What is FRET?
FRET stands for Forester Resonance Energy Transfer. That doesn’t say a lot so we can break things down. A protein system that uses FRET takes advantage of fluorescent proteins. A fluorescent protein can be excited at a certain wavelength of light by a laser and emit an intensity which is absorbed by a detector. Based on the intensity of light detected you can quantify the concentration of fluorescent protein. Using two fluorophores, energy transfer occurs as the FRET donor is excited and transfers energy for the acceptor to emit. The emission intensity of the acceptor detected can help us quantify the concentration of a ligand binding to a protein. Think about this. If we have two fluorophores on a protein that come together to create light as a ligand binds the protein (protein changes conformation) then the intensity of light is directly correlated to ligand concentration! Some people also use fluorescent proteins to not only detect binding events but determine the localization of proteins in cells!
mNeonGreen fluorescent protein used for attachment to N-termini of binding proteins. Protein structures were obtained from the RCSB Protein Data Bank. All protein images were created using PyMOL. A. mNeonGreen protein in a cartoon preset displaying a fluorescent proteins characteristic beta barrel structure. B. A stick preset of mNeonGreen, insightful into the fluorescent proteins volume in 3D space. C. A sliced image of mNeonGreen exposing the chromophore center, vital for fluorescence (atoms are labelled by element).
Selecting Fluorophore Pairs
While typical fluorophores like green or red fluorescent proteins provide reasonable FRET based detection and analyte concentration measurements our team wanted to select a fluorophore pair that was more advanced. Based on an extensive review, mNeonGreen (donor) and mCherry (acceptor) was a pair that has a good energy overlap, strong intensity, good folding/maturation, and is stable in solutions near physiological conditions.
Fluorophore spectrum of the mNeonGreen and mCherry fluorophores. Optical density data for wavelengths 300 nm to 750 nm were plotted for mNeonGreen and mCherry fluorescent proteins, in green and red colors, respectively. Data was obtained from www.fpbase.org. Excitation and emission peaks are labelled as EX and EM, respectively, for each fluorescent protein. Triangular dashed region shows the approximate fluorophore pair overlap, indicating that at an appropriate distance, energy transfer will occur between the donor (mNeonGreen) and acceptor (mCherry).
Bajar BT, Wang ES, Zhang S, Lin MZ, Chu J. A Guide to Fluorescent Protein FRET Pairs. Sensors [Internet]. 2016 Sep 14 [cited 2020 Jun 2];16(9). Available from: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5038762/
Developing a Standard Curve Assay
A standard curve should be generated in order to quantify the concentration of analytes. This is a crucial test that must be done – it will directly affect how the fluorometer reads the interstitial fluid. The most basic method would be to input analytes of known concentration into the fluorometer device (that includes the protein constructs) and record the intensity. The concentration of an analyte based off of the intensity (absorbance), can be determined using the Beer-Lambert law.
A = e c l
Where A is the absorbance, e is the molar extinction coefficient, c is the concentration in moles, and l is the distance between the light source and detector. However, since we do not readily have available the molar extinction coefficient, we can generate a standard curve. That way using a linear regression, at any given intensity we can accurately estimate the concentration of metabolite. Although a non-linear regression may be better suited depending on the data – our proteins are all monomers and so we use a linear model for explanations sake. A non-linear fit may be required should the initial sample data appear that way.
It is also important to note that when conducting this test – repeat the experiment multiple times. Since this data significantly effects how our device reports concentrations do more data collection than the ‘average value of 3 trials’ standard rule. The method described here is for phosphate but may be used with other analytes with small differences.
- Clean the gold chamber of the biosensor apparatus which contains the binding protein with a TRIS buffer solution (used for solvation).
- The buffer.
- Pipette 1 mmol of phosphate solution into the biosensor apparatus.
- Pipette directly into the gold chamber.
- Close the device and turn on the biosensor to activate the fluorometer.
- Wait 5 seconds to ensure an accurate reading.
- Record the intensity value given by the fluorometer in the biosensor apparatus.
- The biosensor apparatus will add a protein buffer solution to wash out the existing phosphate and free the binding protein of any analyte. Wait at least 30 min before repeating the experiment with a different concentration.
- TRIS buffer should be used for this generation step.
Note: Use concentrations at both very low and very high physiological ranges. This is so if a patient using the sensor has an abnormal analyte value, a clinician may use that data accurately for diagnosis and treatment. E.g. Phosphate values between 1-15 mmol. Increments should be 0.5 mmol or smaller for the most accurate standard curve regression.
Hypothetical phosphate standard curve. mCherry intensity at 610 nm was observed at phosphate concentrations ranging from 1-15 mmol in a biosensor apparatus. Using the standard curve, a linear regression may be used to determine phosphate concentrations based on intensity alone. Relationship between absorbance and phosphate concentration can be expressed as Abs = 0.2(phosphate conc. in mmol) - 0.1.