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Relative Fluorescence Units (RFU) or Luminescence Units (LU)

You are not expected to get the same values or even range of values for your assay, even if you use the exact same type/brand of microplate reader and microplates. RFU values are arbitrary and can vary between instruments, instrument settings, and the assay, even day-to-day on the same instrument. What you are expected to get is the same range for the concentration of the compound as shown in the manuals and that your RFU values from lowest to highest concentrations provide a good assay window. 

What is observed is indeed possible and it is not required that you obtain the same RFU values. The light source and the photomultiplier tube (PMT) are sensitive to the instrument settings (gain and voltage on the PMT), electrical input (voltage draws or surges), the age of the light source and the PMT, temperature, humidity, and other factors. 

Therefore, do not expect to get the same RFU values or even range of RFU values, but do expect to attain the expected range for the assay per the detection reagent and a good assay window. Because of this variance in RFU values, it is important to include the standard curve/controls on the same microplate as the sample. 

RFU values are arbitrary and dependent upon the instrument settings (gain and voltage on the PMT), electrical input (voltage draws or surges), the age of the light source and the PMT, temperature, humidity, and other factors. What is important is that the results for concentrations for the unknown samples are consistent per the standard curve generated on the same microplates but from the different reader. The RFU values may be different instruments, but the results should be the same. 

Note: Do not mathematically extrapolate experimental RFU values from standard curves or controls done at other times or on different instruments.

No. Negative RFU values are meaningless and denote an issue with either the instrument or the sample. If the instrument auto-calibrates, it may be calibrating incorrectly or there is something wrong with the baseline or zero controls. Never calibrate zero readings from empty wells. 

We would recommend the following:

  • Make certain the microplate inserts into the assay platform properly and that it is not tilted or off track. 
  • Visually examine the control and sample wells for air bubbles, precipitation/turbidity, contaminants, debris, or that the wells are empty or have a very low volume of liquid. 
  • If using a water blank to calibrate for ‘true zero’ baseline readings, it is possible for a buffer blank to have RFU values lower than water if there is precipitation/turbidity in the buffer (light obstruction). 

This is possible if the controls are highly autofluorescent/luminescent or if the sample wells have air bubbles, precipitation/turbidity, contaminants, or debris. You must also examine the differences in control and sample processing that can cause samples to generate less signal than the controls. For example: Were the same buffers used? Was the same pipettor and type of pipette tips used? Were materials stored in the same types of containers?

This denotes an issue with the light source, the PMT or both. Fluctuations may be due to electrical noise or a loose electrical connection. Consistently increasing RFU values for the water blank may be due to a light source was not properly warmed up before use. Allow at least 30 minutes to warm up a lamp for consistent output. A sudden drop or increase in RFU values and consistent readout thereafter denotes a voltage draw or surge. We would also suggest that the electrical line the instrument uses should not be shared by large appliances/instruments such as refrigerator/freezers, ultracentrifuges or other electrical appliances. 

See the possible causes below.

If using an automated dispenser, there may problems with certain fluidics lines (air bubbles, blockage, etc.). 

Depending upon the optical design of the microplate reader, it may be possible that you have ‘hot spots’ were light is being obstructed or deflected, either during excitation (for fluorescence-based assays) or during emission. To determine this, you can try the following:

For any assay: Fill every well of a microplate with water, with every well filled with the same volume. Take readings from these wells. 

Using another microplate: 

  • For absorbance-based assays: Fill every well of a microplate with a reference dye (ideally of the same color as the assay dye), with every well filled with the same concentration of dye and the same volume. 
  • For fluorescence-based assays: Fill every well of a microplate with a reference dye (ideally of the same fluorescent color as the assay dye), with every well filled with the same concentration of dye and the same volume. 
  • For a bio- or chemiluminescent-based assays: Fill every well of a microplate with either substrate and purified enzyme or, substrate and chemical, with every well filled with the same concentration of substrate/enzyme or substrate/chemical and, the same volume.

Analyze the microplates, collecting the relevant data (either absorbance units, RFU or LU) from every well in the microplate. 

Calculate the average value and then search for deviations from the average. If performing the analysis on an Excel™ spreadsheet, it may help to color the spreadsheet cells per the deviation for more immediate visual inspection. For example, average values are shaded grey, but below average values are shaded blue and above average values are shaded in red. Compare the data with dyes or substrates/enzymes with the water plate. Water plates for luminescence detection should not provide any LU (positive or negative) or only very low numbers. Any large values may denote light leakage in the analysis chamber. 

Are you able to detect any trend regarding where on the microplate these deviations occur? That is, do they occur in a specific area of the microplate (only at the edges or in certain column or row)? Is the data consistent from one area of the microplate to another? Deviations that occur from one half of the microplate to another may be due to the plate not being completely flat. 

If using automated dispenser, you should examine more than one microplate, to determine if there is dispensing errors within certain areas of the microplate. 

Standard Curve

The ‘flat line’ is due to a saturating signal. The signal from the early part of the flat line is the same as the later part of the flat line, even if the actual amount of light hitting the detector is different. This is because the photomultiplier tube (PMT) cannot distinguish the differences in signal in this range. 

You may remedy this by decreasing the voltage of the PMT to make the PMT less sensitive. But in doing so, you may lose signal detection at lower concentrations. When this occurs, you may have to analyze the data at two different instrument settings (high voltage and low voltage) and plot the data on two different graphs (one for lower concentrations and another graph for higher concentrations). The data derived from two different instruments settings cannot be combined or plotted together in a single graph. 

For fluorescent dyes, this may be due to dye-dye quenching. When dye molecules are in close proximity, the fluorescence may be quenched. 

Another possibility is that the dye is being modified to a non-dye structure, either by reduction or oxidation or some other chemical modification. For example, the non-fluorescent ROS indicator Amplex™ Red is oxidized to fluorescent resorufin, as part of the standard assay. But with further oxidation, resorufin can be oxidized to non-fluorescent resazurin or even further to a non-dye structure. 

For other reagents, this may be due to aggregation, turbidity or precipitation; basically anything that can cause the obstruction of light. 

This may be a limitation of your instrument and/or the range of detection reagent in the assay. Not all instruments can be set up in such a manner to cover the complete range of concentrations of the detection reagent. 

This may be a limitation of your instrument and/or the range of detection reagent in the assay. Not all instruments can be set up in such a manner to cover the complete range of concentrations of the detection reagent. 

The graph may become linear simply by plotting one or both axis in log mode, as opposed to linear values. If adjusting the axis from linear to log mode still results in a standard curve that is not linear, you may still use the curve if the plotted data provides a tight curve-fitting formula. If this does not work, we would recommend only using a portion of the standard curve that is linear or segment the curve into ranges that are linear. 

Yes. Use only that range of data that is linear and ignore the rest. You must not extrapolate beyond the linear range. 

Invert the axis: make the X-axis RFU values and the Y-axis concentration. You can now plug in the data X (RFU values) into the curve-fitting formula to solve for Y, where Y is unknown concentration.

Ensure that the microplate is inserted into the analysis chamber in the correct orientation. If the microplate well positions are mapped on the software or spreadsheet, the orientation of the microplate upon insertion into the analysis chamber must match the microplate map. For example, the data for sample in well A1 must be entered into the software map for well A1 and not H12, etc. 

For filter-based microplate readers, make sure that the filters are in the correct slots. Some microplate readers are equipped with the option to physically change the filters and dichroic mirrors seated in slots on a filter wheel or panel within the instrument. For ratiometric or multiple fluorescent color detection, the position of the filters and dichroic mirrors relative to each other on the filter wheel or panel is critical. 



Although they may say the microplates are made from the same plastic, the added ingredients may include plasticizers or other agents that absorb light at certain wavelengths. The molding used to shape the plastic may be made of metals that can inhibit certain enzymes (e.g., copper, stainless steel that may include other metals) and any possible carryover may become solubilized once a buffer is added. And finally, the method of making a no-binding surface may not be the same between manufacturers. 

Certain products specifically mention that the assay has been validated using a specific brand/catalog number of microplate. If possible, do not substitute with any other microplate. If other microplates are used they must be validated by the researcher. 


Discrepancies could result from using different assays. For example, if the published reference performed a chemiluminescent assay and your assay is TR-FRET, the results would not necessarily be the same. Additionally, using any other reagents from other manufacturers may introduce factors that create variations in the results. Even using the same enzyme supplied by two different manufacturers can result in differences in the final results.

If materials are not kept sterile, consider the possibility of microbial contamination. Even if a surface is dry, desiccated microbes or cellular debris can still retain enzymatic activity. 

For assays involving the study of reactive oxygen species (ROS) and not using live cells, degas all solutions. Dissolved oxygen and oxygen radicals from the atmosphere may be enough to convert ROS indicators. 

Any enzymes used in the assay may not be totally free of other enzymes/components that can interfere with the assay. Depending upon the method of extraction and purification, many enzymes still co-elute with other enzymes or proteins that can interfere with an assay. Check the quality of commercially available enzymes for the presence of other proteins/enzymes and any components in their storage buffer that can interfere with the assay. 

Conical and curved-bottom microplate wells concentrate the light signal into a smaller area relative to a flat-bottomed microplate well. The smaller surface at the air-liquid interface also slows the rate of evaporation. Flat-bottomed plates are recommended for cell culture and can be used in non-cell based assays but may not be suitable for non-cell based assays if smaller volumes are used. At lower volumes, a significant amount of sample volume is found at the well walls, the liquid-surface interface, creating a meniscus (less volume in the center, more volume at the edges). 

The droplets of water that collected at the surface of the sealant sheet may have picked up chemicals or other contaminants in the adhesive. 

Silanized pipette tips and containers are suitable for use when working with highly charged materials or very sticky materials (e.g., RNA) that may adhere to charged glass or plastic surfaces (as with surfaces that are not silanized). Silanized surfaces may bind hydrophobic materials used in the assay, thus resulting in less hydrophobic material being transferred to the assay mixture. 

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