SPIE Membership Get updates from SPIE Newsroom
  • Newsroom Home
  • Astronomy
  • Biomedical Optics & Medical Imaging
  • Defense & Security
  • Electronic Imaging & Signal Processing
  • Illumination & Displays
  • Lasers & Sources
  • Micro/Nano Lithography
  • Nanotechnology
  • Optical Design & Engineering
  • Optoelectronics & Communications
  • Remote Sensing
  • Sensing & Measurement
  • Solar & Alternative Energy
  • Sign up for Newsroom E-Alerts
  • Information for:

SPIE Photonics West 2019 | Register Today

SPIE Defense + Commercial Sensing 2019 | Call for Papers



Print PageEmail PageView PDF

Biomedical Optics & Medical Imaging

Fluorescence lifetime imaging for drug discovery and disease research

Decay profiles of the local fluorophore environment can provide new opportunities to study biological tissue and cell signaling.
7 July 2008, SPIE Newsroom. DOI: 10.1117/2.1200806.1175

Fluorescence provides a powerful means to obtain optical molecular contrast for many applications in biomedicine. Typically, fluorescent molecules, or fluorophores, are used to tag biological molecules of interest and fluorescence imaging is used to obtain a map of their distribution. In a conventional wide-field microscope this is a 2D map, but confocal or multiphoton microscopes can provide 3D information. By labeling different types of biomolecules with fluorophores that emit at different wavelengths, one can begin to probe molecular interactions as well as distributions, but this colocalization is limited by the resolution of optical microscopes (typically 300nm in the far field). It is possible, however, to learn considerably more about the local fluorophore environment by analyzing the fluorescence signal with respect to more dimensions.

In order to extract information about molecular phenomena beyond the spatial resolution limit, we are developing instrumentation to measure fluorescence with respect to multiple dimensions including the three spatial dimensions, plus excitation and emission wavelengths, fluorescence lifetime, and polarization. Here we focus particularly on lifetime, which can provide quantitative information concerning mixtures of fluorophores and their environment.

Because our overarching goal is to better understand disease to facilitate more effective diagnosis and therapy, we are particularly interested in using the autofluorescence of biological tissues to provide non-invasive, label-free molecular imaging. This is useful for diagnostic applications and as a means to monitor disease and the response to treatment. We have found that fluorescence lifetime imaging (FLIM), in which one measures the fluorescence decay time for each pixel in an image, provides a powerful tool to study diseased tissue.1 FLIM images of unstained fresh human cervical tissue obtained from a biopsy show a clear label-free lifetime contrast between normal and precancerous tissue (see Figure 1). This contrast was confirmed by hematoxylin- and eosin-stained histological sections. The images where obtained ex vivo with a multiphoton laser scanning microscope using time-correlated single-photon counting (TCSPC) to record the fluorescence decay profiles.

This method (TCSPC) is a useful and accurate technique that we are currently applying to microconfocal endoscopy to provide in vivo FLIM with subcellular resolution. For in vivo diagnostic screening applications, however, laser scanning microscopy and TCSPC are too slow to provide the necessary real-time read out. To address this, we are also working on a portable wide-field FLIM scanner based on time-gated imaging, which we have demonstrated acquiring FLIM images of fresh unstained tissue at up to 10 frames per second (fps).

Figure 1. Label-free FLIM images of sections of fresh human cervix. Precancerous regions exhibit shorter fluorescence lifetimes. (Figure adapted from reference 1.)

High-speed time-gated FLIM can also be useful in molecular biology and drug discovery research wherein the sensitivity of fluorescence lifetime to the local fluorophore environment can be exploited using Förster resonance energy transfer (FRET) experiments. FRET is the phenomena whereby an excited fluorophore can rapidly lose energy to a nearby molecule that is capable of absorbing its fluorescence radiation. Efficient energy transfer, which occurs via a direct dipole-dipole interaction, is only possible if the acceptor molecule is within approximately 10nm of the donor fluorophore. Thus, detecting FRET enables ‘super-colocalization’ if different proteins are tagged with donor and acceptor fluorophores. Because the efficient energy transfer leads to a decrease in the fluorescence lifetime of the donor, FLIM provides an effective means to detect and image FRET and is therefore a powerful tool to study the binding of proteins, which can be important for cell signaling.

Figure 2. (a) Schematic of Nipkow disk confocal FLIM microscope. Inset: FLIM image of live Cos cells exhibiting Förster resonance energy transfer (FRET) between the Ras protein labeled with the mCherry fluorophore and the Raf-RBD protein labeled with EGFP. (b) Nipkow confocal FLIM image, recorded at 5fps, of live Cos cells labeled with EGFP and an EGFP-mRFP FRET construct. (c) FLIM FRET image of live Cos cells exhibiting Ras-RBD-Raf FRET recorded in an automated Nipkow confocal FLIM multiwell plate reader designed for high content analysis. EGFP: enhanced green fluorescent protein. mRFP: monomeric red fluorescent protein.

Because drug discovery typically involves testing large numbers of drug candidates, high-throughput screening is highly desirable. Our high-speed time-gated FLIM technology permits us to acquire FLIM-FRET data at up to 10fps,2 allowing the study of biological signaling processes in real time. We have recently implemented FLIM FRET in an automated spinning disk confocal microscope to rapidly perform 3D FRET experiments with live cells arrayed in multiwell plates typically used by the pharmaceutical industry for high-throughput screening and high content analysis (see Figure 2). By combining spectrally resolved imaging and FLIM, we simultaneously image different protein interactions in the same cell using two distinct donor-acceptor fluorophore combinations. This multiplexed FRET imaging is an important new tool to study the spatio-temporal properties of cell signaling.

Figure 3. (a) Schematic of the excitation-resolved hyperspectral FLIM microscope. (b) Excitation-emission-lifetime resolved fluorescence data for a stained lily-of-the-valley test sample showing the fluorescence intensity image integrated over excitation and emission spectra and fluorescence decay, excitation-emission matrix (EEM) plots corresponding to the regions indicated in the intensity image, and fluorescence decay profiles corresponding to the regions indicated on the EEM plots.

Simultaneously analyzing fluorescence with respect to its spectral and temporal properties is a powerful approach for unmixing signals from multiple fluorophores, for studying the photophysics of complex systems, and for contrasting different biological components contributing to complex signals such as tissue autofluorescence. To this end we have built a line-scanning confocal fluorescence microscope that utilizes a tunable supercontinuum-based excitation source and a spectrometer in conjunction with our wide-field time-gated FLIM technology.3 By exciting a line of fluorescence and stage-scanning the sample, we can acquire the full excitation-emission-lifetime properties for each pixel in the field of view (see figure 3). This line-scanning microscope also provides a confocal optical sectioning capability and thus can resolve the fluorescence signal with respect to six dimensions.

The unprecedented information available from multidimensional fluorescence imaging provides new opportunities but also new challenges for molecular imaging, particularly for data analysis. We are currently working with the Institute of Cancer Research to use FLIM-FRET to monitor signaling in cells in order to learn more about cancer and how drugs may interfere with its progression.

Paul French
Photonics Group
Physics Department
Imperial College London
London, UK