Fluorescence Lifetime Imaging in Scanning Microscopy

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Fluorescence Lifetime Imaging in Scanning Microscopy H.C. Gerritsen, A. Draaijer, D.J. van den Heuvel, and A.V. Agronskaia

INTRODUCTION

Fluorescence Lifetime Spectroscopy

Fluorescence, Lifetime, and Quantum Efficiency It was not until the 1930s that measurement of the fluorescence lifetime (tf ) and the confirmation of theory on this phenomenon became possible (Pringsheim, 1961). Before that time only phosphorescence lifetimes had been measured, evidently because this phenomenon is one or more orders of magnitude slower. The distinction between fluorescence and phosphorescence was first made by the observation of the afterglow. Emission having a noticeable afterglow was called phosphorescence. All other processes that did not have a noticeable afterglow were called fluorescence. Later, the distinction between the two was based on quantum mechanics. In phosphorescence a triplet to ground state transition is involved. Quantum mechanically such a transition is spin forbidden and therefore has a low probability. Consequently, the rate at which such a process occurs is low and the lifetime is long. The fluorescence phenomenon only involves singlet states with transition probabilities that are much higher, making the process faster. In its simplest form, the fluorescence or phosphorescence intensity decay after the excitation source is shut off follows an exponential curve described by: I(t) = I0e-t/t

(1)

where I(t) is the fluorescence intensity at time t, I0 is the the fluorescence intensity at time zero, and tf is the time when the fluorescence has dropped to I0 /e. The fluorescence quantum efficiency Ff (Straughan and Walker, 1976) is defined as the ratio of the number of photons absorbed to the number of photons emitted as fluorescence by the molecule. It is dependent on the relative magnitudes of the rate constants involved in fluorescence and other competing decay processes. The processes involved in the decay of a molecule with singlet ground state, S0, excited singlet state, S1, and excited triplet state, T1 are diagrammed in Figure 27.1 (Jablonski diagram) and listed in Table 27.1. It can be shown that Ff = kf /SkProcesses and tf = 1/ SkProcesses = Ff /kf where Ff is the fluorescence quantum efficiency, kf is the fluorescence decay rate constant, kISC is the rate constant of intersystem crossing, kIC is the rate constant of internal conversion, and kRe is the rate constant for reaction product. In the case of, for example, different microenvironments or different conformations of the fluorescent molecules, the decay curve may become multi-exponential.

Early measurements of nanosecond lifetimes on bulk specimens were performed by means of Pockel or Kerr cells (Pringsheim, 1961). Such devices contain an optically transparent, nonlinear crystalline solid (Pockel cell) or an optically transparent, nonlinear liquid (Kerr cell). The polarization direction of light passing through these cells is rotated by putting a voltage across them. Because detectors and amplifier electronics were not yet fast enough for direct measurements, tf could only be measured by applying a time difference between the excitation and emission cells. This was accomplished by changing the distance between the two cells and measuring the average intensity as a function of the time difference between the two cells. Light travels 30 cm in 1ns. The popularity of fluorescence lifetime spectroscopy has grown considerably in recent decades because of the availability of fast electronics, detectors, and light sources. A number of techniques are available to measure the decay curves of fluorescence processes. In general, tf measurements are either made in the time domain or in the frequency domain. In the frequency domain, one measures the phase shift of the fluorescent light with respect to the phase of a modulated excitation source. Measurements in the time domain are generally performed by measuring the time dependency of the fluorescence intensity with respect to a short excitation pulse. It falls beyond the scope of this chapter to give a detailed overview of all the numerous implementations and applications of fluorescence lifetime spectroscopy. The reader is referred to the existing literature (Lakowicz, 1983; O’Connor and Phillips, 1984; Clark and Hester, 1989). Many applications of fluorescence lifetime spectroscopy can be found in the fields of protein studies, membranes and cells, and energy transfer and ion concentration measurements. We will restrict our consideration to microscopic applications.

Fluorescence Lifetime Imaging Applications Apart from measurements on bulk specimens, fluorescence lifetime spectroscopy equipment can be modified for microspectroscopy and microscopy. This makes it possible to measure tf of small volumes in a specimen. Starting in the last decade of the 20th century, fluorescence lifetime measurements were extended into the field of widefield imaging (Morgan et al., 1990, 1992; Lakowicz and Berndt, 1991; Ni and Melton, 1991; Wang et al., 1991; Gadella et al., 1993; Lakowicz and Szmacinski, 1993; Webb et al., 2002), confocal imaging (Bugiel et al., 1989; Buurman et al., 1992; Sanders et al., 1994), and multi-photon excitation

H.C. Gerritsen, D.J. van den Heuvel, and A.V. Agronskaia • Utrecht University, Utrecht, The Netherlands A. Draaijer • TNO-Voeding, Utrecht, The Netherlands 516

Handbook of Biological Confocal Microscopy, Third Edition, edited by James B. Pawley, Springer Science+Business Media, LLC, New York, 2006.

Fluorescence Lifetime Imaging in Scanning Microscopy • Chapter 27

FIGURE 27.1. Jablonski diagram showing the processes involved in the decay of a molecule with singlet ground state S0, excited singlet state S1, and excited triplet state T1.

imaging (Piston et al., 1992; So et al., 1996; French et al., 1998; Sytsma et al., 1998). Fluorescence lifetime imaging (FLIM) can be used, for example, in multi-labeling experiments, for measuring ion concentration or cell chemistry, and for measuring fluorescence resonance energy transfer (FRET) efficiencies. These uses are discussed below.

Multi-Labeling with Dyes In general, probes that have the same emission wavelength usually do not have the same tf, so they can be distinguished based on their lifetime differences provided the microscope has sufficient lifetime resolution. The number of probes that can be imaged simultaneously depends on the lifetime range of the FLIM system. In the spectral domain, the emission bands are broad and in conventional fluorescence microscopes not more than two or three probes can be imaged simultaneously, with each probe necessarily having its own detection channel. However, given a typical tf resolution in the sub-nanosecond range and a dynamic range of tens of nanoseconds, it should be possible to simultaneously image 10 to 20 probes.1 Recording probes with differing tf can be achieved with only a single detector and one excitation wavelength, and it does not require the use of spectral selection on the emission side. Because no bandpass filters are needed, the complete spectral band is detected and detector sensitivity is increased. Discrimination of the signals from specific probes based on their tf alone can be done using simple image processing. In combination with spectral selection, the number of distinct probes that can be imaged becomes even larger. With some a priori knowledge of the tf of the probes used, signals from mixtures of probes are easily deconvolved and the relative abundances calculated (Verveer et al., 2000). Some probes have even been designed specifically for tf imaging (Sauer et al., 1993).

Concentration Imaging (pH, Ca2+, K+, Na+, O2, etc.) For many existing fluorescent ion indicators, the tf of the free form differs from that of the ion-bound form. In principle this property can be used for the quantitative imaging of ions (pH, Ca2+, K+, Na+)

1

As long as they do not all occur in the same location.

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(Lakowicz and Szmacinski, 1993; Lakowicz et al., 1994; Sanders et al., 1994, 1995). Oxygen and other quenching agents reduce tf (Gerritsen et al., 1997). This effect can also be used for imaging purposes, making it possible to image important biological parameters. Fluorescence lifetime imaging has tremendous advantages compared to other methods that are either invasive (microelectrodes) or require a calibration on the specimen (ratio imaging). Using tf as a parameter for measuring dye concentrations makes the measurements independent of intensity effects such as shading in the image, changes in laser intensity, or absorption in the specimen, and calibration is often easier than calibration of l-ratio images. For several fluorescent ion indicators it was shown that no in vivo calibration procedure is required (Sanders et al., 1995; Herman et al., 1997).

Chemical Environment The tf of a given probe is often sensitive to local differences in chemical environment. This can be exploited to obtain information about micro-environments in general. For example, one could image hydrophobicity, viscosity (molecular friction effects), mobility, or membrane potential.

Quantitative Fluorescence Intensity Measurements Most users of a fluorescence microscope measure the concentration of a fluorescent probe to get information about, for example, the extent of DNA damage or the structure of an object. However, the intensity signal is only a good measure of the dye concentration if the quantum efficiency stays constant and the environment often affects the quantum efficiency. A calibration procedure is required to correlate the fluorescence intensity to the probe concentration and this procedure may be suspect because the calibration environment is different from the specimen environment. Fortunately, tf is closely related to the quantum efficiency and so a measurement of tf can be used to check the validity of the calibration procedure. If the tf measured during calibration is equal to that in the specimen, this indicates that the quantum efficiency is the same in both cases and thus the intensity can be correlated to the concentration. If factors that do not effect tf, such as photobleaching, occur, making the tf measurement increases the reliability of the intensity measurement. In some cases it has been possible to correct for changes in the quantum efficiency by using tf information (Morgan et al., 1990).

Fluorescence Resonance Energy Transfer One of the more recent applications of fluorescence microscopy is the imaging of colocalization on a nanometer scale by means of Förster resonance energy transfer (Förster, 1946; Lakowicz, 1983; Clegg, 1996). This technique is also often referred to as fluorescence resonance energy transfer (FRET). More details on FRET can be found in Chapter 45. In conventional fluorescence microscopy colocalization can be studied by using two differently labeled molecules that emit in two distinct emission bands. As soon as signals from both detection channels occur in the same pixel the two molecules colocalize within the resolution limit of the microscope. Because the resolution of the microscope is much larger than the size of the molecules, colocalization measured in this way does not prove that the two molecules interact. FRET can be employed to study colocalization on a scale of a few nanometers and therefore this method does yield information about molecular interactions. In FRET an excited

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TABLE 27.1. Processes Involved in the Decay of an Excited Molecule Molecule S0 + hn Molecule S1 Molecule S1

æ æÆ K ææÆ K ææ æÆ

Molecule S1 Molecule S1 Molecule S1

æ æÆ K ææ æÆ K ææ æÆ

F

ISC

K IC

Re

NR

Molecule S1 Molecule S0 + hn Molecule T1 Molecule S0 Products Molecule S0

Excitation Fluorescence (F) Intersystem crossing (ISC); non-destructive quenching effect (triplet state build-up) where molecule will become available again only after returning to the ground state (phosphorescence) Internal conversion (IC); non-destructive quenching effect (molecule stays available) Reaction (Re); Destructive quenching effect (molecule is lost for fluorescence) Non-radiative decay (NR); molecule returns to the ground state without fluorescing

fluorescent donor molecule directly transfers its excited state energy to an acceptor molecule. This is a resonant process that occurs through a dipole–dipole interaction between the donor and acceptor. A prerequisite for FRET to occur is that the emission band of the donor overlaps with the absorption band of the acceptor and in addition the two molecules should be within a few nanometers of each other. The presence of the acceptor results in the introduction of an additional decay channel for the excited donor molecules and therefore the total decay rate of the donor molecule is increased. Consequently, FRET results in a reduction of both the donor fluorescence intensity and the fluorescence lifetime (tf = 1/SkProcesses). FRET can be conveniently quantified in terms of the energy transfer efficiency E. E can be expressed in the relative donor intensity reduction E = 1 - IDA/ID, and in the relative donor lifetime reduction E = 1 - tDA/tD. Here, IDA and ID are the intensities of the donor in the presence and absence of the acceptor respectively. tDA and tD are the lifetimes of the donor in the presence and absence of the acceptor, respectively. FRET imaging experiments can be carried out by means of fluorescence intensity measurements. However, this approach requires the use of emission filters and quantification of E is complicated by, for example, filter bleedthrough and direct excitation of the acceptor. In order to correct for these effects multiple reference images need to be recorded. FRET imaging can also be accomplished by means of FLIM (Wouters and Bastiaens, 1999). Now, the energy transfer efficiency E can be directly derived from a single measurement of the donor lifetime, provided that the donor lifetime of dye in the absence of the acceptor is homogeneous throughout the specimen. This approach is straightforward and at present FLIMbased FRET imaging seems to be the preferred method to carry out quantitative FRET experiments on interactions between molecules. From the applications listed above it is obvious that fluorescence lifetime contrast is a versatile and powerful tool in microscopy. In the next section we discuss the basics of frequencyand time-domain methods for implementing FLIM and a number of applications of this technique will then be described.

FLUORESCENCE LIFETIME IMAGING METHODS Introduction FLIM can be implemented in widefield microscopes (WF), confocal laser-scanning microscopes (CLSMs), and multi-photon excitation microscopes (MPEMs). The use of the point scanning geometry in FLIM is advantageous. If a diffraction-limited volume element in a specimen with a lifetime t1 is embedded in a

homogeneous background with lifetime t2 and imaged in a WF microscope, the measured tf will be averaged over the entire excited volume. This will contribute a large t2 background fluorescence component to the measured decay. In the confocal setup, however, only the small volume element near the focal plane is sampled, and therefore much less of the t2 background component is observed. At present, several techniques are being employed for the measurement of tf on bulk specimens. Basically the techniques can be subdivided in frequency-domain–based methods such as phase fluorometry (Gratton and Limkeman, 1983; Jameson and Gratton, 1983; Lakowicz, 1983) and time-domain–based methods such as time-correlated single-photon counting (O’Connor and Phillips, 1984) and methods based on time-gated detection. In either method, both the light source and the detection system have to be modified. A major difference between fluorescence lifetime measurements on bulk specimens and fluorescence lifetime imaging is the number of fluorophores available in the detection volume. In conventional non-imaging spectroscopy experiments, a large number of fluorescent molecules is present in the detection volume and the accuracy of the experiments is, in general, not limited by the strength of the fluorescence signal. In a typical confocal or multi-photon imaging experiment, only a small volume is imaged and in general a low number of fluorescent molecules is present in this volume. On assuming typical dye concentrations of tens of micromolar, we find that on the order of 100 to 200 dye molecules are present in the focus of the microscope objective. Furthermore, photobleaching constrains the maximum number of photons that can be emitted per fluorescent molecules to values between 103 to 105 photons before photobleaching takes place. For conventional confocal and multi-photon fluorescence intensity imaging, images of reasonable quality can be acquired with as few as 20 to 30 detected photons per pixel. FLIM requires much more signal per pixel to produce acceptable images. In general, at least one order of magnitude more signal is required to obtain images of reasonable quality. Therefore, the efficiency of the microscope, the detector, and the lifetime acquisition method are of crucial importance. For this reason we will discuss the factors that determine the sensitivity of the different lifetime imaging methods in detail.

Lifetime Sensing in the Frequency Domain The Phase Fluorometry Method Phase fluorometry is one of the oldest methods used for the determination of tf. The essence of the method is phase-sensitive detection in combination with intensity-modulated excitation of the specimen. Either a pulsed or a sinusoidally modulated light source is used in this technique. In the case of sinusoidally modulated

Fluorescence Lifetime Imaging in Scanning Microscopy • Chapter 27

excitation, the time-dependent excitation intensity, E(t) (see Fig. 27.2) is given by: E(t) = E0(1 + ME sin(wt))

(2)

with w the modulation frequency, E0 the average excitation intensity, and ME the modulation depth of the excitation, defined as the ratio between the AC amplitude and the DC component of the excitation signal. The delay between the absorption of an excitation photon and the emission of the fluorescence introduces a phaseshift f and demodulation of the emission F(t) with respect to the excitation light (see Fig. 27.2). F(t) can be written as: F(t) = F0[1 + MF sin(wt + f)]

(3)

with MF the modulation depth of the emission. For a simple monoexponentially decaying fluorescence signal, the following relation can be derived relating tf, w and f: tf =

1 tan(f ) w

(4)

The tf of a specimen can be accurately determined by measuring the phase difference between the excitation signal and the fluorescence signal. Assuming a strong signal, f can usually be determined to an accuracy of ±0.2°. The phase measurements are usually done at a number of different frequencies ranging from several hundred kilohertz up to many hundred megahertz. Using this approach, fluorescence decays containing more than one decay component can be examined. The value of each tf as well as its relative contribution to the total fluorescence signal can be determined (Gratton et al., 1984). As the ratio MF/ME is affected by tf , this ratio can be also employed to obtain quantitative lifetime information. The relation between the relative modulation (MF/ME) and tf is: 12 È 1 - 1˘ 1Í 2 ˙ tf = ÈM ˘ ˙ wÍÍ F˙ ÍÎ Î ME ˚ ˙˚

(5)

Intensity modulation of the exciting light can easily be accomplished by using electro-optical modulators (EOM) or acousto-

2.0 1.8 1.6

Intensity

1.4 1.2

519

optical modulators (AOM) (see Chapter 3, this volume). These devices allow modulation over a wide range of modulation frequencies, up to about 1 GHz, and can be used in combination with comparatively cheap CW lasers. Alternatively a pico- or femtosecond pulsed laser source can be used as a modulated light source. In this case one can use the harmonic content of the excitation signal to produce a range of modulation frequencies. Depending on the pulse-width of the laser, the frequency content can extend up to 200 GHz, far higher than the best available detectors. On the detection side, a phase-sensitive detector is required. This is often realized by using a gain-modulated photomultiplier tube (PMT) or a micro-channel-plate detector, modulated at exactly the same frequency (homodyning) or slightly different frequency (heterodyning) as the excitation light source. In practice the detector system will limit the maximum useful excitation frequency. Cross-modulation is normally used to transform the high frequency signal to a lower frequency domain because this improves noise immunity. All the noise picked up at high frequencies is simply filtered out. In the case of a fast PMT, the modulation frequencies will be limited to p/2 + fx (t < tx) or smaller than p/2 + fx (t > tx) will give rise to negative and positive values of Ion - Ioff, respectively. This scheme, however, is less suitable for absolute tf determinations because Ion - Ioff still contains constants related to the quantum efficiency, the concentration of the fluorescent material, and various instrumental parameters. This limitation can be overcome by recording three images: one with a detector modulation phase fx, one at fx + p/2, and one with no detector modulation. After subtracting the unmodulated image from each of the modulated images, the values at one pixel for the two detector modulation phases will be I(fx) - Ioff = 0.5 A MD MF cos(fF - fx)

(8)

I(fx + p/2) - Ioff = 0.5 A MD MF cos(fF - fx - p/2)

(9)

and

respectively. After taking the ratio of Eqs. 8 and 9 we obtain I=

cos(f F - f x ) = tan(f F - f x ) = wtF , x cos(f F - f x - p 2)

(10)

For fx = 0, Eq. 10 only depends on fF and the known modulation frequency w. Therefore, this mode of operation does provide fully quantitative tf imaging. However, in the case of multiexponential decay of the fluorescence, the measured tf will be concentration- and quantum-efficiency–weighted average lifetime over all the decay components. Consequently, a mixture of a number of fluorescent species with different lifetimes will yield only a single value, and truly selective imaging of one particular species is difficult. Alternatively, a large number of images recorded at different detector phase angles and modulation frequencies can be recorded. After a full analysis of each pixel, using a nonlinear, least-squares fitting procedure, the tf and relative contribution of each individual fluorescence decay can be resolved (Jameson and Gratton, 1983; Gratton et al., 1984) and the relative abundance of each species can be determined directly. However, because pointscanning confocal data acquisition times are on the order of 1 to 10 s/frame, the total data acquisition time may become long, and as the images are recorded sequentially, this method is sensitive to photobleaching. The effects of bleaching may be reduced by averaging a number of interleaved data sets, each one recorded using a different phase sequence. A significant amount of computational power is required for the analyses of complete images, and one must also consider the non-uniform response of the image intensifier. At high modulation frequencies, a difference in the modulation depth and the phase shift is observed between the center and the edge of intensifiers (Lakowicz and Berndt, 1991). This can be compensated for by a calibration method using a uniform reference specimen for which tf is known. Another way to reduce this effect is to use small diameter intensifiers.

Disk-Scanning Implementations Implementation of tf imaging based on phase fluorometry is straightforward when the method is combined with a spinning-disk

type of confocal microscope. Here, the light source can easily be modulated using EOMs or AOMs or by using a pulsed-laser system and any of the phase-sensitive detection schemes described above can be used.

Point-Scanning Implementations The phase-fluorometry–based method of tf sensing can be easily implemented in a point-scanning microscope. An example of a phase-fluorometry–based detection method in a scanning twophoton excitation microscope is given below. This work concerns one of the first lifetime imaging scanning microscopes (Piston et al., 1992). Here, a mode-locked dye laser producing femtosecond pulses at a repetition frequency of 75.6 MHz was used as a light source and a simple modulated PMT detector was employed, but the detection scheme was somewhat different from that discussed above. The detector was modulated at a frequency slightly higher than the excitation frequency and as a result the fluorescence is detected at Df, the difference frequency between the excitation and detection modulation frequencies. This cross-correlation detection scheme (heterodyning) transforms the signal to the low-frequency domain and assures high noise immunity. In order to obtain tf images, three images are acquired, each one with an additional 120° phase shift. Test measurements on rat basophilic leukemia cells, stained with the DNA indicator Hoechst 33342, were recorded with a total pixel dwell time of 18 ms (3 ¥ 6 ms/pixel). In the original paper describing this work, only fluorescence lifetime line profiles are shown. Based on the noise level in the lifetime traces, the accuracy in the lifetime is estimated to be 10%. Higher pixel dwell times will result in more accurate lifetime images. In Figure 27.3 the original lifetime images from which the line profiles were extracted are shown (image courtesy Dr. Dave Piston). Areas outside the nuclei yield low signal and therefore have lifetimes assigned as zero. A clear difference is visible between the lifetime calculated from the demodulation and that of the phase shift. This difference in lifetime is indicative of a multiexponential decay of the fluorescence. Another example of frequency-domain fluorescence lifetime imaging can be found in So and colleagues (1996). This implementation is also based on a multi-photon microscope. However, these approaches could easily also be implemented on a conventional CLSM.

Fluorescence Lifetime Sensing in the Time Domain Time-Domain–Based Methods Streak Camera Implementations In time-domain methods, the fluorescence intensity decay as a function of time is recorded after excitation with a pulsed light source. A good example of this is the combination of a picosecond pulsed laser for the excitation of the specimen coupled with a streak camera (O’Connor and Phillips, 1984). Here, the streak camera records the whole fluorescence decay curve after the specimen has been excited with a laser pulse.

Time Correlated Single-Photon Counting Implementations A more common way to record tf is time-correlated single-photon counting (TCSPC) (Lakowicz, 1983; O’Connor and Phillips, 1984). Here, the fluorescent molecules are excited using a brief

Fluorescence Lifetime Imaging in Scanning Microscopy • Chapter 27

521

FIGURE 27.3. Multi-photon phase-fluorometry–based FLIM image of rat basophilic leukemia cells, stained with Hoechst 33342.

light pulse after which the timing of single-photon emission is recorded. Using this approach the probability distribution for the emission of a single photon, and thus the fluorescence decay curve, is recorded. The high time resolution, 25 to 250 ps, and wide dynamic range of tf of this technique have made it popular for spectroscopic applications. A typical experimental geometry is depicted in Figure 27.4. The specimen is excited by the pulsed light source. A trigger pulse synchronized with the excitation light pulse is used to start a time-to-amplitude converter (TAC). The fluorescence emitted by the specimen is detected by a PMT, sent through a discriminator (DISC), and then used to stop the TAC. The output from the TAC will be proportional to the time difference between the start and stop pulses. The TAC output is now converted to a digital word by means of an analog-to-digital converter (ADC). This digital word is used as a pointer to an address in a histogramming memory. Finally, the value at this specific address is incremented. After repeating this process numerous times, a histogram of the fluorescence decay curve is recorded in the memory. The dead-time of the TAC electronics is comparatively long, typically 300 to 1000 ns. Therefore, care must be taken that the count rate of the experiment is sufficiently low to prevent pulse pile-up. The TAC usually operates in the reversed start–stop geometry. Here, the TAC is started by the fluorescence signal and stopped by

FIGURE 27.4. A schematic diagram of a TCSPC setup.

the laser trigger. In this way the TAC is only triggered by usable events, and not by laser trigger pulses that do not result in a detected fluorescence photon. Therefore, this mode of operation suffers less from dead-time effects. In the reversed start–stop geometry, pile-up is minimized by reducing the excitation intensity to about 1 to 5 detected photons per 100 excitation pulses. Furthermore, in spectroscopy applications excitation frequencies not exceeding 10 MHz are employed to ensure that the fluorescence decay signal from one excitation pulse is not affected by that of other excitation pulses. Therefore, the maximum count rate in conventional TCSPC is less than 100 kHz. The time required to transfer the decay curve from the histogramming memory to the computer system can be substantial. Instead of the TAC, time-todigital converters (TDCs) can be used. Here, the timing of events is determined by means of solid-state delay lines and directly translated into a digital word. In practice TDCs have very similar properties as TACs. In general, the decay curves recorded by TCSPC are fitted to a (multi) exponential decay. Here, usually the time response of the instrument is taken into account by employing an iterative deconvolution technique. In general, the use of conventional TCSPC equipment for imaging results in very long acquisition times. Conventional TCSPC equipment, however, has been employed in CLSM for fluorescence spectroscopy on discrete microscopic volumes (Ghiggino et al., 1992; Vanderoord et al., 1995) and for fluorescence lifetime imaging at a low acquisition speed (Bugiel et al., 1989). Operating the TCSPC detection system at too high detection rates, above 5% of the excitation frequency, results in distortion of the recorded decay curve (Lakowicz, 1999). More recently, TCSPC plug-in cards for PCs have been developed that are optimized for imaging applications. These cards have a much lower dead time than the conventional TCSPC electronics intended for use in spectroscopy (Becker et al., 2003; Kwak and Vanden Bout, 2003).

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Time-Gating Implementations Another approach to fluorescence lifetime imaging is based on combining pulsed excitation with time-gated detection. In timegated detection, use is made of one or more time channels, each with a different time offset with respect to the excitation pulse. Time gating can be implemented using both single-photon counting and analog detection methods. Here, we will restrict ourselves to implementations based on single-photon counting. In time-gated photon counting, the dead time of the detection electronics can be very low (sub-nanoseconds) and consequently such a detection system can be operated at very high count rates. In principle, multiple fluorescence photons can be detected for each excitation pulse. By sampling the fluorescence decay at different offsets with respect to the excitation pulse, information about the fluorescence lifetime can be obtained. In the early days of lifetime imaging often only two time-gated detection channels were used (see Fig. 27.5), each with a different delay time with respect to the excitation pulse. For a mono-exponential fluorescence decay, tf can be calculated from the ratio IA/IB of the intensities recorded in the two time windows A and B. t=

Dt log( I A I B )

(11)

with Dt the time difference between the start of the two time windows. This equation is only valid for windows of equal width and provided the excitation pulse is short compared to tf. It must be noted here that, although the fluorescence signal is convolved with the excitation pulse, the intrinsic fluorescence decay is observed after the excitation pulse has died out. The optimum gate width Dt for a particular tf amounts to 2.5 tf (Ballew and Demas, 1989). In two-channel, time-gated detection, only one decay time can be determined, and a multi-exponential fluorescence decay yields only a single effective tf. This limitation can be overcome in principle by using more detection windows. The time gates can be acquired in one single pass or in multiple passes. In the case of a multiple-pass implementation, all photons arriving at the detector when it is gated off are lost. As a result, sensing more channels reduces the efficiency of the system. If the time gates are all activated sequentially after each and every excitation pulse, a much better photon economy can be realized even when a large number of gates are employed.

Time-Domain Fluorescence Lifetime Imaging Widefield, Slit-Scanning, and Spinning-Disk Microscopy Implementations Several groups have been working on WF imaging systems that employ time-gated detection (see, e.g., Ni and Melton, 1991; Wang et al., 1991; Schneckenburger and Konig, 1992; Periasamy et al., 1996). Often, gated, image intensifiers are used with minimum gate times of several nanoseconds. This limits the use of these imaging systems to measuring fluorescence lifetimes greater than 1 ns. After the acquisition of two images, Eq. 11 can be used to transform the two intensity images into a tf image. Because the images are acquired sequentially, this approach is sensitive to photobleaching effects. Much faster gated, image intensifiers that allow sub-nanosecond time gating are also available (Scully et al., 1997). Using this type of system, complete decays can be sampled by recording sequences of images at different gate offsets. The efficiency of such systems goes down inversely with the number of time gates. Again, averaging schemes can be employed to compensate somewhat for the effect of photobleaching on the measurement of tf. A streak-camera–based lifetime imaging system was employed by, for example, Minami and Hirayama (1990). More recently, Krishann and colleagues (2003) employed a two-dimensional (2D) streak camera in combination with a slit-scanning multi-photon excitation microscope. Time traces were recorded for all the points imaged onto the entrance slit. The system can be employed for lifetime imaging with picosecond time resolution.

Point Scanning Implementations Most of the present implementations of FLIM in point-scanning microscopes are based on photon-counting techniques. Several commercial suppliers exist for both TCSPC PC plug-in cards and for fast time-gated photon counting detection systems (Van der Oord et al., 2001; Becker et al., 2003; Kwak and Vanden Bout, 2003).

TCSPC FLIM TCSPC PC plug-in cards have been optimized for high throughput data collection. Some of the cards have comparatively large onboard memory banks that remove the memory transfer bottleneck. Here, the maximum frame size is restricted by the amount of on-board memory. In other plug-in cards, the memory of the

Excitation

8 channel time-gating

Intensity

Intensity

2 channel time-gating

I1

I2

I1 I2

∆t1

∆t2

∆t1∆t2 ∆t3 Excitation

Time [ns]

FIGURE 27.5. Time gating.

I3

I4

I5

I6

I7

I8

∆t4

∆t5

∆t6

∆t7

∆t8

Time [ns]

Fluorescence Lifetime Imaging in Scanning Microscopy • Chapter 27

host computer is used and the timing histogram is built up in the computer memory. Furthermore, the dead time of the electronics has been reduced to 150 to 300 ns. Therefore, these TCSPC cards are considerably faster than conventional TCSPC electronics designed for spectroscopy applications (Becker et al., 2003; Kwak and Vanden Bout, 2003). The dead time of the TCSPC detection electronics is dominated by that of the TAC or TDC. FLIM systems based on this type of acquisition electronics typically employ a large number of time channels (typically 64–4096) that are comparatively narrow (25–200 ps). In principle, these FLIM systems are still based on the concepts used in the conventional spectroscopic TCSPC electronics. Therefore, they can in principle provide spectroscopic information as long as sufficient fluorescence signal is present. It should be stressed though that a large number of detected photons is required in order to realize spectroscopic accuracy. In spectroscopy experiments whole decay curves are recorded with as many as one million counts in the decay. This is far more than is available in fluorescence imaging experiments. Furthermore, care must be taken to not operate the TCSPC electronics at too high count rates. Operation of detection systems at a rate of one over the dead time at the input will result in a loss of 50% of the detection efficiency due to pile-up. Moreover, operation of TCSPC at too high count rates will result in distortion of the decays. True spectroscopic accuracy can only be realized when the detection count rate is below 5% of the excitation rate (O’Connor and Phillips, 1984; Lakowicz, 1999).

Time-Gated FLIM Our group has implemented time-gated detection in several pointscanning microscopes. Originally we employed a system based on two time gates (Buurman et al., 1992; Draaijer and Houpt, 1988). Here, the detection channel of an existing CLSM was modified to accommodate the time-gated detection technique in combination with single-photon counting detection. A fast (0.8 ns rise time) redsensitive PMT was followed by a fast discriminator with a 3 ns pulse pair resolution. The output signal from the discriminator was fed to two time-gated counter circuits that were both synchronized with the excitation source. All the time-critical electronics were built using emitter coupled logic (ECL) electronics. This detection circuit could be used at count rates of up to about 107 cps without serious pile-up effects visible in the lifetime. The minimum gate width of this system was approximately 2 ns and using this setup lifetimes of less than 1 ns could be imaged. An important advantage of using two time-gated detection channels, which are opened sequentially after every laser pulse, is the relative insensitivity to photobleaching. Furthermore, at the optimum gate width of 2.5 tf , the total acquisition time per excitation pulse amounts to 5 tf and approximately 99.3% of the total decay is captured within this period. More recently we investigated the performance of time-gating systems with 4 and 8 channels, respectively (Van der Oord et al., 2001; de Grauw and Gerritsen, 2001). Here, the same design philosophy was used. Again all the gates open sequentially after each and every excitation pulse. This approach ensures both high photon efficiency and a high maximum count rate capability. The dead time of the electronics is below 1 ns and therefore doesn’t limit the maximum count rate of the lifetime acquisition system. In practice the performance of such a lifetime imaging systems is dominated by the properties of the detector.

523

Comparison of Confocal Fluorescence Lifetime Imaging Methods Three detection methods have been used so far for recording fluorescence lifetime images in point-scanning microscopes: time gating, TCSPC, and phase fluorometry. In this section we will attempt to compare these approaches. Aspects that we take into consideration are the shortest observable lifetimes, the possibility of recording multiple lifetimes per pixel (multi-exponential decays), the sensitivity for bleaching, the photon economy, and the acquisition time. It must be stressed, though, that the limitations of all methods are to a large extent determined by the present stateof-the-art of the technology. The properties of the detector have a profound influence on the performance of the different lifetime imaging methods. Therefore, a separate section is included to summarize detector properties.

Shortest Lifetime The shortest lifetime that can be imaged using phase fluorometry is limited by the speed of the detector and the modulation frequency or pulse repetition frequency of the light source. Based on the experience in non-imaging experiments using the fastest detectors, lifetime imaging down to 0.2 ns should be feasible (Ven, personal communication). In the case of time-gated detection, the minimum lifetime is determined by the minimum gate width, the width of the excitation pulse, and the speed of the detector. Gate widths of less than 1 ns can be realized by the use of fast electronics. In combination with the fastest PMTs, the minimum observable lifetime is again estimated to be about 0.2 ns. The TCSPC systems can be operated with channel widths down to 10 to 40 ps. Therefore, the minimum observable lifetime in this type of system will be only limited by the timing properties of the detector and the laser pulse width rather than the performance of the detection electronics. In practice the fastest available detectors (MCP-PMT) exhibit a timing jitter (transit time spread, TTS) of about 25 ps (Hamamatsu, 1997). Accurate determination of lifetimes close to or shorter than the TTS is difficult. Therefore, the limiting lifetime for TCSPC–based systems will be in practice on the order of 50 to 100 ps when a fast detector is employed. Experience with conventional (TCSPC, phase fluorometry) fluorescence lifetime spectroscopy has shown that measuring short lifetimes (10-4 s Low

Fluorescence Lifetime Imaging in Scanning Microscopy • Chapter 27

maximum count rate is limited to about 1 to 3 MHz and their timing resolution is about 350 ps. The above figures can be used to make an estimate of the maximum frame rate for a specific type of detector. Assuming a 256 ¥ 256 pixel image and an average number of counts per pixel of 250, we find frame acquisition times of 3.2 to 1.6 s, 500 to 160 s, 16 to 8 s, and 16 to 5 s for the PMT, MCP-PMT, SPAD, and GaAs PMT, respectively. Very short lifetimes of about 25 ps can only be measured with a MCP-PMT at the price of long acquisition times. Fast acquisition speeds, in combination with a lower lifetime limit of a few hundred picoseconds, can be realized with fast PMTs. Interestingly, the GaAs PMT combines a lifetime limit comparable with that of fast PMTs, with high quantum efficiency. This makes this detector an interesting alternative for low light level FLIM. The acquisition time performance not only depends on the properties of the detector but also on the properties of the acquisition electronics. The dead time of the acquisition electronics is particularly important; large dead times will result in the loss of counts at high count rates. In lifetime imaging, the probability of a photon arriving at the detector usually follows a (multi) exponential distribution. Therefore, the probability of pile-up is non-uniform in time and peaks at time equals zero of the exponential function. Consequently, when pile-up takes place the recorded fluorescence decay is distorted and the fitted lifetimes show a count-rate dependency. In our experience, a pile-up of 5% to 10% does not significantly affect the recorded fluorescence lifetimes. For a detection system with a dead time of 10 ns, this corresponds to a maximum count rate of 5 to 10 MHz. This goes down to 250 to 500 kHz for a detection system with a dead time of 200 ns.

APPLICATIONS Multi-Labeling and Segmentation Fluorescence lifetime imaging is able to discriminate between a large number of probes based on lifetime differences alone. In Figure 27.8, an example of fluorescence-lifetime–based discrimination of fluorescent probes is shown. Here, a mixture of seven different fluorescent probes (Yellow Green beads, NYO beads, FITC

NR

YG

employed in scanning microscopes usually show very poor timing properties and transit-time spreads (TTS) of several nanoseconds are not uncommon. Therefore, such detectors are not usable for FLIM. Fast, head-on PMTs exhibit a TTS as low as several hundred picoseconds and are in general much better for timingcritical applications. Usually their quantum efficiency does not exceed 10% and they can operate at maximum count rates of 5 to 10 MHz. At higher count rates, pulse pile-up degrades their performance and they may even be damaged when the maximum anode current is exceeded. The shortest lifetimes that can be measured with a lifetime imaging system depend on both the timing properties of the detector and of the detection electronics. PMTs optimized for fast timing applications have a TTS as low as 200 to 400 ps. This limits the shortest lifetimes that can be measured with this type of tube to similar values. In order to measure shorter lifetimes microchannel plate PMTs (MCP-PMTs) (McLoskey et al., 1996) can be employed. These detectors have a quantum efficiency of about 10% and a TTS as low as 25 ps. At present the MCP-PMTs have the best timing performance for the measurement of fast decays. However, the MCP-PMTs are expensive and vulnerable and their maximum count rate is limited. For example, the popular Hamamatsu R3809U-50 MCP-PMT has a maximum average anode output current of 100 nA, corresponding to a maximum count rate of 100 to 300 kHz. However, Hamamatsu advises operating this device at less than a maximum count rate of 20 kHz (Hamamatsu, 1997). In general, all detectors exhibit a wavelengthdependent time response that complicates the accurate determination of short decay times (Lakowicz, 1999). Both PMTs and MCP-PMTs show a rather poor quantum efficiency of about 10%. Alternatively, single-photon counting avalanche photo diodes (SPADs ) or PMTs with a GaAs photo cathode can be used. The SPADs have high quantum efficiencies, about 70% at 650 nm, and can be used for single-photon counting applications at count rates of up to 1 to 2 MHz. Their typical timing jitter is specified by PerkinElmer to be about 350 picoseconds and no maximum value is specified. In practice we found that values on the order of 700 ps are more common. Moreover, the timing properties of these devices are count-rate dependent. These properties make the SPADs less suitable for lifetime imaging. The GaAs photocathodes, also have comparatively high quantum efficiencies of about 40% in the red part of the spectrum. Their

527

C

800

0

B

benzopyrylene

NYO

Lifetime

200

2 A

400

LY

6

FITC BB

Occurrence

600

2

3

4

5

6

7 8 Lifetime [ns]

9

10

11

12

C

FIGURE 27.8. (A) Intensity image of a specimen containing a number of different fluorescent beads and crystals. (B) Corresponding fluorescence lifetime image. (C) The histogram of fluorescence lifetimes.

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Chapter 27 • H.C. Gerritsen et al.

beads, Bright Blue beads, Nile Red beads, Lucifer Yellow solution, and benzopyrylene crystals) was imaged with a homemade two-photon fluorescence lifetime microscope equipped with an 8channel time-gated detector (Sytsma et al., 1998). In the lifetime image and in the lifetime histogram, six different lifetimes are clearly visible. The areas with the highest intensities yield the narrowest lifetime distributions. The fluorescence lifetimes of the beads were determined by means of TCSPC. The yellow-green beads and the Lucifer Yellow solution showed mono-exponential decays with decay times of 4.27 ns and 4.95 ns, respectively. The other beads and the crystal showed multi-exponential decays. The average lifetimes of these decays were 2.8 ns for NYO beads, 3.1 ns for FITC beads, 3.2 ns for Bright Blue beads, 3.6 ns for Nile Red beads, and 10.8 ns for the benzopyrylene. The lifetimes of FITC and Bright Blue beads are separated by only 0.1 ns and show up as one broadened peak in the histogram. The widths of peaks in the histogram are determined by statistics alone. Here, the minimum and maximum intensity per pixel amounted to 70 counts and 140,000 counts for the Lucifer Yellow solution and benzopyrylene, respectively. A higher signal per pixel improves the signal-to-noise ratio and, therefore, reduces the widths of the lifetime peaks in the histogram. Here, the time between two excitation pulses amounts to 12.2 ns (82 MHz Ti:Sa laser). Consequently, the lifetime sensitivity for the long lifetime benzopyrylene (10.8 ns) is poor. In Figure 27.9, confocal fluorescence intensity and fluorescence lifetime images from the alga Gymnodinium nagasakience are shown. The images were recorded with a confocal microscope equipped with a simple time-gated detection system with two, 2 ns-wide gates (Buurman et al., 1992). The alga was stained with an antibody–FITC conjugate against its outer membrane. The fluorescence intensity image in Figure 27.9(A) shows signal from both the antibody–FITC and the autofluorescence of the chlorophyll contained in the chloroplasts because no spectral discrimination was used to separate the fluorescence of these two substances. Figure 27.9(B) shows a fluorescence lifetime image of the alga; the gray value is a measure for tf. Chloroplasts show up as dark spots because for chlorophyll tf ª 0.7 ns, while the areas around the membrane are lighter because of the FITC tf ª 1.1 ns. The lifetime image can be segmented and used as a mask to obtain two separate images: one with fluorescence intensity representing chlorophyll [Fig. 27.10(A)] and the other with fluorescence intensity representing FITC [Fig. 27.10(B)]. For comparison, two images obtained by filtering the emission in the conventional way are also shown. Figure 27.10(C) shows the chlorophyll signal

A

B

A

B

FIGURE 27.9. Confocal fluorescence intensity (A) and fluorescence lifetime images (B) from the alga Gymnodinium nagasakience stained with antibody–FITC conjugate against its outer membrane (tf ª 1.1 ns). In addition, autofluorescence of chlorophyll is visible (tf ª 0.7 ns).

recorded through a long-pass 580 filter and Figure 27.10(D) shows the FITC signal detected through a bandpass filter. There is a high degree of correlation between the FLIM and spectral images. However, a subtle detail coming from a dimple in the center of the alga shell shows up in both members of the spectrally resolved image pair showing that this filter combination is not good enough to discriminate between chlorophyll and FITC. This detail is not visible in the lifetime image.

Ion-Concentration Determination Ion concentrations in biological systems, such as pH, Ca2+, and Na+, are widely studied with fluorescent probes. The probes have a high selectivity for specific ions and exhibit marked changes in their photophysical properties upon binding. Often the probes exist in two states, bound to the ion and free, that have distinct fluorescence lifetimes. The average fluorescence lifetime is a good measure for the ion concentration and can be used for the quantitative imaging. Some of the examples obtained in our lab are discussed below. Others can be seen in Chapter 42, this volume.

C

D

FIGURE 27.10. The chlorophyll (A,C) and FITC (B,D) signals after fluorescence lifetime–based segmentation (A,B) and segmentation based on differences in emission spectra (C,D).

Fluorescence Lifetime Imaging in Scanning Microscopy • Chapter 27

Calcium Imaging Calcium plays a central role as a second messenger in plant and animal cells and is involved in controlling numerous biological processes. Fluorescent probes are widely used in macroscopic and microscopic studies of [Ca2+]. These probes have a high selectivity for free Ca2+ over other ions and the probes undergo a significant change in fluorescence properties upon binding to Ca2+. Current fluorescent calcium probes can be divided in two groups: ratio or dual-wavelength probes and non-ratio or single-wavelength probes. Due to calibration problems, single-wavelength probes produce very ambiguous results. The fluorescence intensity increases as a result of an increase in [Ca2+]. However, the fluorescence intensity not only depends on the [Ca2+] but also on the dye concentration, which can vary as a result of unequal cytosol thickness, inhomogeneous dye distribution within or between cells, and leakage and/or photobleaching during an experiment. Quantification of [Ca2+] can be more accurate when ratio probes are used. These probes show a marked shift in either excitation or emission spectra upon binding to Ca2+ and are referred to as dual-excitation or dual-emission probes. This feature allows these probes to be used for ratio analysis (Rink et al., 1982; Grynkiewicz et al., 1985; Tsien and Poenie, 1986). The ratioing method is independent of dye distribution inhomogeneities within or between cells, and leakage and/or photobleaching during an experiment. Unfortunately, a laborious in situ calibration procedure is required when ratio probes are being used in fluorescence microscopy. Most of the commercially available ratio probes such as Fura-2 and Indo-1 have to be excited in the ultraviolet (UV). UV wavelengths can potentially injure cells and tend to excite autofluorescence; therefore, it is preferable to use probes that have their excitation wavelength in the visible. FLIM is an interesting alternative for the imaging of free Ca2+. The binding of Ca2+ to the single-wavelength probes results in a large increase of the quantum efficiency of the probe. This is in general accompanied by a substantial increase of the fluorescence lifetime of the probe. Depending on the free calcium concentration and the Kd of the probe, different amounts of bound and free probe will be present in the specimen, Kd = ([free probe]· [ Ca2+])/[bound probe]. In general the fluorescence decay will be (at least) bi-exponential. The amplitudes of the different lifetime components depend on the calcium concentration and it turns out that the average lifetime is a good measure of the calcium concentration (Sanders et al., 1994).

A

FIGURE 27.11. Intensity image (A) and fluorescence lifetime image (B) of an Indo-1 stained beating rat myocyte. The beating, approximately 2 Hz, results in the bands in the image. The 256 ¥ 256 pixel image was recorded in 1 s using a two-photon excitation microscope equipped with a time-gated detection system.

10 µm

529

Interestingly, in principle FLIM allows quantitative imaging of Ca2+ using single wavelength probes that are excited with visible light. There are several indications that this approach is more reliable than the approaches based on ratio imaging (Sanders et al., 1995; Herman et al., 1997). In general, the free probe has both a short lifetime and low quantum efficiency. The ion-bound probe, on the other hand, has both a long lifetime and high quantum efficiency. Therefore, the ion-bound form dominates the fluorescence decay and the usable ion concentration range of the probe is shifted to lower ion concentrations. Early work on the quantitative FLIM of calcium required comparatively long acquisition times. Recently, we carried out fast-time-gating-based FLIM measurements of Indo-1 stained rat myocytes (see Fig. 27.11) (Gerritsen et al., 2002). The rat myocytes spontaneously beat at rates of 0.5 to 2 Hz at 37°C and the beating is accompanied by large Ca2+ fluxes (Berlin and Konishi, 1993). The images were recorded in a homemade twophoton excitation microscope (Sytsma et al., 1998) equipped with an 8-channel time-gated detector and a femtosecond titanium:sapphire laser operating at 82 MHz and an excitation wavelength of 700 nm. Indo-1 exhibits a 1.4 ns lifetime for the free probe and a 1.66 ns lifetime for the ion-bound probe (Lakowicz and Szmacinski, 1993). Despite the small lifetime difference between the free and bound forms of the probe, lifetime images (256 ¥ 256 pixels) could be recorded at a rate of 1 frame per second. Here, a Hamamatsu R1894 PMT was used at an average count rate close to 10 MHz. Both the intensity [Fig. 27.11(A)] and the lifetime image [Fig. 27.11(B)] were median-filtered. The edges of the lifetime image are somewhat sharpened due to the thresholding, lifetimes were only calculated for pixels with 50 or more counts. Pixels with fewer counts were set to zero. The calibration bar below the lifetime image gives an indication of the lifetimes. No attempt was made to convert the lifetimes into calcium concentrations. The beating of the myocyte during image acquisition shows up as a band in both the FLIM and the intensity image. Here, the myocyte beat about twice during the image acquisition.

pH Imaging Dental biofilm can be several hundred micrometers thick and exhibits an intricate three-dimensional (3D) structure (see also Chapter 51, this volume). This type of specimen is strongly scattering and two-photon excitation (TPE) microscopy is therefore the preferred imaging method to study it. pH is one of the key

B

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Chapter 27 • H.C. Gerritsen et al.

4.0

τ [ns]

3.5

3.0

2.5 2

3

4

5

6

7

8

pH FIGURE 27.12. The pH calibration (carboxy–fluorescein) of the two-photon FLIM system.

parameters in biofilm research. The pH in the biofilm is in the acidic range and quantitative imaging of the pH in this range is usually accomplished by means of excitation-ratio imaging. Implementation of excitation ratio-imaging in TPE imaging is hampered by the requirement of two measurements at different excitation wavelengths. Fluorescence lifetime imaging is an excellent alternative for the quantitative imaging of pH in the acidic range. Here, we employed TPE in combination with FLIM to study the pH in a biofilm (Bradshaw et al., 1998; Vroom et al., 1999). Carboxy–fluorescein was used as a fluorescent pH indicator. The carboxy form of the probe is cell impermeable, so it senses only the extracellular pH, the pH of interest in the biofilms. TCSPC measurements on a series of carboxy–fluorescein buffers of different pH revealed a clear pH-dependent average fluorescence lifetime of 4.0 ns at high pH (pH > 7) and 3.0 ns at low pH (pH < 3). In addition to the fluorescence lifetime effect, the quantum yield of carboxy–fluorescein went down to 5% at pH 3, relative to that at pH 7. At an excitation wavelength of 800 nm, the biofilms show some autofluorescence with an average fluorescence lifetime of

A

B

FIGURE 27.13. Intensity image of biofilm (A) with individual bacteria visible. Fluorescence lifetime image (B) shows a homogeneous pH of 6.2 ± 0.3.

less than 1 ns. The difference in fluorescence lifetime between the autofluorescence and the carboxy–fluorescein was exploited to suppress the autofluorescence in the pH measurements. This was achieved by opening the first gate 1 ns after the excitation pulse. This suppresses 80% of the autofluorescence and only 20% of the carboxy–fluorescein signal. In addition, a comparatively high probe concentration of 50 to 100 mM was used in the pH imaging experiments. The autofluorescence was less than 1% of the total fluorescence intensity in all the measurements, even at low pH. In Figure 27.12, the pH calibration of the microscope is shown. Intensity and pH xy-images (30 ¥ 30 mm2, z = 60 mm) of carboxy–fluorescein stained biofilm are shown in Figure 27.13. The intensity image [Fig. 27.13(A)] shows individual bacteria, while the fluorescence lifetime image [Fig. 27.13(B)] shows an almost homogeneous pH of pH 6.2 ± 0.3. At a constant pH the quantum efficiency of the carboxy–fluorescein is constant. Therefore, the heterogeneous fluorescence intensity distribution can be attributed to a non-homogeneous probe distribution. Binding of some of the probe to the bacteria may cause this. A pH gradient was induced by overlaying the specimen with a 14 mM sucrose solution. The fermentation of sucrose lowers the pH outside the bacteria. In Figure 27.14, the intensity [Fig. 27.14(A)] and fluorescence lifetime [Fig. 27.14(B)] images 94 min after the addition of the sucrose are shown. These images were recorded at the same position as that of Figure 27.13. The fluorescence intensity image shows some brightly fluorescing areas. At the same position in the fluorescence lifetime image, a pH is found of less than 3. This observation is somewhat unexpected because the quantum efficiency of the probe goes down with pH. It suggests that a high local probe concentration is present at the bright spots. This effect may be caused by probe precipitation and makes the measurements at these locations less reliable. Therefore, the bright spots were excluded from the pH analysis. The average pH of this image is 5.2 ± 0.4, one pH unit lower than the average pH of the reference image.

Probes for Fluorescence Lifetime Microscopy A literature study and some measurements on probes in our own laboratory revealed a large list of probes which can potentially be used for fluorescence lifetime imaging, this list is summarized in Table 27.3.

A

B

FIGURE 27.14. Intensity (A) and lifetime (B) images after overlaying the biofilm specimen with a 14 mM sucrose solution. The average pH is 5.2 ± 0.4.

Fluorescence Lifetime Imaging in Scanning Microscopy • Chapter 27

531

TABLE 27.3. Ion Sensitive Probes PH Probes Probe SNAFL-1 Carboxy-SNAFL-1 Carboxy SNAFL-2 Carboxy SNAFL-3 Carboxy SNARF-5 SNARF-6 Carboxy SNARF-6 SNARF-1 SNARF-2 Carboxy SNARF-1 Carboxy SNARF-1 Carboxy SNARF-2 2-Naphtol Acridine Virginiamycin Sc BCECF DM-NERF Carboxy-fluoresceind Cl-NERF Acridine Orange

tacida (ns)

tbasea (ns)

Reference

295 /500 295a/500b 295a/500b 295a/500b 295a/500b

3.58 3.44 4.19 3.63 4.21

1.14 0.95 0.87 1.16 0.73

295a/500b 295a/500b 295a/500b 295a/500b 295a/500b 295a/500b 313/360

4.50 0.46 0.30 0.60 0.60 0.27 0.80 (0.35) 7.30 (1.91) 9.6 0.48 3.0 2.3 3.0 1.3 1.8

1.04 1.54 1.75 1.32 1.32 1.67 4.82

Whitaker et al. (1991) Whitaker et al. (1991) Whitaker et al. (1991) Whitaker et al. (1991) Whitaker et al. (1991) Whitaker et al. (1991) Whitaker et al. (1991) Whitaker et al. (1991) Whitaker et al. (1991) Whitaker et al. (1991) Whitaker et al. (1991) Whitaker et al. (1991) Laws and Brand (1979)

lexc/lem (nm) a

b

348/560 330/420 490/520 490/520 490/520 490/520 490/520

31.1 1.9 3.8 4.0 4.0 4.0 5.3

Gafni and Brand (1978) Clays et al. (1991) This laboratory This laboratory This laboratory This laboratory This laboratory

Ca2+ Probes Probe

lexc/lem (nm)

tno Ca++a (ns)

tCa++a (ns)

Fura-2

340/420 380/420

0.77(0.49) 1.5(0.51)

1.77

Keating et al. (1989)

488/520

1.6

3.5

Lakowicz et al. (1994) This laboratory

488/520

2.1

3.6

This laboratory

488/520

0.5

3.4

This laboratory

488/520

1.5

3.7

This laboratory

Reference

Quin-2 Calcium green Bapta 1 Calcium green Bapta 2 Calcium green Bapta 5N Oregon Green Bapta 1,2 Oregon Green Bapta 5N BTC Indo 1

488/520

0.5

2.8

This laboratory

340/400–475

0.7 1.4

1.2 1.7

Indo 5F

340/400–475

1.4

1.4

This laboratory Lakowicz and Szmacinski (1993) This laboratory

Cl Probes Probe SPQ/Cl-

lexc/lem (nm)

tnoCla (ns)

tCla (ns)

—/—

26.0

2.0

Reference Dix and Verkman (1990)

DNA or RNA Selective Probes Lifetime Differences with Free Probe and Probe Bound to DNA lexc/lem (nm)

lfreea (ns)

tbounda

Proflavine, acriflavine, acridine yellow (208C)

—/—

4.5

4 (0.7)

9-Aminoacridine

—/—

Rivanol

—/—

6.5

Quinacrine

—/—

4

532/— 488/520 488/520 488/>630

1.7 — — —

Probe

Ethidium bromide Acridine orange, DNA RNA RNA

15

7 (0.3) 10.5 (0.7) 31 (0.3) 5.5 (0.7) 13 (0.3) 3.5 (0.7) 19 (0.3) 24.2 1.7–1.9 1.7–1.9 5–20

Reference Duportail et al. (1977)

Duportail et al. (1977) Duportail et al. (1977) Duportail et al. (1977) Atherton and Beaumont (1984) Marriott et al. (1991b) Marriott et al. (1991b) Marriott et al. (1991b) continued

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Chapter 27 • H.C. Gerritsen et al.

TABLE 27.3. Ion Sensitive Probes (Continued) Lifetime Differences with Different Amounts of A + T Probe

lexc/lem (nm)

Rivanol

—/—

Quinacrine

—/—

Quinacrine

435/516

tmuch ATa (ns) e

5.5 (0.8) 13 (0.2) 2.2 (0.9) 15 (0.1) 2.6 (0.35)f 11 (0.39) 27 (0.26)

tlittle ATa (ns)

Reference

11

Duportail et al. (1977)

16

Duportail et al. (1977)

1.1 (0.06) 7.90 (0.33) 26 (0.60)

Arndt-Jovin et al. (1979) Arndt-Jovin et al. (1979) Arndt-Jovin et al. (1979)

Probes Which Are Sensitive to Other Factors as Ions or DNA or RNA Probe SR33557 Calcium— entry blocker

lexc/lem (nm) H2O/EtOH [50/50] EtOH Erythrocytes Egg PC vesicle

Fluoranthene

FITC FITCtransferin FITC-KE2

tfreea (ns)

tbounda (ns)

336/400

4

Chatelain et al. (1992)

336/403 —/—

8.5 13.3 (0.91) 2.9 (0.09) 13.6 (0.81) 5.0 (0.19) 51 51 51 3.9 1.5 (0.38) 3.6 (0.62) 1.32 (0.28) 4.11 (0.72)

Chatelain et al. (1992) Chatelain et al. (1992)

—/—

In N2 gas In air In O2 gas pH 7.2 pH 7.4

355/— 32 14 490/520 490/520

pH 7.4 Fab complex

485/—

Reference

Chatelain et al. (1992) Ni et al. (1991)

This laboratory This laboratory Matko et al. (1992)

lexc (nm), excitation wavelength; lem (nm), emission wavelength; tf, fluorescence lifetime, a is the amplitude of the respective components in case of multi-exponential decay. a Absorption maxima between 480 and 580 nm. b 500 nm long-pass filter. c Virginiamycin S has three different pKs and thus four different protolytic forms. d Affected by other effects such as binding. e 28% A + T. f 55% A + T.

SUMMARY Fluorescence lifetime imaging is a powerful tool that can be used for a large number of different applications. Since its introduction around 1990, it has significantly matured and at present several companies supply complete FLIM detection systems. The advantages and disadvantages of FLIM and the different implementations are summarized below.

•

• Fluorescence lifetime imaging is insensitive to intensity effects

•

• •

such as shading, photobleaching, absorption, and light source noise. This can be an important advantage, especially in confocal and multi-photon studies of thick specimens. Here, absorption effects and photobleaching are important limitations. Fluorescence lifetime imaging is a powerful tool in the field of ion-concentration imaging. The comparison between emission-ratio imaging and lifetime-imaging suggests that lifetimeimaging can yield quantitative results without a cumbersome in vivo calibration procedure. In theory, lifetime imaging is able to separate the signals from a large number of different probes in parallel using a single detector. Image-processing can be simplified by using fluorescence lifetime data as the basis of segmentation. In recent years it has been demonstrated that FLIM is an excellent and straightforward method to carry out FRET experi-

•

•

ments. The simple calibration of FLIM/FRET experiments is particularly appealing (Fig. 27.15). The image acquisition times of lifetime-imaging microscopes strongly differ. Potentially, the phase-fluorometry–based technique combined with pulsed excitation has the shortest acquisition times due to its ability to cope with large signal intensities. The phase technique employing sinusoidally modulated excitation, however, will require very long acquisition times when combined with scanning microscopy. The acquisition time of the time-gated or phase techniques in combination with pulsed lasers will be at least one order of magnitude shorter than in TCSPC. In principle the photon economy of all methods employing pulsed excitation is excellent. The photon economy of phase of fluorometry with sinuoidal excitation, however, is poor. Furthermore, the photon economy may vary per implementation. For example, systems with a long electronics dead time (TCSPC) or slow detectors will show a reduction of the photon economy (detection efficiency) above a certain count rate. At present the systems equipped with TCSPC are best suited for the imaging of very short lifetimes, provided that they are equipped with a fast detector (MCP-PMT). This type of detector should, however, be operated at very low count rates (50–200 kHz).

Fluorescence Lifetime Imaging in Scanning Microscopy • Chapter 27

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Figure 27.15. A FRET-FLIM experiment involving living HER14 cells was designed to determine if the EGF receptor (labeled with EGF-Alexa-488) co-localized with the lipid raft marker CTB (labeled with Alexa-594). The top row shows donor fluorescence intensity images and the bottom row donor lifetime images (256 ¥ 256 pixel). The graph on the right plots a histogram of the tf values derived from pixels in the regions of interest (gray boxes) under the conditions noted. (A) The donor-only lifetime image (lower) yields an average lifetime of 4.0 ns (yellow curve). (B) In the presence of the acceptor, FRET reduces the donor lifetime to about 3.5 ns (lower image and blue curve). (C) After photobleaching the acceptor, the donor lifetime (lower image and red curve) returns almost completely to the donor-only lifetime (experiment on a different cell).

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