Confocal Luminescence Lifetime Imaging with Variable Scan Velocity

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Confocal luminescence lifetime imaging with variable scan velocity and its application to oxygen sensing Zdenek Petrasek, Juan M. Bolivar, and Bernd Nidetzky Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.6b03363 • Publication Date (Web): 30 Sep 2016 Downloaded from http://pubs.acs.org on October 5, 2016

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Analytical Chemistry

Confocal luminescence lifetime imaging with variable scan velocity and its application to oxygen sensing Zdenˇek Petráˇsek∗,† , Juan M. Bolivar† and Bernd Nidetzky∗,†,‡ †

Institute of Biotechnology and Biochemical Engineering, Graz University of Technology, NAWI Graz, Petersgasse 12, A-8010 Graz, Austria ‡ Austrian Centre of Industrial Biotechnology, Petersgasse 14, A-8010 Graz, Austria ∗ ∗

Fax: +43 316 873 8434. E-mail: [email protected] Fax: +43 316 873 8434. E-mail: [email protected]

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Abstract The dependence of the luminescence lifetime on the probe environment is the basis of a range of sensing techniques. The major advantage of using the lifetime as the sensitive parameter is its independence on the probe concentration. However, the instrumentation for lifetime measurements is complex, generally requiring time-resolved excitation and detection. Here we present a simple method for the measurement of luminescence lifetimes on the microsecond scale based on variable excitation time determined by the scanning velocity. The technique is implemented in a confocal laser scanning microscope (CLSM), thus allowing not only simple lifetime measurement but also phosphorescence lifetime imaging. Since the method exploits the spatio-temporal dependence of sample excitation in a CLSM, there is no need for a pulsed or modulated light source or for additional time-resolved detection. The method can be realized in a standard CLSM without any modifications. The principle is demonstrated on oxygen sensing by collisional quenching of an oxygen-sensitive ruthenium(II) complex.

Keywords: luminescence, phosphorescence, lifetime imaging, oxygen sensing, confocal laser scanning microscopy

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Introduction The environmental dependence of the intensity, spectrum, lifetime or polarization of fluorescence and phosphorescence is the basis of a broad range of noninvasive techniques for studying molecular interactions and sensing the molecular environment 1–3 . The lifetime of the excited state is a particularly attractive parameter, as it is largely independent of the probe concentration and can therefore be employed also in situations where the control of the probe concentration is difficult, such as in microscopy 4,5 . While the lifetimes of fluorescent dyes lie usually in the nanosecond range, the phosphorescence lifetimes are often much longer, typically in the range of microseconds to milliseconds. The long lifetime of the emissive state of phosphorescent probes means that the dye has more time to interact with its environment, resulting in increased sensitivity 6 . For example, in case of collisional quenching much lower concentration of quencher can be detected than in case of a short-lifetime fluorescent probe. One of the most widespread applications is oxygen sensing 7–9 , employing long-lived ruthenium complexes 10 or platinum and palladium-containing porphyrins with long luminescence lifetimes 11,12 . Other applications of long-lifetime probes include iridium probes for various ions and small molecules 13 , and probes containing lanthanide ions (Eu3+ , Tb3+ , Yb3+ , . . . ) as sensors for pH, as ion hydration probes exploiting quenching of lanthanide luminescence by coordinated water molecules, and as energy donors in probes based on resonance energy transfer 14,15 . Lifetime-based sensing is often combined with imaging. For example, the imaging of oxygen is highly important in different branches of life sciences and medicine, including the study of the oxygen concentration in brain tissue 16 , in the eye retina 17 , in tissue during wound healing 18,19 and in tumors 20 . The measurement of oxygen distribution and consumption plays an important role in biotechnology and biocatalysis 21,22 . Oxygen sensing is furthermore the functional principle of pressure-sensitive paints used in fluid mechanics and aerodynamics 2 . Temperature-sensitive photoluminescence is the basis of temperature measurements of turbine blades 23 and other industrial thermometry applications 24 . Temperature-sensitive lifetimes of gold nanoclusters enabled intracellular nanothermometry in biological systems 25 . Other applications include imaging of defects in silicon for solar cells in photovoltaics 26,27 , and lifetime multiplexing with luminescent Tm- and Yb-doped nanocrystals 28 , with possible applications in data storage and security. Luminescence lifetimes can be determined by a range of methods of varying degree of complexity. While the advanced methods, such as time-correlated single-photon counting (TCSPC), provide fluorescence and phosphorescence decays with high resolution that can be subjected to a detailed analysis and reveal possibly complex kinetics 29–31 , there is an interest in technically simple and robust methods for fast measurement of monoexponential decays described by a single lifetime. One such method is rapid lifetime determination (RLD) 32–34 , which requires two luminescence intensity values measured in two different time windows following the excitation pulse to calculate the lifetime. This can be realized by using a pulsed light source (LED, laser diode) and a time-gated detector (gated CCD). Dual lifetime referencing (DLR) is an interesting approach employing the frequency-domain lifetime measurement to detect intensity changes of the sensor dye 35,36 . Its principle is the frequency-modulation method applied to a mixture of an analyte-insensitive dye with a long lifetime and an analyte-sensitive dye with a lifetime in the nanosecond range. The changes in the emission intensity of the sensitive dye are then detected as an effective change of the lifetime of the mixture of the two dyes. A similar referencing principle of using two dyes with their lifetimes on different time scales can be applied in the time domain, leading to time-domain dual lifetime referencing (t-DLR) 37 , and in combination with RLD to a technique simultaneously detecting two analytes 38,39 . The above-mentioned methods are typically combined with imaging to achieve two-dimensional mapping of lifetimes 40 . While employing a CCD camera results in wide-field 2D imaging, many applications would benefit from or require 3D resolution, as provided by confocal microscopy. Combination of lifetime imaging and 3D resolution can be realized by TCSPC 29–31 or by frequency modulation approach 41 . Recently, two methods for confocal imaging of long luminescence lifetimes, not requiring additional instrumentation for high temporal resolution such as TCSPC, have been suggested. One method employs the FRAP (fluorescence recovery after photobleaching) function

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of the microscope, but is rather slow and applicable only to lifetimes on the millisecond scale or longer 42 . The other method uses confocal pinhole shifted to at least two different positions to achieve the same effect as time gating, but can be implemented only in instruments that allow pinhole shifting 43 . Here we introduce a method to perform lifetime imaging in a common commercial CLSM based on a different principle, without requiring any additional modification or adjustment of the microscope. It takes advantage of the short time of excitation by a moving laser focus in a CLSM, and is based on measuring the relaxation of populations between the ground and the excited states upon the start of the excitation. The kinetics of this relaxation depend on the excitation and the emission rates. Consequently, for luminescent probes where the time scale of these processes lies in the same range as the illumination time by the scanning beam, the luminescence intensity will depend on the scanning velocity. We show that by analysing the intensities at two or more scanning velocities, the luminescence lifetimes can be determined, and lifetime imaging performed. We illustrate the principle on Ru(dpp)3 dye and show an application for oxygen sensing.

Experimental Methods Samples Tris-(4,7-diphenyl-1,10-phenanthroline)ruthenium(II) dichloride, in short Ru(dpp)3 , was purchased from ABCR GmbH (Karlsruhe, Germany). Phenyl sepharose CL-4B beads were obtained from GE Healthcare Europe GmbH. Lactate oxidase from Aerococcus viridans was a gift from Roche Diagnostics GmbH (Wien, Austria). L-(+)-lactic acid, and all other chemicals were obtained from Sigma-Aldrich, unless otherwise stated. We have shown in previous work that stable labeling of hydrophobic materials with Ru(dpp)3 can be achieved solely through dye adsorption by the carrier 44–46 . A stock solution of Ru(dpp)3 in ethanol (5 mg/ml) was prepared. A mixture of about 5 mg of Ru(dpp)3 per 1 g of wet beads was incubated in 50 mM potassium phosphate buffer (pH 7) while mixing in an end-over-end rotator (20 rpm). After a 1 h incubation at room temperature, the beads were washed thoroughly with 50 mM potassium phosphate buffer, pH 7.0, and stored at 4◦C until further use. No Ru(dpp)3 leakage occurred upon transferring the labeled carriers to buffers (e.g. 50 mM potassium phosphate, pH 7.0) containing up to 2 M NaCl. Immobilization of lactate oxidase follows the principle of hydrophobic adsorption. Native or luminescently labeled carriers were first washed with 50 mM potassium phosphate buffer, pH 7.0. Then, 1 ml of protein solution containing between 5 units and 50 units of lactate oxidase was incubated with 100 mg of wet carrier in 50 mM potassium phosphate buffer containing 2.0 M of NaCl, pH 7.0. After two hours of incubation, the immobilizate was washed with loading buffer and stored at 4◦C until further use. To follow the progress of immobilization during incubation, samples of supernatant were withdrawn and activity and protein concentration determined. The activity of lactate oxidase was measured by following the oxygen consumption in 50 mM L-lactate solution in air-saturated 50 mM potassium phosphate pH 7 at 25◦C. Liquid flow was delivered from a New Era NE-1000 syringe pump (Next Advance, NY, USA). Teflon tubings (250 µm diameter) and connection parts were purchased from Micronit Microfluidics (Enschede, The Netherlands) and Ibidi GmbH (Martinsried, Germany). The microscopy flow chamber µ-Slide VI 0.4 (Ibidi GmbH) had a channel size of 0.4 mm × 3.8 mm × 17 mm (H×W×L). Microscopy The microscopy experiments were performed on a Leica TCS SPE confocal laser scanning microscope. The samples were excited at 488 nm and the emission was detected in the range 500–700 nm. In all experiments, the water immersion objective HCX APO LU-V-I 20×/0.50 W was used. The pinhole size was always set to one Airy unit. The image size was typically 512 × 512 pixels; for the lifetime imaging analysis 4 × 4 pixels were binned to reduce the noise. The scan velocity is determined by two parameters that can be controlled by the user: the scan frequency and the magnification. The microscope used provides for only three different scan

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frequencies: 400, 600 and 800 Hz. Thus, for a given magnification selected by the continuous zoom setting, only three scan velocities are possible. The actual values of these velocities then depend on the selected magnification. In order to achieve a broader range of different scan velocities, we had to combine different frequency and zoom settings. Employing all three possible scan frequencies and several zoom values between 1 and 5 we obtained 15 scan velocities in the range 0.15–1.12 m·s−1 . Using different magnifications does not present any problem for the measurements on homogeneous solutions, but would complicate measurements on beads or any other structured sample. Therefore, in the imaging experiments with beads, we used used a single magnification, with the field of view of 183 µm, and the frequencies 400 and 800 Hz, resulting in the scan velocities 0.25 m·s−1 and 0.50 m·s−1 . We have noticed that the scan velocity is not constant across the whole field of view, but decreases towards the edges. This is very likely caused by the fact that the scanned beam slows down before it reaches the end of line and turns back to return to start scanning the next line (one-directional scanning was used). If not taken into account, this effect would cause artificial variation of the luminescence lifetime across the field of view. We have therefore applied the following correction: a homogeneous solution of Ru(dpp)3 was imaged with the two scan velocities, as described above. The rate k1 was calculated in the center of the image using the nominal values of the scan velocities. Subsequently, this value of k1 was used to calculate the corrected scan velocity in all all other parts of the image. The resulting map of the corrected scan velocity was then used for the calculation of the lifetime images of the beads. This type of correction should not be necessary with new laser scanning microscopes which are equipped with linear rather than sinusoidal scanning, resulting in a constant velocity scan. For calibration purposes (the factor α in eq. 6) it was assumed that the lifetime of Ru(dpp)3 in air-equilibrated ethylene glycol is τ = 2.3 µs 47 . The samples measured in the absence of flow were mounted in a well made of a microscope slide and a coverslip separated by a spacer (thickness 90–180 µm) with a circular hole of ∼10 mm diameter. For the experiments under flow, between 20-40 mg of labeled phenyl sepharose beads were packed in the microscopy flow chamber. Depending on the experiment, different mixtures of airsaturated 50 mM potassium phosphate buffer, pH 7, were flowed through the packed bed: only the buffer, the buffer containing 50 mM of L-lactate, or the buffer containing 50 mM of L-lactate and 50 unit/ml of lactate oxidase.

Theory We consider a simplified system of energy levels of the luminescent molecule with a ground state and one excited state (Fig. 1A). The luminescence at any given moment is proportional to the population of the molecules in the excited state. Let’s assume that at the onset of excitation (t = 0) with light of constant intensity all molecules are in the ground state. The steady state of the populations of the ground state and the excited state is reached only after a certain time from the onset of excitation (Fig. 1B). This time is determined by two rate constants: the rate constant of excitation k0 , which depends on the excitation intensity, and the decay rate constant of the excited state k1 , which depends on the radiative luminescence lifetime and the rates of all non-radiative processes, including collisional quenching by the analyte molecule. The temporal evolution of the population of the excited state y(t) is described by the following differential equation: dy/dt = k0 (yT − y) − k1 y, where yT is the total number (or concentration) of the luminescent molecules. Assuming that there are no molecules in the excited state at time t = 0, the solution is (Fig. 1B): y(t) = yT

k0 1 − e−(k0 +k1 )t k0 + k1

(1)

When the excitation intensity is chosen sufficiently low, so that k0 k1 , the rate of reaching the steady state depends practically only on the emission rate constant k1 . The condition k0 k1 is equivalent to operating in a non-saturating regime; that is, the situation when the steady-state

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A

excited state

excita�on

emission

ground state

B

0 Figure 1: The evolution of the populations of the ground and the excited states after the onset of excitation. A: the molecules are excited with the rate k0 and decay back to the ground state with the rate k1 . B: Initially, all molecules are in the ground state; with the start of excitation at time t = 0 the population of the excited state y(t) approaches a steady state with a time constant 1/(k0 + k1 ). population ys = yT k0 /(k0 + k1 ) (as follows from eq. 1) is small compared to the total number of molecules yT . Monitoring how fast the steady state of y(t) is reached upon excitation provides an alternative way to determine the decay rate constant k1 , or, equivalently, the luminescence lifetime τ = 1/k1 . In contrast, the common time-domain methods for lifetime determination monitor the decay of the excited state after the end of the excitation period. One possibility to utilize the approach towards the steady state described by eq. 1 and Fig. 1B for lifetime determination is to record time-resolved luminescence while exciting the sample. This would, however, require a detector with high temporal resolution. Another possibility, which is the basis of the method presented here, is to measure the average luminescence during excitation pulses of variable duration. The average fluorescence is low for excitation pulses shorter than the time needed to approach the steady state, and increases until it reaches a higher level for excitation pulses much longer than the characteristic time needed to reach the steady state: 1/(k0 + k1 ). The measurement of average luminescence for variable excitation times is easily realized in a confocal laser scanning microscope (CLSM), and can thus be applied not only to measure luminescence lifetimes but also to perform lifetime imaging. In a CLSM, the focused excitation beam moves across the sample, illuminating any location only for a short time. The emitted light is detected simultaneously from the same location. This is shown schematically in Fig. 2, where for simplicity the spatial profile of the excitation and detection is approximated as a rectangle with a width 2a and 2b, respectively. The location that has just started to be illuminated (at x = a in Fig. 2) does not contain any excited molecules yet and therefore there is no emission originating from this point. The point on the other side of the rectangular excitation region (at x = −a) has been illuminated for the time t = 2a/v, where v is the scan velocity, and therefore contains a

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A

B

Figure 2: A: The principle of lifetime imaging with a variable scan speed. The molecules are excited with a rectangular beam with a width 2a (gray line) moving to the right with velocity v. The population of the excited state (solid black line, gray shaded area) depends on the time for which the molecules have been excited: it is lowest at the leading edge of the rectangle (right) and highest at the other side (left). The emission is detected from a region selected by the confocal pinhole, here represented by a rectangle with a width 2b (dashed gray line). The black dotted line shows the population of the excited state obtained with scan velocity 4× lower than that shown with solid black line. B: The dependence of the average detected luminescence on the scan velocity v for different emission rates k1 (eq. 6). Measurements at different scanning velocities v allow the determination of the excited state lifetime τ = 1/k1 . certain population of excited (and emitting) molecules. The excited state population, and therefore the emission, increases from location at x = a to x = −a. The total intensity under the excitation rectangle thus represents the average luminescence for a pulse of duration t = 2a/v. By changing the scan velocity v the average luminescence for pulses of different duration can be measured. The decay rate k1 can then be calculated from the dependence of the mean luminescence intensity F on the scan velocity v (Fig. 2B). After the scanning beam leaves a particular location (x < −a) the luminescence decays with a rate constant k1 . As the size of the detection area (2b) can be controlled by changing the size of the confocal pinhole, luminescence from regions not excited any more may still be detected while scanning (particularly for larger pinhole sizes). Following the approximate geometry in Fig. 2, the luminescence intensity F (v) detected within the rectangle (−b, b) for a given scan velocity v can be divided into two parts: Z a Z −a F (v) = F1 (v) + F2 (v), F1 (v) = y1 (x) dx, F2 (v) = y2 (x) dx (2) −a

−b

where y1 (x) is y(t) from eq. 1 with substitution t = (a − x)/v, and y2 (x) = y(2a/v)e−k1 ((−a−x)/v)

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The solutions are (assuming b ≥ a): F1 (v) = 2ayT and F2 (v) = vyT

k0 k0 + k1

1−

v 1 − e−(k0 +k1 )2a/v 2a k0 + k1

k0 1 − e−(k0 +k1 )2a/v 1 − ek1 (a−b)/v k1 (k0 + k1 )

(4)

(5)

By varying the scan velocity v it is in principle possible to use eq. 2 to determine the decay rate k1 . The particular expressions for F1 and F2 are however based on idealized assumptions of rectangular excitation and detection profiles. We found that for practical implementation it is sufficient to use an approximate formula for F (v) derived from F1 (v): v (6) 1 − e−αk1 /v F (v) = F0 1 − αk1 where α is a calibration factor determined from a measurement of a sample with a known luminescence lifetime τ = 1/k1 , and F0 is the luminescence intensity in the limit of zero scan velocity (continuous excitation). In eq. 6 it is furthermore assumed that the excitation intensity is below the saturation limit (k0 k1 ). Examples of the theoretical dependence of the luminescence intensity F (v) on the scan velocity for several decay rates k1 are shown in Fig. 2B. The decay rate k1 can be determined from the experimental data by pixelwise fitting of the luminescence intensity dependence F (v) on the scan velocity v to eq. 6, with two fitting parameters: F0 and k1 . Alternatively, when only two velocities are used, k1 can be calculated from the ratio F (v2 )/F (v1 ) derived from eq. 6. The necessary requirement for the described lifetime measurement method is that the scan velocity can be varied in a range where the illumination time of a single location is comparable to the luminescence lifetime. The lower lifetime limit for common CLSMs lies in the sub-microsecond to microsecond range, similar to the range of attainable pixel dwell times. These time scales are too long for most fluorescent dyes (nanosecond range), but sufficient for the measurement of lifetimes of transition metal complexes (µs) and lanthanides (µs to ms). The luminescence lifetime of the oxygen-sensitive ruthenium complex Ru(dpp)3 used in this work is on the scale of several µs, and so lies within the lifetime range measurable with a common CLSM.

Results and Discussion Dye in solution To demonstrate the principle of the method we first measured air-equilibrated solution of Ru(dpp)3 in ethylene glycol at concentration 0.2 mM. The sample was placed in a well on a microscope slide and excited with a low intensity. The scan velocity was varied in the range 0.15–1.12 m·s−1 , as described in the Materials and methods section. The luminescence intensity for every scan velocity was determined by averaging the signal in the center part of the corresponding image. As expected, the luminescence decreases with increasing scan velocity. The dependence is described well by eq. 6 (Fig. 3A). The variation of the luminescence with scan velocity depends on the luminescence lifetime τ . In order to produce samples with different luminescence lifetimes we added FeCl3 at concentrations of 4 mM and 10 mM to the Ru(dpp)3 solution. The Fe3+ ions quench the luminescence of Ru(dpp)3 48 , leading to shorter lifetimes. The addition of FeCl3 changes the dependence of luminescence intensity on the scanning velocity, as expected (Fig. 3A). Fitting the data to eq. 6 and assuming the lifetime of unquenched Ru(dpp)3 equal to τ = 2.3 µs 47 , we obtained lifetimes of 1.76 µs and 0.71 µs for the quencher concentrations 4 mM and 10 mM, respectively. As described in the Theory section, the dependence of the luminescence intensity on the scan velocity has its origin in the approach to the stationary state upon start of excitation and depends on the sum of the excitation and emission rates k0 + k1 (eq. 1, Fig. 1B). It is advantageous for practical applications to keep the excitation intensity sufficiently low so that k0 k1 and the

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A

B

Figure 3: A: The experimental dependence of the luminescence of Ru(dpp)3 solutions on the scan velocity for different quencher concentrations, and fits to eq. 6. B. The experimental dependence of the luminescence of Ru(dpp)3 without a quencher on the scan velocity at different relative excitation intensities, together with fits to eq. 6. contribution of the excitation rate to the overall rate can be neglected. We have tested the effect of high excitation rates on the measured F (v) dependence by increasing the excitation intensity by a factor of approx. 3 and 12 (Fig. 3B). While at the highest excitation intensity there is a difference with respect to the lowest excitation intensity, the excitation intensity ∼ 3 higher than the intensity used for all the other experiments did not lead to a significant alteration of the measured F (v) dependence. This result confirms that the lowest excitation intensity used results in sufficiently low excitation rate, so that the approximation k0 k1 is valid. Dye immobilized on a bead To test the method in imaging mode, we prepared porous agarose beads (phenyl sepharose beads) with the oxygen-sensitive dye Ru(dpp)3 immobilized inside the pores of the beads. We imaged two types of samples: in the first type, the beads were placed in air-saturated buffer. The oxygen present in the buffer is expected to partially quench the luminescence of Ru(dpp)3 . In the second sample, the buffer additionally contained the enzyme lactate oxidase and lactate as its substrate. The enzymatic reaction depleted oxygen from the solution, reducing the quenching of the Ru(dpp)3 dye. The beads were imaged using two scan velocities, 0.25 and 0.5 m·s−1 . In accordance with the experiments in solution described above, the luminescence intensity was found to depend on the scan velocity, with lower velocity resulting in higher intensity (Fig. 4). Using eq. 6 and the intensity ratio F (v2 )/F (v1 ), the lifetime images were calculated (Fig. 4D, J). As expected, the mean lifetime of the beads with oxygen depleted from the solution, 3.36 µs, was longer than the

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A

B

C

D

E

F

G

H

J

K

Figure 4: Luminescence lifetime imaging of agarose beads with immobilized Ru(dpp)3 immersed in air-saturated buffer (A-E) and in buffer with enzymatically depleted oxygen (F-K). The luminescence intensity of the beads is lower at higher scan speed (A, B, F, G). C, H: the intensity profiles across the central part of the beads, as marked by two white lines in A. D, J: the luminescence lifetime images. E, K: pixel lifetime distributions of the beads shown in D and J with the mean value marked by a red line. lifetime of the beads in the solution containing oxygen, 2.88 µs (Fig. 4E, K). The distributions of lifetimes (Fig. 4E, K) in individual pixels are relatively broad. This is, however, not surprising considering that the total measurement time per binned pixel in the lifetime images was only about 1 ms. The width of the lifetime distributions is largely determined by the intensity noise in individual pixels. The signal-to-noise ratio could be thus improved by extending the measurement time. Another possibility to increase the precision of lifetime determination is to optimize the choice of scan velocities. Without performing a detailed analysis we note that a good combination of two velocities would be the following: a very slow velocity v1 : v1 /(αk1 ) 1, providing a ‘reference’ intensity value which is minimally influenced by the transient effects, and the second velocity v2 being matched to the luminescence time scale: v2 /(αk1 ) ≈ 1 (v = 1 in Fig. 2B), and thus ideally sensing the effect of finite luminescence lifetime on the detected fluorescence intensity. Here we were limited by the microscope design to two scan velocities differing only by a factor of two. This is however not a principal limitation, as newer confocal laser scanning microscopes typically allow a much broader range of scanning speeds, largely independent of the magnification. In a second set of experiments we immobilized the enzyme together with the Ru(dpp)3 dye inside the porous agarose beads. The beads were packed in a microfluidic channel together with a larger fraction of unlabeled beads without enzyme. The unlabeled beads made it possible to achieve dense packing while keeping the density of labeled beads relatively low. First, the microchannel was filled with only buffer without the enzyme substrate. Imaging the beads under flow at two scan velocities and calculating the lifetime image, as described above, yielded a lifetime influenced by oxygen quenching (Fig. 5A, B). Afterwards, lactate was added to the buffer. The lifetime images showed increase in the average lifetime of Ru(dpp)3 (Fig. 5C, D) indicating the reduction of oxygen quenching as a result of oxygen consumption by the reaction of the immobilized enzyme with the substrate added to the buffer. A limited motion of the beads due to the liquid flow, despite the packing of the beads, could not be avoided between the acquisition of the two images at different scan speeds. When small and when limited to the lateral plane, the shift could be partially corrected for by manual adjustment during the analysis. These experiments were conducted in optically rather unfavourable setting, as the focused light had to partially pass through beads located in layers above the imaged beads, thus distorting the laser focus and so affecting the detected luminescence intensity. The results

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A

B

C

D

Figure 5: Luminescence lifetime images of beads with immobilized enzyme and Ru(dpp)3 , imaged under flow without (A, B) and with (C, D) substrate, resulting in different oxygen concentrations. B, D: pixel lifetime distributions of the beads shown in A and C with the mean value marked by a red line. nevertheless show that even under these difficult but realistic conditions differences in luminescence lifetimes can be clearly detected and visualized.

Conclusions and Implications In this work we introduce a method for the measurement of luminescence lifetimes based on monitoring the relaxation of the dye molecules to a new steady state upon the onset of excitation. This contrasts the more conventional time-domain methods for the determination of excited state lifetimes, where relaxation to the ground state is measured after the excitation is finished. In the latter methods, the excitation period is followed by the detection period; ideally one strives to avoid any temporal overlap between the excitation and the emission. In contrast to this, in the method presented here the excitation and the detection take place simultaneously. As this is the normal mode of operation of a CLSM, the method can be readily implemented in a scanning microscope. The principle of operation is not limited to a common point-scanning CLSM, but can be applied also in much faster line-scanning microscopes 49,50 . The major advantage of the technique is that there is no additional need for pulsed or timemodulated excitation light sources or fast time-gated detectors. The temporal resolution has its origin in the time for which the luminescence is measured: short times mean that the stationary state is not reached and the average luminescence is lower than for longer measurement times. The control of the measurement time is conveniently realized by changing the scan velocity in a CLSM. We have shown that two scan velocities, that is, two images, are sufficient to perform lifetime imaging. Since the detection of the emission takes place during the excitation, any fluorescence from a short-lived probe can be also detected. Consequently, the time-domain DLR 37 principle of detecting the intensity changes of a short-lifetime fluorescent sensing probe by measuring the effective changes of a long lifetime of an insensitive phosphorescent probe could possibly be exploited. The presented method is technically similar to the imaging of transient states by CLSM 6,51 . There, the relaxation to a steady state involving three energy states: the ground and the excited singlet states and the triplet (or other transient) state, was probed. The focus was on monitoring the population of the non-emissive triplet state, not its lifetime. In contrast, here we consider only two energy states: the ground and the excited state; and the goal is the determination of the lifetime of the emitting state. Similarly to the time-gating method, where the application of more than two time gates allows the analysis of multiexponential decays 52 , it should be possible to extend the presented method to

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two-exponential decays. This would require measurements at more than two scanning speeds, with more speeds increasing the precision. An attractive application of two-exponential decay analysis are time-resolved emission anisotropy experiments 53 , whereby the long luminescence lifetimes of phosphorescent dyes would allow the quantification of rotational motion, and therefore the size, of molecules with a large molecular weight, such as proteins or protein complexes. Although the luminescence lifetime measurement has been realized here as lifetime imaging in a CLSM, the imaging mode is not a necessary requirement. The only important aspect of the described method is the motion of the excitation focus, and the confocal detection from the same volume. The same principle could be realized, for example, by scanning the beam in a circle with a diameter large enough so that the excited state fully decays before the beam returns to the same position. The scanning could be realized mechanically or by electrooptical scanners. The device could be possibly incorporated into a small probe or endoscope with the scanned circle being either outside of the probe, within the liquid sample as in a CLSM, or within a thin permeable layer on the probe surface containing the immobilized luminescent dye, as in commonly used optical probes 54,55 . Another potential implementation is to use fixed excitation and detection volume and let the sample flow through the volume with different velocities. This configuration would allow integration in a microfluidic channel, with the flow velocity controlled by external pump or by having a channel with variable width and multiple measurement spots at positions of different channel width, therefore different flow velocity. In this way it might be possible to realize lifetime measurements of luminophores with lifetimes in the millisecond range, such as Eu complexes, and perhaps even lifetimes in the range of tens of µs. It is the maximum achievable flow velocity that eventually determines the lower limit of measurable lifetimes.

References [1] Siraj, N.; El-Zahab, B.; Hamdan, S.; Karam, T. E.; Haber, L. H.; Li, M.; Fakayode, S. O.; Das, S.; Valle, B.; Strongin, R. M.; Patonay, G.; Sintim, H. O.; Baker, G. A.; Powe, A.; Lowry, M.; Karolin, J. O.; Geddes, C. D.; Warner, I. M. Anal. Chem. 2016, 88, 170–202, PMID: 26575092. [2] Sch¨ aferling, M. Angew. Chem. Int. Ed. 2012, 51, 3532–3554. [3] Lakowicz, J. R. Principles of Fluorescence Spectroscopy, 3rd ed.; Springer, 2006; Chapter 20, pp 679–681. [4] Suhling, K.; French, P. M. W.; Phillips, D. Photochem. Photobiol. Sci. 2005, 4, 13–22. [5] Becker, W. J. Microsc. 2012, 247, 119–136. [6] Sandén, T.; Persson, G.; Widengren, J. Anal. Chem. 2008, 80, 9589–9596, PMID: 19007245. [7] Wang, X.-d.; Wolfbeis, O. S. Chem. Soc. Rev. 2014, 43, 3666–3761. [8] Quaranta, M.; Borisov, S. M.; Klimant, I. Bioanalytical Reviews 2012, 4, 115–157. [9] Papkovsky, D. B.; Dmitriev, R. I. Chem. Soc. Rev. 2013, 42, 8700–8732. [10] Wolfbeis, O. S.; Klimant, I.; Werner, T.; Huber, C.; Kosch, U.; Krause, C.; Neurauter, G.; Drkop, A. Sens. Actuators, B 1998, 51, 17–24. [11] Dmitriev, R. I.; Papkovsky, D. B. Cell. Mol. Life Sci. 2012, 69, 2025–2039. [12] Finikova, O. S.; Lebedev, A. Y.; Aprelev, A.; Troxler, T.; Gao, F.; Garnacho, C.; Muro, S.; Hochstrasser, R. M.; Vinogradov, S. A. ChemPhysChem 2008, 9, 1673–1679. [13] Lo, K. K.-W.; Li, S. P.-Y.; Zhang, K. Y. New J. Chem. 2011, 35, 265–287. [14] Berezin, M. Y.; Achilefu, S. Chem. Rev. 2010, 110, 2641–2684, PMID: 20356094.

12


Page 12 of 15

Page 13 of 15

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


[15] Lakowicz, J. R. Principles of Fluorescence Spectroscopy, 3rd ed.; Springer, 2006; Chapter 20, pp 679–681. [16] Sakadˇzić, S.; Mandeville, E. T.; Gagnon, L.; Musacchia, J. J.; Yaseen, M. A.; Yucel, M. A.; Lefebvre, J.; Lesage, F.; Dale, A. M.; Eikermann-Haerter, K.; Ayata, C.; Srinivasan, V. J.; Lo, E. H.; Devor, A.; Boas, D. A. Nat. Commun. 2014, 5, 5734. [17] Wanek, J.; Blair, N. P.; Shahidi, M. Curr. Eye Res. 2012, 37, 132–137. [18] Schreml, S.; Meier, R. J.; Wolfbeis, O. S.; Maisch, T.; Szeimies, R. M.; Landthaler, M.; Regensburger, J.; Santarelli, F.; Klimant, I.; Babilas, P. Exp. Dermatol. 2011, 20, 550–554. [19] Li, Z.; Roussakis, E.; Koolen, P. G. L.; Ibrahim, A. M. S.; Kim, K.; Rose, L. F.; Wu, J.; Nichols, A. J.; Baek, Y.; Birngruber, R.; Apiou-Sbirlea, G.; Matyal, R.; Huang, T.; Chan, R.; Lin, S. J.; Evans, C. L. Biomed. Opt. Express 2014, 5, 3748–3764. [20] Vinogradov, S. A.; Lo, L. W.; Jenkins, W. T.; Evans, S. M.; Koch, C.; Wilson, D. F. Biophys. J. 1996, 70, 1609–1617. [21] Bolivar, J. M.; Eisl, I.; Nidetzky, B. Catal. Today 2016, 259, 66–80. [22] Bolivar, J. M.; Consolati, T.; Mayr, T.; Nidetzky, B. Trends Biotechnol. 2013, 31, 194–203. [23] Aldén, M.; Omrane, A.; Richter, M.; Särner, G. Prog. Energy Combust. Sci. 2011, 37, 422– 461. [24] Allison, S. W.; Gillies, G. T. Rev. Sci. Instrum. 1997, 68, 2615–2650. [25] Shang, L.; Stockmar, F.; Azadfar, N.; Nienhaus, G. U. Angew. Chem. Int. Ed. 2013, 52, 11154–11157. [26] Giesecke, J. A.; Schubert, M. C.; Schindler, F.; Warta, W. IEEE Journal of Photovoltaics 2015, 5, 313–319. [27] Wilson, M.; Lagowski, J.; Edelman, P.; Korsos, F.; Nadudvari, G.; Kiss, Z.; Schmauder, J.; Mihailetchi, V.; Olibet, S. Energy Procedia 2013, 38, 209 – 215. [28] Lu, Y.; Zhao, J.; Zhang, R.; Liu, Y.; Liu, D.; Goldys, E. M.; Yang, X.; Xi, P.; Sunna, A.; Lu, J.; Shi, Y.; Leif, R. C.; Huo, Y.; Shen, J.; Piper, J. A.; Robinson, J. P.; Jin, D. Nat. Photon. 2014, 8, 32–36. [29] Becker, W.; Su, B.; Bergmann, A.; Weisshart, K.; Holub, O. Proc. SPIE 2011, 7903, 790320. [30] Jahn, K.; Buschmann, V.; Hille, C. Sci. Rep. 2015, 5, 14334. [31] Shcheslavskiy, V.; Neubauer, A.; Bukowiecki, R.; Dinter, F.; Becker, W. Appl. Phys. Lett. 2016, 108, 091111. [32] Woods, R. J.; Scypinski, S.; Love, L. J. C.; Ashworth, H. A. Anal. Chem. 1984, 56, 1395–1400. [33] Ballew, R. M.; Demas, J. N. Anal. Chem. 1989, 61, 30–33. [34] Liebsch, G.; Klimant, I.; Frank, B.; Holst, G.; Wolfbeis, O. S. Appl. Spectrosc. 2000, 54, 548–559. [35] Lakowicz, J. R.; Castellano, F. N.; Dattelbaum, J. D.; Tolosa, L.; Rao, G.; Gryczynski, I. Anal. Chem. 1998, 70, 5115–5121. [36] Huber, C.; Klimant, I.; Krause, C.; Wolfbeis, O. S. Anal. Chem. 2001, 73, 2097–2103. [37] Liebsch, G.; Klimant, I.; Krause, C.; Wolfbeis, O. S. Anal. Chem. 2001, 73, 4354–4363. [38] Schroeder, C. R.; Neurauter, G.; Klimant, I. Microchim. Acta 2007, 158, 205–218.

13



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

[39] Schr¨ oder, C. R.; Polerecky, L.; Klimant, I. Anal. Chem. 2007, 79, 60–70. [40] Kondrashina, A. V.; Dmitriev, R. I.; Borisov, S. M.; Klimant, I.; O’Brien, I.; Nolan, Y. M.; Zhdanov, A. V.; Papkovsky, D. B. Adv. Funct. Mater. 2012, 22, 4931–4939. [41] Choi, H.; Tzeranis, D. S.; Cha, J. W.; Clémenceau, P.; de Jong, S. J.; van Geest, L. K.; Moon, J. H.; Yannas, I. V.; So, P. T. Opt. Express 2012, 20, 26219–26235. [42] Grichine, A.; Haefele, A.; Pascal, S.; Duperray, A.; Michel, R.; Andraud, C.; Maury, O. Chem. Sci. 2014, 5, 3475–3485. [43] Ramshesh, V. K.; Lemasters, J. J. J. Biomed. Opt. 2008, 13, 064001. [44] Boniello, C.; Mayr, T.; Klimant, I.; Koenig, B.; Riethorst, W.; Nidetzky, B. Biotechnol. Bioeng. 2010, 106, 528–540. [45] Boniello, C.; Mayr, T.; Bolivar, J. M.; Nidetzky, B. BMC Biotechnol. 2012, 12, 11. [46] Bolivar, J. M.; Consolati, T.; Mayr, T.; Nidetzky, B. Biotechnol. Bioeng. 2013, 110, 2086– 2095. [47] Morris, K. J.; Roach, M. S.; Xu, W.; Demas, J. N.; DeGraff, B. A. Anal. Chem. 2007, 79, 9310–9314, PMID: 17985845. [48] Hoffman, M. Z.; Bolletta, F.; Moggi, L.; Hug, G. L. J. Phys. Chem. Ref. Data 1989, 18, 219–543. [49] Scully, A. D.; Ostler, R. B.; MacRobert, A. J.; Parker, A. W.; de Lara, C.; O’Neill, P.; Phillips, D. Photochem. Photobiol. 1998, 68, 199–204. [50] Im, K.-B.; Han, S.; Park, H.; Kim, D.; Kim, B.-M. Opt. Express 2005, 13, 5151–5156. [51] Widengren, J. J. R. Soc. Interface 2010, 7, 1135–1144. [52] Sharman, K. K.; Periasamy, A.; Ashworth, H.; Demas, J. N.; Snow, N. H. Anal. Chem. 1999, 71, 947–952. [53] Hirvonen, L. M.; Fruhwirth, G. O.; Srikantha, N.; Barber, M. J.; Neffendorf, J. E.; Suhling, K.; Jackson, T. L. Pharm. Res. 2016, 33, 2025–2032. [54] Orellana, G.; Haigh, D. Curr. Anal. Chem. 2008, 4, 273–295. [55] Wang, X.-d.; Wolfbeis, O. S. Anal. Chem. 2016, 88, 203–227.

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Confocal Luminescence Lifetime Imaging with Variable Scan Velocity

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