A Guide to Fluorescence Lifetime Microscopy and Förster's Resonance Energy Transfer in Neuroscience

Abstract

Fluorescence lifetime microscopy (FLIM) and Förster's resonance energy transfer (FRET) are advanced optical tools that neuroscientists can employ to interrogate the structure and function of complex biological systems in vitro and in vivo using light. In neurobiology they are primarily used to study protein-protein interactions, to study conformational changes in protein complexes, and to monitor genetically encoded FRET-based biosensors. These methods are ideally suited to optically monitor changes in neurons that are triggered optogenetically. Utilization of this technique by neuroscientists has been limited, since a broad understanding of FLIM and FRET requires familiarity with the interactions of light and matter on a quantum mechanical level, and because the ultra-fast instrumentation used to measure fluorescent lifetimes and resonance energy transfer are more at home in a physics lab than in a biology lab. In this overview, we aim to help neuroscientists overcome these obstacles and thus feel more comfortable with the FLIM-FRET method. Our goal is to aid researchers in the neuroscience community to achieve a better understanding of the fundamentals of FLIM-FRET and encourage them to fully leverage its powerful ability as a research tool. Published 2020. U.S. Government.

Keywords: FLIM; FRET; FRET-based biosensor; conformational change; fiber-photometry; microscopy; protein-protein interaction.

Figure 1

Lifetime measurements using TCSPC. (A) Schematic depicting laser pulses from an 80-MHz pulsed laser (green spikes) used to excite a sample containing a fluorophore, and detected photons (yellow spikes) emitted from the sample. The time interval (Δt) between an excitation pulse and a detected photon is measured (see dashed red lines). (B) A Monte Carlo–simulated population of 10,000 Δt measurements from a fluorophore having a 3-ns fluorescence lifetime. (C) A histogram of the Δt values (blue bars). Dashed red line is a fit of the histogram values to a single exponential decay model which accurately recovers the simulated 3-ns lifetime. (D) A more realistic rendition of a simulated lifetime decay plotted on a semi-log plot. Note the rising phase and peak count (observed here between 0 and 1 ns) resulting from the convolution of the true underlying lifetime decay (3 ns) and the instrument response function (150 ps). The red dashed line indicates a fit to a single exponential decay model using only the data between the peak count and the count at 12.5 ns. This fit successfully recovered the 3 ns tau value. On a semi-log plot, a single exponential decay looks like a straight line with a negative slope. Also note how the fractional noise of the photon count gets larger (see the spread of blue dots around the dashed red fit) as the count gets smaller with time.

Figure 2

How the instrument response function (IRF) influences our ability to recover lifetime decay information. (A) Simulated lifetime decay (black line) with two decay components (100 ps and 1000 ps) of equal amplitude. The red dashed line is a fit of the decay data to a double exponential decay model which accurately recovers both Tau1 and Tau2 as well as their relative amplitudes (A1 and A2). The inset in panel A shows three IRFs based on commercially available photo-detectors with 20 (black trace), 70 (blue trace), and 150 ps (green trace) decay constants. The un-convolved lifetime decay depicted in panel A was convolved with either the 20 ps IRF (B), the 70 ps IRF (C), or the 150 ps IRF (D). A rising phase and peak count is seen in all three cases. In panels B and C, data between the peak count and 9 ns was fit to a double exponential decay model, while the data in panel D was fit to a single exponential decay model (red dashed lines).

Figure 3

FRET and how it influences donor lifetime decays. (A) Cartoon depicting a donor fluorophore in the absence of a FRET acceptor (yellow circle) being excited by a photon (blue star) and emitting a red-shifted photon (yellow star) at rate K_E. (B) Cartoon depicting a donor fluorophore (yellow circle) in the presence of a FRET acceptor (red circle). The donor fluorophore is excited by a photon (blue star), but now has a choice of either emitting a red-shifted photon (yellow star) or transferring its excitation energy via FRET to a nearby acceptor (red circle). (C) Simulated lifetime decay data for a donor alone (blue trace) or a donor in the presence of an acceptor (green trace). In this simulation, the donor alone had a lifetime of 3 ns, the acceptor was Föster’s distance (R₀) away from the donor, and it was assumed that the donor and acceptor were in the isotropic dynamic regime with κ² equal to 2/3. Dashed and dotted red lines are fits to a single exponential

Figure 4

The speed of molecular rotation relative to the donor lifetime can influence FRET donor-lifetime decays. (A) The average FRET efficiency (<E>) as a function of the donor-acceptor separation (R) divided by the Föster distance (R₀) in the isotropic dynamic regime (dashed blue line) or in the isotropic static regime (red line). (B) The κ² probability distribution (green trace). Note that when an isotropic population of fluorophores rotate faster than their excited state lifetime, it can be assumed that κ² has a value of 2/3, the average value of the κ² probability distribution. If isotropic fluorophores rotate slower than their excited state lifetime, the heterogenous nature of the isotropic κ² probability distribution may impart heterogeneous lifetime decay characteristics to a lifetime decay curve. (C) The influence of dynamic (top panels) versus static (bottom panels) regimes on FRET donor lifetime decays. Dynamic regime is typically observed with small organic fluorophores that rotate rapidly, while the static regime is typically observed with large fluorophores, such as FPs, which rotate slowly. Four Monte Carlo–simulated decay curves are depicted: when R/R₀ = 1 (FRET) or 2 (no FRET), for the dynamic or static regimes. Note that molecular rotation does not alter the donor lifetime decay in the absence of FRET. In the presence of FRET, the donor lifetime decays as a single exponential in the dynamic regime, but decays multiexponentially in the static regime.

Figure 5

Homo-FRET is measured using a variant of FLIM called time-resolved anisotropy, a technique that measures the time-dependent polarization of light emitted from a fluorophore. (A) Cartoon depicting the polarization of light emitted from a fluorophore in the absence or presence of homo-FRET. On the left is shown an excited fluorophore emitting light with an orientation correlated with the orientation of the linear polarized excitation light (i.e., polarized emission). In contrast, the right depicts how the orientation of emitted light changes due to homo-FRET. In homo-FRET, excited energy can migrate back and forth between like fluorophores before de-excitation. Light emitted from the initially excited ‘donor’ fluorophore is correlated to the excitation light, while light emitted from the ‘acceptor’ fluorophore is uncorrelated or depolarized. (B) Photon decay histograms collected from a sample of mVenus fluorophores. Lifetime decays measured using either a detector configured to collect light that had an orientation parallel to the linear polarized excitation light (black trace) or from a detector configured to collect light that had an orientation perpendicular to the excitation light (green trace). (C) Anisotropy decay curves, calculated from orthogonal polarization lifetime decay histograms similar to those shown in (B) using Equation 16, for mVenus (green trace), mUranus-17-mVenus (blue trace), mVenus-17-mVenus (orange trace), or mCherry-17-mVenus samples (red trace, 17 in these construct names denotes the length of the amino acid linker used, and mUranus is a point mutant of an FP that cannot form its chromophore). Starting the moment after excitation, mVenus anisotropy decays over time and is well described by a single exponential decay model. The decay time constant of this curve is the rotational correlation time of mVenus, and represents depolarization due to molecular rotation. Similarly, the mVenus-17-mUranus anisotropy curve decays according to a single exponential model. This curve (and the mCherry-17-mVenus curve) demonstrates how molecular rotation can slow down when a fluorophore is tethered to another protein. The mVenus-17-mVenus anisotropy decay curve is well described by a double exponential model. The fast decay time constant is associated with homoFRET-dependent depolarization, while the slow decay constant is due to molecular rotation. The gray-shaded area in between the mUranus-17-mVenus and mVenus-17-mVenus decay curves represents depolarization due primarily to homo-FRET. (D) Photon decay histograms for the samples described in (C). A decrease in donor fluorescence lifetime due to hetero-FRET is clearly shown as the gray-shaded area in between the mUranus-17-mVenus and mCherry-17-mVenus decay curves. Notice that the change in anisotropy for homo-FRET and lifetime for hetero-FRET in (C and D), respectively, are qualitatively similar.

Figure 6

Cartoon depicting conformational changes in the CaMKII holoenzyme structure observed, in part, using FRET microscopy. The autoinhibited enzyme, as depicted in (A), is composed of an assembly of 12 subunits each having a catalytic domain (blue) on its N-terminal end and an oligomerization domain (gray) on its C-terminal end with a regulatory (yellow)/linker (red) domain connecting the two. The pairing of catalytic domains was clearly observed by both homo-FRET and hetero-FRET microscopy. The holoenzyme is activated when calcium-calmodulin (green U-shaped blobs) bind to the regulatory domain. If the kinase is activated in the absence of T-site ligands (orange circles), the holoenzyme assumes a paired but extended structure as depicted in (B1). This was observed by a combination of homo-FRET microscopy and FCS diffusion analysis. If the kinase is activated in the presence of T-site ligands (orange circles), the holoenzyme assumes an unpaired extended structure as depicted in (C). This was also observed by a combination of homo-FRET microscopy and FCS diffusion analysis. It is not known if the binding of T-site ligands upon activation, such as to the NMDA receptor, proceeds via an (A to B1 to C) pathway or alternatively via an (A) to (B2 to C) pathway. Some homo-FRET experiments suggest that a transient unpaired activated holoenzyme structure as depicted in (B2) might exist.

Figure 7

Schematic of FRET biosensors. (A) Enhanced FRET can be observed when the biological factor of interest (red square) interacts with the sensing domain (blue circles) of the biosensor. (B) Similarly, in some biosensors, diminishing FRET can be observed when the biological factor of interest interacts with the sensing domain. (C) Diminishing FRET can also be observed when a protease (red square) cleaves the protease’s specific peptide substrate (blue circle) linking the donor and acceptor. (D) For some biosensors, FRET can only be detected when a biological factor promotes the association of two separate sensor components. (E) Cartoon depicting the expected fluorescence lifetime decay curve changes in response to FLIM-FRET biosensor activity.