Viewing entries in
LIFA

Automated lifetime-based screening and characterization of fluorescent proteins

Lambert Instruments has provided a versatile library of interface functions (API) that allows direct communication with the LIFA software via MATLAB, in order to control the LIFA camera and the microscope. With the aid of this API, a custom made MATLAB graphical user interface (GUI) was developed that allows multi-position acquisition.

Fluorescence Lifetime Imaging with a Time-Domain FLIM System on a Widefield Microscope

Fluorescence lifetime can be recorded for every pixel in the image simultaneously with a time-domain FLIM camera. This method requires an intensified camera, a pulsed laser and a widefield fluorescence microscope. This is typically more cost-effective than alternative methods that need a confocal set-up.

One of the most popular methods for fluorescence lifetime imaging microscopy (FLIM) is time-correlated single photon counting (TCSPC). This method requires a confocal microscope with a pulsed laser and a photomultiplier tube (PMT). The sample is briefly illuminated by a laser pulse after which the PMT counts the number of emitted fluorescence photons. The intensity I of the fluorescence emission decays exponentially after the laser pulse has excited
the sample:

I(t) = I_0\exp\left(-\frac{t}{\tau}\right).

The fluorescence lifetime $$\tau$$ quantifies the rate of decay of the fluorescence light. By scanning the sample with a focused laser beam, TCSPC systems can construct a fluorescence lifetime image of the sample one pixel at a time.

As an alternative to TCSPC on a confocal microscope, Lambert Instruments has developed a new system that brings time-domain FLIM to widefield microscopes. By carefully timing the exposure of the camera in the subnanosecond range, a light pulse profile of the fluorescence light can be captured. This method requires a pulsed laser and an intensified camera to record the raw data. Custom Lambert Instruments software then processes this data to automatically calculate the fluorescence lifetime.

Set-up

Images were recorded with the LIFA-TD, which has a CCD camera with a fiber-optically coupled image intensifier. The image intensifier boosts the incoming light levels and it can achieve gate widths of less than 3 ns. A 485 nm pulsed laser (Picoquant LDH-D-C-485 laser head with a PDL 800-B laser driver) with a fiber-optical output was coupled into a widefield fluorescence microscope (Nikon Eclipse Ti) to provide 85 ps excitation pulses.

Methods

The set-up was calibrated by recording the light pulse profile of the laser by placing a highly reflective material in the sample holder of the microscope. Next, the fluorescence decay profile of a convallaria (lily of the valley) sample was recorded. The fluorescence lifetime is determined by correlating the fluorescence emission to the light pulse profile.

Results

Figure 1 shows the fluorescence lifetime of a convallaria sample overlayed on the original image. The LIFA-TD is able to detect the small variations in fluorescence lifetime between different parts of the sample, stained with different dyes.

Figure 1: Fluorescence intensity (left) recording of convallaria sample and corresponding fluorescence lifetimes (right) overlayed on the original image.

Conclusion

Time-domain fluorescence lifetime imaging microscopy can be done on a widefield fluorescence microscope by using an intensified camera and a pulsed laser. The LIFA-TD is an entry-level FLIM system that offers an integrated solution.

Revealing Cancer's Infrastructure

Lambert Instruments has been shipping the LIFA to cancer research facilities all over the world for years. We visited Dr. Kees Jalink of the Biophysics of Cell Signaling group at the Netherlands Cancer Institute. His research group purchased the first ever LIFA to leave the labs of Lambert Instruments. Ten years later, the LIFA is still their fluorescence lifetime imaging method of choice for studying signal transduction pathways in living cells.

Confocal FLIM Applications

Confocal imaging on a widefield fluorescence microscope can now be done in combination with frequency-domain fluorescence lifetime imaging microscopy (FLIM). The increased spatial resolution in the z-direction results in lifetime images with enhanced contrast as the detection of out-of-focus emission is reduced significantly. This allows you to see differences in fluorescence lifetime e.g. between the cell membrane and the cytoplasm.

The data below were obtained with the Lambert Instruments FLIM Attachment (LIFA), either widefield (with LED light; 468nm peak) or confocal (with spinning disk CSU10 and 470nm-diode laser). The fluorescence lifetime images are generated at 2 different z-positions, z1 and z2. Snapshots of several z-positions are shown, as well as movies through even more.

z2

These images show different pollen grains: the lifetime in pseudo colours and the intensity in grey scale. Because of the pinholes in the spinning disk, the exposure time is higher with confocal imaging. In this image 220 ms exposure time per phase step was taken for the confocal image versus 195 ms for the widefield (LED light). However, when a diode laser with higher power is used, the exposure time can be shortened.

z2

These images show YFP-transfected mammalian cells and were taken with 12 phase steps of each 400 ms for the confocal, versus 70 ms for the widefield image. The calculated lifetimes are 2.34 ns (z1) and 2.42 ns (z2) in the confocal images, versus 2.58 ns (z1) and 2.57 ns (z2) in the widefield images.

The differences in lifetime could be due to the fact that the diode laser has one excitation wavelength, exactly 470 nm, while the LED has a range of wavelengths for which we used the emission band pass filter of 465-495 nm.

Total Internal Reflection Fluorescence Lifetime Imaging Microscopy

Total Internal Reflection Fluorescence (TIRF) microscopy facilitates extremely high-contrast visualization and thereby high sensitivity of fluorescence near the cover glass. Typically, the optical section adjacent to the cover glass is about 100 nm. TIRF does not disturb cellular activity, thus enabling tracking of biomolecules, and the study of their dynamic activity and interactions at the molecular level. TIRF enables the selective visualisation of processes and structures of the cell membrane and pre-membrane space like vesicle release and transport, cell adhesion, secretion, membrane protein dynamics and distribution or receptor-ligand interactions. The unique combination of TIRF and frequency domain FLIM makes it possible to measure lifetimes of, for instance, small focal adhesions near the cover glass.

Widefield

These cells (kindly provided by Ms. S.E. Le Devedec, Leiden University, The Netherlands) express dSH2-GFP in small focal adhesions as well as in the nuclei as shown by widefield microscopy. However, by the use of TIRF only fluorescence close to the coverslip is obtained, thus only the focal adhesions are excited.

Fluorescence lifetime images give a more accurate measurement in TIRF mode, as out of focus light is emitted from the average lifetime in the focal adhesions.

The images shown here are taken with the Nikon TE2000-U widefield microscope with white-TIRF illuminator, combined with the Lambert Instruments Fluorescence lifetime imaging Attachment (LIFA). As light source the modulated LED of 468nm 3W was used and as demonstrated here enough intensity was generated to obtain fluorescence lifetime images with TIRF.

Widefield

Cells (kindly provided by Ms. S.E. Le Devedec, Leiden University, The Netherlands) expressing dSH2-GFP in small focal adhesions as well as in the nuclei.

The images shown here are taken with the Olympus TIRFM (laser-TIRF), combined with the Lambert Instruments FLIM Attachment (LIFA). As light source the modulated diode laser of 473 nm 20 mW was used and as demonstrated here enough intensity was generated to obtain fluorescence lifetime images with TIRF.

TIRF

Cells (kindly provided by Ms. S.E. Le Devedec, Leiden University, The Netherlands) expressing dSH2-GFP in small focal adhesions as well as in the nuclei.

Fluorescence-lifetime imaging of synapse specific interactions in live neurons

The group of Daniel Choquet at the University of Bordeaux Segalen aims at gaining a better understanding of protein accumulation at synapses to unravel the molecular mechanisms underlying memory storage in the brain.

Biomolecular interactions

Inside cells specific interactions between biomolecules are involved in almost any physiological process. Sensing extracellular signals is a matter of receptor to adapter interactions and an intricate network of structural protein interactions maintains the shape of the cell. Finding interactions between proteins involved in common cellular functions is a way to get a broader view of how they work co-operatively in a cell. One way to observe biomolecular interactions is by doing Forster Resonance Energy Transfer (FRET) measurements. In this article some examples of different interactions are given, with the link to the paper in question.

Protein-Protein Interactions

Signal transduction pathways inside cells involve the coupling of ligand-receptor interactions to many intracellular events. These events include phosphorylation by tyrosine kinases and/or serine/threonine kinases. Protein phosphorylation change enzyme activities and protein conformations. The eventual outcome is an alteration in cellular activity and changes in the program of genes expressed within the responding cells. Phosphorylation dynamics can be imaged by FRET, by labelling two proteins-, domains-, or phospho-epitopes that come in close proximity during a phosphorylation event.

Epidermal-Growth Factor Receptor (EGFR) phosphorylation with the eYFP-(acceptor)-labelled phosphotyrosine-binding domain and eCFP (donor)-tagged EGFR.Beta-secretase (BACE) phosphorylation with BACE-GFP (donor) transfected cells fixed and stained with phosphoserine-Cy3 (acceptor).

When the enzyme is labelled by one fluorophore of a FRET pair, and the substrate by the other, FRET is expected when the enzyme cleaves the substrate.

Presenilin 1 (PS1) is a critical component of the gamma-secretase complex. This complex is involved in the cleavage of several substrates, including the amyloid precursor protein (APP). By FLIM-FRET is shown that the low-density receptor-related protein (LRP) is a PS1 interactor and can compete with APP for gamma-secretase enzymatic activity.

Endosome fusion can also be imaged by FRET:

Conformational Changes

When the N-terminus is tagged with the donor fluorophore and the C-terminus with the acceptor fluorophore (or vice versa), the conformational change of the macromolecule can be visualised by the occurrence of FRET. In the 'open' conformation no FRET will occur, while the 'closed' conformation will cause FRET. Different dyes bind to different regions in DNA and so FRET occurrence can give information on the condensation of DNA:

An example is the staining of nuclei with Hoechst, that binds to AT-rich regions and with 7-AAD (7-aminoactinomycin D) that binds to GC-rich regions. These stained nuclei give a non-homogenous FRET signal in total nuclei, hence an increased FRET efficiency is shown when the cell progresses from G1 to G2/M (condensed DNA formation) phase.

Oligomerization kinetics is used to reveal the composition of macromolecules, and can be observed by FRET:

Lipid-protein interactions

Interactions between lipids and proteins can be visualised by FRET by incorporation of fluorescent lipids in the membrane and fluorescence-tagged peripheral membrane proteins.

Intensity-based FRET

In the intensity-based Forster Resonance Energy Transfer (FRET) method, change in emission intensities from donor and acceptor fluorophores is measured. During FRET, the amount of emitted photons (emission intensity) from the donor fluorophore decreases and the emission intensity from the acceptor fluorophore increases. The FRET efficiency is basically calculated from the ratio of emission intensities from donor and acceptor before and after FRET occurrence.

To obtain accurate FRET data by sensitized emission, three images have to be acquired:

1. Donor excitation with donor emission,

2. Donor excitation with acceptor emission,

3. Acceptor excitation with acceptor emission.

The major advantage of this method over fluorescence lifetime imaging microscopy (FLIM)—which is a donor-based FRET detection—is that it can be carried out with standard wide-field or confocal fluorescence microscopes that are available in most laboratories. Moreover, it yields additional data on the acceptor population. However, quantitative sensitized emission requires significant attention for corrections and calibration, whereas FLIM-based FRET techniques are inherently quantitative from first physical principles. [Ref. Gadella TW Jr., FRET and FLIM techniques, 33, 2008]

FRET Efficiency

Forster Resonance Energy Transfer (FRET) efficiency $$E$$ indicates the percentage of the excitation photons that contribute to FRET and is defined as:

E = 1−\frac{\tau_{DA}}{\tau_D}

where $$\tau_{DA}$$ is the fluorescence lifetime of the donor in the presence of an acceptor, and $$\tau_D$$ in the abscence of an acceptor. As you can see, the more FRET occurs, the more decrease in donor fluorescence lifetime.

FRET strongly depends on the distance between the donor and acceptor fluorophores (sixth-power relationship). Fluorescence lifetime of a fluorescent molecule is inversely proportional to its FRET efficiency, thus the higher the FRET efficiency the lower the fluorescence lifetime of the donor molecule will be.

The efficiency also depends on the donor-to-acceptor separation distance R with an inverse 6th order law due to the dipole-dipole coupling mechanism:

E = \frac{R^6_0}{R^6_0 + R^6}

with $$R$$ being the distance between donor and acceptor pair and R0 being the Förster distance between donor and acceptor at which the FRET efficiency is 50%.

FRET efficiency in a single pixel of an image, does not give exact conclusions about the interactions between fluorophores. The entire 2D image gives a better overview of the interactions that occur. For example: in case of 50% FRET efficiency in a single pixel, it could be possible that 50% of the donor fluorophores have had 100% energy transfer to acceptor fluorophores, but it also could be possible that 100% of the donor fluorophores have had 50% energy transfer to acceptor fluorophores.

FLIM-FRET Experiments

Fluorescence Lifetime Imaging Microscopy Forster Resonance Energy Transfer (FLIM-FRET) has a lot of advantages over other FRET detection techniques. A major advantage is that FLIM-FRET measurements are more robust and quantitative than the FRET measurements done by, for example, sensitised emission FRET. Another advantage is that only the lifetime of the donor fluorophore has to be measured; steps to determine acceptor lifetimes are not needed. The acceptor fluorophore may therefore have an inefficient emission, or even may be a quencher, and still good quality FRET-data can be retrieved. This makes the FLIM-FRET method more versatile, faster, and easier. Furthermore, no corrections are needed for donor fluorophore emission bleed through in the acceptor emission channel.

Acceptor Photobleaching

When photobleaching the acceptor fluorophore during FRET, the non-radiative transfer of energy from the donor to the acceptor decreases. The donor fluorophore, in its turn, loses less energy and its fluorescence lifetime, with respect to FRET without a photobleached acceptor. Only when the acceptor is bleached completely, the lifetime of the donor fluorophore will be similar to the situation of no FRET occurrence (a donor-only situation).

Enhanced Acceptor Fluorescence (EAF)

In the case where the donor and acceptor fluorophores are both excited with the same excitation light wavelength, e.g. in the FRET pair GFP-YFP, a special kind of FRET can be detected. Namely, the average lifetime that is calculated is the contribution of both donor and acceptor fluorophores. Taking GFP and YFP as an example, GFP has a small lifetime compared to YFP. When no FRET is occurring, the average lifetime is measured of both GFP and YFP that are both excited by the 480nm wavelength light source. However, when FRET occurs, the energy of the GFP proteins transfers non-radiatively to the YFP proteins, so relatively more YFP emission (with a long lifetime) is taken into account. So, the average lifetime increases instead of decreases, as is normally the case when you measure only the lifetime of the donor fluorophore.

Forster Resonance Energy Transfer

Forster Resonance Energy Transfer (FRET) is the non-radiative transfer of energy from a molecule in the excited state (donor) to a molecule in the ground state (acceptor). A fluorescent donor molecule can return to the ground state by losing its energy through emission of a photon (fluorescence), or by transferring its energy to a nearby (1 - 9nm) acceptor molecule (FRET). Compared to a molecule that exhibits no FRET, the donor has more options to lose its energy. Therefore, it returns faster to the ground state, which decreases its lifetime.

FRET is a useful tool to quantify molecular dynamics like interactions of two fluorophores by microscopy. The proteins under investigation are labelled with donor fluorophores or acceptor fluorophores. Interaction between the two fluorophores is accompanied by direct energy transfer from donor to acceptor (FRET). When FRET occurs, it means that the two proteins of interest are in such close proximity that they can interact with each other.

During FRET, a quantum of energy is transferred from a donor fluorophore to an acceptor fluorophore in a nonradiative process. So, in case of no FRET, the donor fluorophore is excited and emits photons. The acceptor fluorophore does not emit photons, because it is not excited. In case of FRET, the donor fluorophore is excited, but in stead of emitting all its energy as photons, it transfers some of its energy to the acceptor fluorophore that becomes excited and emits light.

Summarising, in case of no FRET only the donor fluorophore emits photons, and in case of FRET both donor and acceptor emit photons.

FRET only occurs if...

1. The donor fluorescence emission spectrum overlaps with the acceptor absorbance.

2. The donor and acceptor fluorophores are in close proximity (i.e. 1 - 9nm, which is at the scale of protein size).

3. The transition dipole moments of the donor and acceptor fluorophores are not perpendicular.

FRET pairs

To let FRET occur, the emission spectrum of the donor fluorophore has to overlap the excitation spectrum (absorbance) of the acceptor fluorophore. Some examples are BFP-YFP, CFP-YFP, GFP-DsRed, GFP-Cy3, GFP-mOrange, YFP-RFP, and Cy3-Cy5.

Browser based calculator to find the critical distance and FRET efficiency with known spectral overlap.

Probing the refractive index of the microenvironment

Fluorescence lifetime is a property which is almost completely insensitive to fluorophore concentration. It provides the means of discrimination among molecules with a spectrally overlapped emission. A further important feature is the dependence of the fluorescence decay time to the microenvironment. This dependence varies between fluorophores and certain factors.

The fluorescence lifetime of e.g. GFP can be used to probe the direct local environment of the fluorophore, because the local refractive index affects fluorescence decay. The inverse GFP fluorescence lifetime scales approximately with the square of the refractive index.

Cell membranes normally have a higher refractive index than the cytoplasm, namely 1.46 - 1.60 and 1.35 respectively. From fluorescence lifetime measurements of GFP in a PBS solution with increasing glycerol concentrations, the expected lifetime of GFP differs from 2.17 ns in the cell membrane to 2.67 ns in the cytoplasm.

Reference: Klaus Suhling, Jan Siegel, David Phillips, Paul M. W. French, Sandrine Leveque-Fort, Stephen E. D. Webb, and Daniel M. Davis. "Imaging the environment of green fluorescent protein". Biophysical Journal, 83:3589-3595 (2002).

There was, however, no correlation observed between GFP fluorescence lifetime and the viscosity of the surrounding solution. This was researched with a variety of solutes added to GFP in buffer.

Reference: Suhling, K., D. M. Davis, and D. Phillips. "The influence of solvent viscosity on the fluorescence decay and time-resolved anisotropy of green fluorescent protein". J. Fluoresc. 12:91–95 (2002).

Ion imaging

For ion imaging, several fluorescent indicators (sensor, construct, tracer, etc) are available that have a change in quantum yield upon ion binding. This means that they emit photons with different energy, thus have different emission wavelength. Their fluorescence lifetime could also change. Therefore, there are two methods in which ion imaging can be done by use of indicators: the ratiometric method and the FLIM method.

Another method ion imaging is by the use of Forster Resonance Energy Transfer based (FRET-based) indicators that change their conformation upon ion binding. Upon the conformational change of a FRET-based indicator, its FRET efficiency changes, which is used as indicator of ion concentration. Examples of these indicators are cameleons. Cameleons are genetically-encoded fluorescent indicators for Ca2+ based on green fluorescent protein variants and calmodulin (CaM).

Reference: Miyawaki A, Griesbeck O, Heim R, Tsien RY. "Dynamic and quantitative Ca2+ measurements using improved cameleons". Proc Natl Acad Sci USA (PNAS) 96(5):2135-40 (1999)

Demonstration of the Lambert Instruments Toggel camera for single-image FLIM (siFLIM) detection of histamine-induced alterations in Ca2+ concentration. Tiny oscillations in Ca2+ levels (~2.5 s periods) are observed after addition of histamine. Such small and rapid transients would go completely unnoticed when recorded by conventional FLIM.

Video courtesy of the Netherlands Cancer Institute.

Calcium Imaging

Calcium (Ca2+) is important for signal transduction pathways.

Proton (pH) Imaging

The intracellular proton (H+) concentration (pH), as well as intracellular calcium, is important in the regulation of cellular functions including growth, differentiation, motility, exocytosis and endocytosis. To study this in more detail, measurements of the intracellular pH of resting cells can be done and the pH fluctuations inside cells after environmental perturbations can be followed.

Reference: Hai-Jui Lin, Petr Herman, and Joseph R. Lakowicz. "Fluorescence Lifetime-Resolved pH Imaging of Living Cells". Cytometry Part A 52A:77–89 (2003).

Zinc Imaging

Zinc (Zn2+) is involved in enzyme catalysis, protein structure, protein-protein interactions, and protein-oligonucleotide interactions. Zinc interacts with extracellular binding sites, which are likely to include binding sites involved in the subsequent translocation of this ion to the cell interior. Inside the cell, Zinc binds to cytosolic and organelle binding sites or is taken up by intracellular organelles.

Sodium Imaging

Sodium (Na+) is important in the signal transduction in the central nerve system.

Magnesium Imaging

Many enzymes (like kinases) require the presence of magnesium Mg2+ ions for their catalytic action, especially enzymes utilising ATP.

Chloride Imaging

Chloride (Cl-) plays a role in the central nervous system.

Potassium

Potassium (K+) plays a role in cell growth and cell viability.

Indicators for Ion Imaging by FLIM

• BCECF (pH)

• Bis-BTC (heavy metals)

• Calcium-crimson (Calcium, orange excitation)

• Calcium-green (Calcium, blue excitation)

• Carboxyfluorescein (pH)

• Carboxy-SNAFL-1 (pH)

• Carboxy-SNAFL-2 (cytosol pH)

• DM-NERF dextrans (lysosoml pH)

• Fluo-3 (Calcium)

• Fura-2 (Calcium)

• LysoSensor DND-160 (lysosomal pH)

• LysoSensor probe (pH)

• Magnesium-green (Magnesium)

• Mag-quin-1 (Magnesium)

• Mag-quin-2 (Magnesium)

• MQAE (Chloride)

• Newport Green DCF (Zinc)

• OG-514 carboxylic acid dextrans (lysosoml pH)

• PBFI (Potassium)

• Quin-2 (Calcium, blue excitation)

• SPQ (Chloride)