results, in our hands the 5 mm detector probe was slightly superior in regards to detection limit performance. Although we only utilized white opaque microplates with reading from the top, clear bottom plates could also be utilized for bottom reading. The use of opaque sides is still recommended. Regardless of the read mode, only glow-type chemiluminescent reactions are appropriate for measurement with the FL600. Flash-type reactions occur within seconds of adding substrate requiring a fluidics injector located within the instrument. Glow-type reactions, take place at much slower rates allowing the user to add reagents to the microplates and begin luminescence determinations before the reactions have gone to completion.
(1) Van Dyke, K. ed. (1985) Bioluminescence and Chemiluminescence: Instruments and Applications vol. 1, CRC Press, Inc. Boca Raton, Florida.
(2) Van Dyke, K and R Van Dyke Eds. (1990) Luminescence Immunoassay and Molecular Applications, CRC Press, Boston, Massachusetts.
(3) Crowther, J. (1995) ELISA Theory and Practice, Humana Press, Totowa, New Jersey.
(4) Haugland, R. (1996) Handbook of Fluorerscent Probes and Research Chemicals 6th edition, Molecular Probes Inc., Eugene, Oregon.
(5) Van Dyke, K. ed. (1985) Bioluminescence and Chemiluminescence: Instruments and Applications vol. 2, CRC Press, Inc. Boca Raton, Florida.
What is a dual monochromator spectrofluorometer system?
Some fluorescence microplate readers use dual monochromators instead of filters to select the wavelength of light that excites the sample, as well as the wavelength that is emitted by the sample. Monochromators use a diffraction grating to spatially separate the colors of light and can select a wide range of wavelengths without the need to install a separate filter for each wavelength required by an application.
Our SpectraMax Gemini dual-monochromator spectrofluorometer uses two scanning monochromators for selection of the optimal excitation and emission wavelengths.
DNA Quantification using Hoechst 33258
An essential element of cellular and molecular biology is the ability to quantitate DNA in large numbers of samples at a sensitivity that enables determination of small amounts of sample. Here we describe a fluorescent method to quantitate DNA with Hoechst 33258 dye and the BioTek FL600 fluorescence microplate reader. This assay provides linear results over three orders of magnitude and can routinely detect 8 ng of DNA. Reduction of the dye concentration was found to improve sensitivity to as low as 600 pg/well.
Many techniques of cellular and molecular biology require the ability to quantitate dsDNA in large numbers of samples at sensitivities that only require a small amount of the total sample. For example, cycle sequencing reactions require that appropriate concentrations of bacterial miniprep DNA are used in order to be successful on a consistent basis. Many biochemical studies that involve the growth kinetics of cell cultures or cell cycle studies require normalization by DNA content.
Although there are many different methods to quantitate DNA, most methods have disadvantages that preclude their use in many applications. Absorbance measurements at 260 nm (A260) is the most commonly used method for DNA concentration determination, but it suffers from the interfering absorbance of contaminating molecules (1). Many of these contaminants, which include nucleotides, RNA, EDTA and phenol, are commonly found in nucleic acid preparations. The fluorescent bisbenzimide (Hoechst) dyes circumvent many of these problems. Hoechst 33258 dye is relatively selective for dsDNA and in high salt does not show fluorescent enhancement in the presence of either protein or RNA. The dye, weakly fluorescent itself in solution, binds specifically to the A-T base pairs in dsDNA resulting in an increase in fluorescence and a shift in the emission maximum from 500 to 460 nm (2, 3). The use of Hoechst 33258 in conjunction with the BioTek FL600 fluorescence microplate reader offers high specificity, as well as high sensitivity for dsDNA quantitation.
Materials and Methods
Bisbenzimide (Hoechst 33258), catalogue number B-2883, was purchased from Sigma Chemical Co. (St. Louis, Missouri), as were sodium chloride and sodium monobasic phosphate. The 96-well black microplates with clear bottom, catalogue number 3603, were purchased from Costar (Cambridge, MA). Low fluorescent background black flat-bottom plates, catalogue number 011-010-7805, were obtained from Dynatech Laboratories, Inc. (Chantilly, VA). Assay buffer (2M NaCl, 50mM NaH2PO4, pH 7.4) was previously prepared and sterilized by autoclaving and stored at 4°C. Prior to use a portion of the buffer was allowed to warm to room temperature. Hoechst dye stock (1 mg/ml in distilled H2O) was previously prepared and sterilized by filtration through a 0.22 µm filter and stored at 4°C in a light tight container. Working assay solution was prepared fresh prior to each assay by mixing 1 µl of concentrated dye stock solution for every 1 ml of assay buffer required resulting in a final Hoechst 33258 concentration of 1 µg/ml. Working assay solutions containing final Hoechst dye concentrations of 0.1 µg/ml and 0.01 µg/ml were prepared by diluting the 1.0 µg/ml solution with assay buffer 1:10 and 1:100 respectively.
Dilutions of sonicated herring sperm DNA were made using the three working assay solutions, each containing different dye concentrations, as the diluent. After the dilutions were made, 200 ml aliquots were pipetted into microplate wells in replicates of six. Fluorescence was determined using a BioTek Instruments FL600 fluorescent plate reader with a 360 nm, 40 nm bandwidth, excitation filter and a 460 nm, 40 nm bandwidth emission filter. The data collected from either the top or the bottom using static sampling with a 0.35 second delay, 100 reads per well at several differing sensitivity settings as required by experimental conditions. Although in these experiments the plates were read immediately, if they remained sealed and protected from light the reaction was found to be stable for several hours.
The fluorescence intensity was determined for DNA concentrations ranging from 0 to 20 µg/ml (figure 1). Over this range the intensity increased in a linear fashion.
Using Microsoft® Excel TM a least means squared linear regression analysis can be generated with a coefficient of determination (r 2 ) value of 0.999.
The average coefficient of variation (%CV) of the standards was less than 2%, with the greatest percent variation taking place in the lower DNA concentrations tested (data not shown). In terms of sensitivity, the assay was found to be sensitive to the nanogram level. Under appropriate sensitivity settings, DNA concentrations as low as 40 ng/ml were found to be statistically different (p=0.005) from the working assay solution only, 0 ng/ml DNA, control. Quantitation of dsDNA using the fluorescent properties of Hoechst dye 33258 in conjunction with the BioTek FL600 allows researchers to quantitate as little as 8 ng/well (40 ng/ml in a 0.2 ml total volume). Using the described conditions the assay was found to be linear over three orders of magnitude.
In order to achieve greater sensitivity a number of measures can be undertaken to reduce background fluorescence. The use of different microplates that utilize low fluorescent plastics would be expected to reduce background and therefore increase sensitivity. Likewise, the reduction of background dye fluorescence would also be expected to increase sensitivity. Because of the inherent background fluorescence of bisbenzimide in solution, the reduction of Hoechst 33258 dye concentrations to below 1 µg/ml has been suggested as a means of decreasing background fluorescence and thus improving the detection of lower DNA concentrations (3). In order to optimize our detection limits, we sought to examine the effect of dye concentration on the fluorescent signal obtained over a range of DNA concentrations.
The fluorescent intensity was determined for DNA concentrations ranging from 0 to 400 ng/ml using three different concentrations of bisbenzimide dye. Figure 2 demonstrates that decreasing dye concentrations result in lower fluorescent signals for a given DNA concentration even after correction for differing baseline fluorescent signals. Dye concentrations of 1.0 µg/ml and 0.1 µg/ml both produce a linear response with respect to DNA concentrations from 0 to 400 ng/ml, with coefficient of determination values indicating a high degree of confidence (r 2 =0.97 and r 2 =0.98 respectively).
Further dilution of the dye concentration to 0.01 µg/ml results in a hyperbolic shaped curve indicating that for DNA levels above 200 ng/ml the bisbenzimide dye is no longer in excess. From these data we decided that reducing the bisbenzimide dye concentration to 0.1 µg/ml would result in lower background fluorescence, yet provide adequate amounts of dye to provide linearity.
As demonstrated in figure 3, a ten fold reduction in dye concentration along with the use of microplates with low background fluorescence result in lower detection limits. In this experiment, using the lower Hoechst 33258 concentration of 0.1 µg/ml reduced the detection limit to 3 ng/ml (p=0.03) or approximately 600 pg/well. The average coefficient of variation (%CV) of the standards was still less than 2% (data not shown).
Because a low fluorescent signal can be offset by an increase in the sensitivity setting on the FL600, the effect of different sensitivity settings on the fluorescent signal was tested using 0.1 µg/ml Hoechst 33258 dye with DNA concentrations ranging from 0 ng/ml to 400 ng/ml. As shown in figure 4, increasing the sensitivity will increase the fluorescent signal obtained from a given DNA and bisbenzimide dye combination. It is important to note that by correcting each sensitivity setting to zero for the 0 ng/ml DNA sample, any increase in signal due to an increase in the background fluorescence has been eliminated. After correction, the signal at each specific DNA concentration is amplified with an increase in the sensitivity, resulting in divergent increasing values. A simple increase in the baseline fluorescence would result in parallel lines that would be superimposed upon one another after correction.
Optimization of assay conditions must be determined empirically, as several factors must be weighed against one another. For most investigators using the conditions described by Labarca and Paigen (3) will be adequate. In fact, using a Hoechst 33258 dye concentration of 1.0 µg/ml would be expected give a linear response to a wider range of DNA concentrations than lower dye concentrations. Alternatively for low levels of DNA the use of 0.1 µg/ml dye concentrations may be more appropriate, as background fluorescence is reduced. The caveat of this condition being a decrease in fluorescent signal response resulting in a flatter dose response. The reduction in fluorescent signal in turn can be compensated for by increasing the sensitivity setting on the FL600 fluorescent plate reader, but in doing so, earlier saturation of the photomultiplier tube results in the loss of ability to quantitate higher levels of DNA. The ability to adjust both the assay and the FL600 allows the user to adjust the assay to meet his or her individual needs.
The ability to perform this assay in microplates offers several advantages over the conventional tube-based fluorescence assays. Like most assays that are performed in microplates, the ability to use multi-channel pipettes greatly reduces the manual labor required to perform the assay. The microplate format also lends itself to 'off the shelf' automation for laboratories with high volume requirements. The smaller reaction volumes in microplates will lead to lower per assay costs by reducing the amount of expensive reagents necessary to perform the assay.
(1) Maniatis, T., E.F. Fritsch, and J. Sambrook (1982) Molecular Cloning A Laboratory Manual, Cold Spring Harbor Laboratory, Cold Springs Harbor, NY.
(2) Daxhelet, G.A., M.M. Coene, P.P. Hoet, and C.G. Cocito (1989) Analytical Biochemistry 179:401-403.
(3) Labarca, C. and K. Paigen (1980). Analytical Biochemistry 102:344-352.
Fluorescence Microplate Assays
Combining the sensitivity of a fluorescence-based assay with a microplate format enables a rapid, quantitative readout suitable for high-throughput analysis. In a microplate well, the fluorescent signal can be generated within whole cells, in cell lysates, or in purified enzyme preparations and may then be analyzed by measuring fluorescence intensity from the well without the need for cellular imaging.
Many of these assays include substrates, buffers, and calibration standards as well as kinetic or endpoint protocols.
To understand the difference between fluorescence and phosphorescence, we need to take a little detour into electron spin. Spin is a fundamental, unvarying property of the electron and a form of angular momentum that defines behavior in an electromagnetic field. Electron spin can only have the value of ½ and the spin orientation is either up or down. An electron’s spin is therefore designated as +½ or -½, or alternatively as ↑ or↓. Two electrons in a single orbital will always have antiparallel spin at singlet ground state (S0). Upon promotion of one electron into excited state, the electron maintains its spin orientation and a singlet excited state (S1) is formed, where the both spin orientations remain paired as antiparallel. All relaxation events in fluorescence are spin neutral and the spin orientation of the electron is maintained at all times.
However, this is different for phosphorescence. Fast (10 -11 to 10 -6 sec) Intersystem crossing from singlet exited state (S1) to an energetically favorable triplet excited state (T1) leads to inversion of the electron spin. Triplet excited states are characterized by parallel spin of both electrons and are metastable. Relaxation occurs via phosphorescence, which results in another flip of the electron spin and the emittance of a photon. The return to relaxed singlet ground state (S0) might occur after considerable delay (10 -3 to >100 sec). Additionally, more energy is dissipated by non-radiative processes during phosphorescent relaxation than in fluorescence, therefore the energy difference between the absorbed and emitted photon is bigger and the wavelength shift more pronounced. Thus, phosphorescence is characterized by a bigger Stokes shift than fluorescence.