Fluorescence Lifetime Imaging of Molecular Rotors in Living Cells

The overall goal of the following experiment is to perform fluorescent lifetime imaging microscopy or flimm of fluorescent molecular rotors in living cells. This is achieved by first staining cells with modified hydrophobic. Bo dippy dies that act as molecular rotors, fluorescent molecules whose fluorescent lifetime is a function of the viscosity of their microenvironments.

As a second step time domain phlegm is performed using a confocal laser scanning microscope. Next, the data is fitted to an exponential decay to yield a fluorescence lifetime for each pixel. A flim map accompanied by fluorescence lifetime histogram plot can then be generated by plotting the lifetimes extracted from each pixel of the flim image.

A histogram of fluorescence lifetimes can be obtained across the whole image, allowing for mapping the micro viscosity in distinct cellular regions of living cells. Viscosity maps of single cells have until recently been hard to obtain. Mapping viscosity with fluorescence techniques is advantageous because its minimally invasive, non-destructive, and can therefore be applied to living cells and tissues.

Demonstrating a procedure will be my PhD student and Dr.James Levit, a postdoctoral research assistant in my laboratory. Fluorescence is a multi parameter signal that can be characterized by position, intensity, wavelength, lifetime, and polarization. And we are particularly interested in the position and the lifetime Prior to the start of this procedure.

LAR cells in a multi-well plate or dish, which can be heated at 37 degrees Celsius and can be supplied with 5%carbon dioxide on a microscope stage until the cells are approximately 80%confluence. Next, use an accurate balance to weigh out enough BO dippy C 12 dye for a 10 milliliter D stock solution as a concentration of approximately one milligram per milliliter dissolve the D in methanol with a pipette. Following this, add 10 to 20 microliters of the D stock solution to the living cells growing in four milliliters of cell culture medium per well.

This yields a micromolar dye concentration in the well. Return the cells to the incubator to allow for staining after the incubation. Wash the cells three to four times with four milliliters of optically clear cell culture.

Medium to remove excess dye. Prepare to image the cells by transferring the multi-well plate to the microscope stage connected to a temperature controller and 5%carbon dioxide gas inlet as required. Before starting a flim acquisition of fluorescent molecular rotors in cells, obtain a transmission and fluorescence image to identify the fluorescent cells.

Verify that the fluorescence emanates from the locations expected inside the cells. This should be a punctate distribution accompanied by a more diffuse staining throughout the cell. Obtain a fluorescence emission spectrum by scanning the image in different wavelength regions.

Verify that the spectrum is that of the diet or protein expected. The emission spectrum of the molecular rotor demonstrated here should rise sharply at around 490 nanometers peak at around 515 nanometers and trail off around 600 nanometers as a negative control image, a non-ST stain sample and verify that it does not fluoresce. Next, switch to flim mode by moving a mirror out of the fluorescence detection beam path.

Include an appropriate fluorescence emission filter in the fluorescence detection beam path to block any exciting light from reaching the detector. Once in flimm mode, prepare to scan the sample. Ensure that the detect count rate is no more than about 1%of the laser repetition rate.

If so, reduce the laser excitation intensity, for example, by placing a neutral density filter in the laser beam path. To avoid collecting pileup distorted For fluorescence decay curves, acquire a slim image typically for three to five minutes following acquisition, stop scanning and save the raw data. The raw data represents a three dimensional data cube consisting of spatial coordinates X and y, as well as time open the raw data in the fluorescence decay analysis software package.

Shown here is A SPC image by Becker and Hickle. This displays the fluorescence intensity image, which is simply the integrated fluorescence decay. In each pixel, select a typical pixel by placing the cursor on it.

Inspect the fluorescence decay in that pixel. If the peak count is below 100, use spatial bidding of pixels by adding the counts of adjacent pixels into the central pixel to obtain a higher peak count. This provides a higher statistical accuracy for the next step.

Next, select a global pixel threshold value and apply a single exponential decay fit to the image. The result is a fluorescence lifetime for each pixel above the threshold. Using a rainbow color scheme, the software assigns a color to each pixel.

Each pixel is then colored with the result of the fit and a flim map is obtained from the phlegm image, the frequency of certain fluorescence lifetimes versus the fluorescence lifetime itself is plotted to obtain the fluorescence lifetime histogram. This plot should be a bell-shaped curve and its mean value should be in agreement with other studies of Bo JPC 12 in cells adjust the color range such that the fluorescent lifetime distribution fits into the color range assigned to the lifetime range. At the pixels, check the reduced chi squared values for various pixels.

A value of approximately one to 1.3 indicates a good fit inspect the corresponding residuals, which should be randomly distributed around zero. If a mono exponential fit does not yield a good kai squared value and there is a systematic deviation of the residuals from zero, a more sophisticated model is required. Using an appropriate software package fit the data to a double exponential decay to account for two different probe environments.

The fit will also yield the pre-ex exponential factors or amplitudes, which give an indication of the relative amount of diet in one environment versus another. The results for the fluorescent lifetimes and pre-ex exponential factors can then be encoded in color, as before each pixel is colored according to its value and contrasted with each other. To obtain the lifetime ratio and the pre-ex exponential factor ratio for each pixel, check that the reduced chi-squared values indicate a good fit.

Inspect the residuals, which again should be randomly distributed around zero fluorescent lifetime histogram plot should accompany all images for easy visualization of average fluorescent lifetime values and the fluorescent lifetime distribution fluorescence decays measured for the molecular rotor at increasing viscosity. In methanol glycerol mixtures are shown. The fluorescence decays are mono exponential and the fluorescence lifetime varies markedly as a function of viscosity.

It increases from around 300 picoseconds in methanol to 3.4 nanoseconds in 95%Glycerol. A logarithmic calibration plot of log fluorescence lifetime versus log viscosity. For BO DPC 12 yields a straight line in accordance with the forced Hoffman equation.

Fluorescence intensity of he our cells incubated with the molecular rotor clearly shows the intracellular uptake of the molecular rotor with punctate D distribution. Probably in the lipid droplets. A phlegm image of the LAR cells incubated with Bo DPC 12 is shown.

The bright punctate regions exhibit a shorter lifetime than the cytoplasmic regions, which correspond to a lower viscosity by the Forster Hoffman equation. The fluorescence decays in every pixel of the image can be adequately fitted using a single exponential decay model. By plotting the lifetimes extracted from every pixel, a histogram of fluorescent lifetimes can be obtained across the whole image.

After watching this video, you should have a good understanding of how to map viscosity in living cells using fluorescent molecular rotors.