Photosynth Res DOI 10.1007/s11120-015-0104-2
EMERGING TECHNIQUES
Characterizing non-photochemical quenching in leaves through fluorescence lifetime snapshots Emily J. Sylak-Glassman1,2,3 • Julia Zaks2,4,5 • Kapil Amarnath1,2,6 Michelle Leuenberger1,2 • Graham R. Fleming1,2,4
•
Received: 19 November 2014 / Accepted: 16 February 2015 Ó Springer Science+Business Media Dordrecht 2015
Abstract We describe a technique to measure the fluorescence decay profiles of intact leaves during adaptation to high light and subsequent relaxation to dark conditions. We show how to ensure that photosystem II reaction centers are closed and compare data for wild type Arabidopsis thaliana with conventional pulse-amplitude modulated (PAM) fluorescence measurements. Unlike PAM measurements, the lifetime measurements are not sensitive to photobleaching or chloroplast shielding, and the form of the fluorescence decay provides additional information to test quantitative models of excitation dynamics in intact leaves.
Electronic supplementary material The online version of this article (doi:10.1007/s11120-015-0104-2) contains supplementary material, which is available to authorized users. & Graham R. Fleming [email protected] 1
Department of Chemistry, University of California Berkeley, Berkeley, CA 94720, USA
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Physical Biosciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, USA
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Present Address: IDA Science and Technology Policy Institute, 1899 Pennsylvania Avenue Suite 520, Washington, DC 20006, USA
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Graduate Program in Applied Science and Technology, University of California Berkeley, Berkeley, CA 94720, USA
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Present Address: Agilent Technologies, 5301 Stevens Creek Blvd, Santa Clara, CA 95051, USA
6
Present Address: FAS Center for Systems Biology, Harvard University, 52 Oxford Street, Rm. 440, Cambridge, MA 02138, USA
Keywords Arabidopsis thaliana Chlorophyll Fluorescence lifetime Fluorescence yield Nonphotochemical quenching
Introduction Plants dynamically regulate the photosynthetic apparatus in response to fluctuations in light intensity (Blankenship 2002). To minimize damage (Barber 1995; Melis 1999) from excess light plants utilize a set of mechanisms collectively known as non-photochemical quenching (NPQ) to dissipate singlet excitations safely as heat (Demmig-Adams and Adams 1996). The most rapid component of NPQ is known as energy-dependent quenching (qE). An improved understanding of NPQ in general and qE in particular is of importance as mutant plants impaired in NPQ have decreased fitness (Ku¨lheim et al. 2002) and because altering the dynamics of NPQ may improve photosynthetic efficiency (Zhu et al. 2010). The amount of NPQ is generally measured through changes in the chlorophyll fluorescence yield, most often using a pulse-amplitude modulated (PAM) fluorometer. In a typical experiment, a dark-adapted leaf is monitored while it is exposed first to excess light to turn on NPQ and then to darkness as NPQ relaxes (Cazzaniga et al. 2013). While changes in the chlorophyll fluorescence yield are interpreted as the result of changes in the amount of NPQ, these fluorescence yield changes can have multiple causes. For instance, photobleaching and chloroplast movement can also decrease the fluorescence yield, thus complicating the interpretation of PAM data (Baker 2008). In contrast, the time-resolved fluorescence decay of chlorophyll is not sensitive to either photobleaching or chloroplast movement. The form of the fluorescence decay is sensitive to the
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number of quenchers and their qE rates and holds the potential for greater insight into the entire NPQ phenomenon. Because the amount of qE turns on and off in seconds to minutes, the fluorescence decays must be collected in time increments (‘‘snapshots’’) commensurate with the intrinsic timescales on which qE turns on and off. Amarnath et al. solved this problem by building a set of computer-controlled shutters to measure the fluorescence decay of Chlamydomonas reinhardtii for 80 ms every 200 ms, ensuring that the decay profile was effectively constant during the measurement (Amarnath et al. 2012). Repeated measurements were used to produce adequate statistics. In this paper, we describe the extension of this fluorescence lifetime snapshot approach to detached leaves of Arabidopsis thaliana (A. thaliana) plants.
Methods Measurements were made on medium-sized A. thaliana leaves detached from plants before the stage of bolting. Plants were grown in a growth chamber with 100 lmol photons 9 m-2 s-1 light. Measurements were carried out using a similar procedure to that described by Sylak-Glassman et al. (2014). Plants were each dark-adapted for 30 min prior to detaching a leaf. To prevent any plant from having its light cycle disturbed, measurements were performed on multiple plants during each day of data collection so that any given plant would not be dark-acclimated for more than 30 min in a 1.5 h period. Between dark adaptation times, plants were placed in a 100 lmol photons 9 m-2 s-1 light chamber for re-light acclimation. Each leaf was selected from the middle of the rosette to control for leaf age and size. After leaf selection, the leaf was immediately placed in a home-built holder (shown in Online Resource 1). To keep the leaf from overheating or dehydrating, the holder keeps most of the leaf open to air, and allows the leaf to take up water. Similar to the set-up described by Amarnath et al., the light exposure of the leaf and detector were controlled by shutters programmed in LabVIEW. The set-up is shown in Fig. 1a. The excitation pulse hits the leaf at 70° to the adaxial leaf surface (Fig. 1b). Excitation is into the chlorophyll a (chl a) Soret band at 420 nm. The average power of the laser at the sample was 5 mW with a pulse energy of 66 pJ. This laser power corresponds to an approximate intensity of 5000 lmol photons 9 m-2 s-1, which is enough to close reaction centers (Schansker et al. 2006) and is similar to the intensity used in PAM fluorometers (Brooks and Niyogi 2011; Cazzaniga et al. 2013; Porcar-Castell et al. 2008). An actinic light (Schott KL1500) was used to turn on qE. The leaf’s exposure to the actinic light was controlled
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by a shutter. The actinic light intensity was set at either 500 lmol photons 9 m-2 s-1 or 1200 lmol photons 9 m-2 s-1 for experiments. The light was collimated by a lens, forming a 1 cm diameter illumination spot size at a 45° angle to the adaxial side of the leaf. Detection was at a 45° angle to the adaxial side of the leaf and at a 65° angle to the laser. Fluorescence was collimated with a 5 cm lens and passed through a polarizer set to the magic angle (54.7°). The light was then focused into a monochromator (HORIBA Jobin–Yvon; H-20) with a 10 cm lens. The detection wavelength was centered at 684, 715, or 720 nm. 684 nm was chosen to look at fluorescence emanating from the Qy band of chlorophyll a, and 715 and 720 nm were chosen to mimic the spectrum of light collected by PAM fluorometers. The light from the monochromator was collimated with a 5 cm lens before being sent to the detector (Hamamatsu; R3809U). The instrument response function was 35–39 ps FWHM. Following Amarnath et al., the sequence of shutters opening and closing was designed in MATLAB and executed in LabVIEW (Amarnath et al. 2012). A typical measurement sequence consisted of an initial measurement taken on a dark-adapted leaf. After 2 min, the leaves were illuminated with actinic light. After 10 min of illumination, the actinic light was blocked, and the leaf was allowed to relax for 10 min. Data were collected at intervals ranging from 10 to 60 s during the light acclimation period and subsequent relaxation period in dark. Unless otherwise specified, identical data collection routines were performed on multiple leaves from multiple plants (6 leaves total from 3 different plants). The process for data summing, aligning, and fitting is identical to that described by Sylak-Glassman et al. (2014). The data from each leaf were aligned according to the maximum of a cross-correlation [xcorAlign.m (Carmichael 2010)]. The curves were fit to a sum of four exponentials: three decay functions and one fast rise (1–4 ps) (Picoquant Fluofit Pro 4.5). A comparison of the fits resulting from fitting the data to three decay functions versus three decay functions and a fast rise are shown in Online Resource 2. Following data fitting, curves were reconstructed by plotting the fluorescence intensity calculated by the exponential decays: P Ai et=si FðtÞ ¼ i P ; ð1Þ i Ai where Ai are the amplitudes and si are the fluorescence lifetime components. Amplitude-weighted average lifetimes of the leaves, savg, were calculated using the three exponential decays as follows: P i Ai s i : ð2Þ savg ¼ P i Ai
Photosynth Res Fig. 1 A top-view schematic of the apparatus is shown in the top (a), and the detection geometry is shown below (b). A diode laser pumps an ultrafast Ti:Sapphire oscillator. The resulting 840 nm light is frequency doubled to 420 nm. A portion of the beam is sent through a shutter to the leaf in its holder (green line). An actinic light is also sent through a shutter such that both beams can impinge on the leaf. Fluorescence is collected at the magic angle. The illumination and detection geometry is shown in more detail in b. The leaf is held at a 45° angle to the actinic light and the detector. The angle between the laser and actinic light is 25°
The standard deviations of the values of savg were calculated by performing bootstrapping on the lifetimes. Bootstrapping involves selecting a subset of points in the fluorescence decay and determining the fluorescence lifetime components required to fit the subset of points. Repeating this process one thousand times gives an estimate of the error in the fluorescence lifetime fitting parameters. The amplitudes were bootstrapped separately to determine the error in the amplitudes of each component.
Results We confirmed that detached leaves exhibit the same qE characteristics as leaves attached to an intact plant provided that the detached leaf is able to take up water during the measurement (Fig. 2a and Fig. 2b). The amount of qE is significantly reduced for detached leaves without access to water (Fig. 2c) In order to selectively track changes in the fluorescence decay profile due to NPQ rather than photochemistry, it is necessary to measure the fluorescence when the reaction centers are closed (Stirbet 2011). This corresponds to the longest fluorescence lifetime, and we determined via multiple fluorescence decay measurements on leaves while they are exposed to the laser for different lengths of time. In data collection, the total exposure time was 1 s total divided into five 0.2 s bins. This is shown schematically in
Fig. 3a. Figure 3b shows how the fluorescence lifetime changes over seven overlapping 0.2 s bins for leaves measured at 715 nm for wild type A. thaliana. Five leaves were summed for each 0.2 s bin, and the average fluorescence lifetime reached a peak at 0.4 s of laser exposure before the lifetime measurement. Once the bin with the longest lifetime is identified, it is used for further data analysis, and the other bins are discarded. The high concentration of chloroplasts in leaves leads to re-absorption and subsequent re-emission of fluorescence. This has a large impact on the leaf’s emission spectrum, which shows a peak at 735 nm that is absent in measurements on isolated chloroplasts (Moise and Moya 2004). It has been hypothesized that the re-emitted fluorescence prevents the accurate measurement of fluorescence lifetimes (Johnson and Ruban 2009). While relative changes in the fluorescence lifetime during qE would still be instructive, for the measurements to be used to parameterize models, the fluorescence lifetimes must be accurate. To verify the role of chlorophyll concentration on the measured fluorescence lifetime, the fluorescence lifetimes were compared between the spinach leaves and solutions of spinach chloroplasts with varying concentrations at two different wavelengths, 684 and 720 nm. These wavelengths were chosen to correspond to fluorescence from the Qy band of chlorophyll (684 nm) and to a wavelength range closer to that measured in a PAM fluorometer (720 nm). It is expected that the average fluorescence lifetimes at 684
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Fig. 2 The fluorescence decays for 3 leaves with different sample holder conditions and measured after 3 different illumination conditions are shown in (a, b, and c). The fluorescence decays for dark-adapted leaves are shown in a. The fluorescence lifetimes for the same leaves after 10 min of illumination are shown in b. The same leaves after 60 min of relaxation in the dark are shown in c. In each of
Fig. 3 Multiple fluorescence measurements are required to determine the maximum average lifetime. A schematic for how a single measurement period is subdivided is shown in a. The measurement period is 1 s long, during which time the leaf is exposed to the laser and fluorescence is detected. The measurement period is subdivided into 5 steps, each with a 0.2 s duration. The time increment with the maximum average lifetime (shown in gray, with the label peak savg), is the only step that is retained. The duration of laser exposure prior to the step with the maximum average fluorescence lifetime is referred to as the saturation time, because it is the time required to close all the PSII reaction centers. Measurements of savg summed for 5 darkadapted leaves are shown in b, measured at 715 nm. Exposure to the laser begins at 0 s. The horizontal bars indicate the time span over which the measurement was made. The vertical bars indicate the standard deviation in the average fluorescence lifetime value. The saturation time for this measurement is 0.3 s
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the images, the fluorescence decays for the leaf attached to a plant are shown in black, the fluorescence decays for the leaf detached from the plant with the stem immersed in water are shown in gray, and the fluorescence decays for the leaf detached from the plant without water are shown in red
and 720 nm will differ because of differing fractions of fluorescence from PSII and PSI antennas (Borisov and Il’Ina 1973; Moise and Moya 2004), each of which exhibits different chlorophyll energy transfer dynamics. The average fluorescence lifetimes are shown in Table 1. The spinach leaf and chloroplast solutions with a concentration of chlorophyll of 0.025 and 0.125 lg/lL all had nearly equivalent fluorescence lifetimes at 684 nm. At 720 nm, the spinach leaf has a fluorescence lifetime most similar to the lowest concentration of chloroplasts, 0.0025 lg/lL. Given that a typical spinach leaf has a chlorophyll concentration of approximately 2.7 lg/lL (Mehler 1951; Porra et al. 1989) and an average fluorescence lifetime similar to that of the chloroplast solutions, the average fluorescence lifetime does not show a concentration dependence over 3–4 orders of magnitude of chlorophyll concentration. The very dilute sample of chloroplasts (0.0025 lg/lL) displayed a slightly shorter average fluorescence lifetime. The consistency of the fluorescence lifetimes between the isolated chloroplasts and the leaves indicates that the fluorescence lifetimes that we measure for leaves are uncontaminated by non-primary fluorescence. To examine the effect of the actinic light intensity on the measured fluorescence lifetime, two actinic light intensities Table 1 Average fluorescence lifetimes of a spinach leaf and of spinach chloroplasts, measured at 684 and 720 nm [Chl] (lg/lL)
savg (ns) at 684 nm
savg (ns) at 720 nm
0.0025
1.20 ± 0.06
0.47 ± 0.03
0.025
1.37 ± 0.07
0.64 ± 0.03
0.125
1.44 ± 0.08
0.64 ± 0.03
Leaf
1.44 ± 0.07
0.51 ± 0.02
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were selected that are typically used in NPQ measurements: 500 and 1200 lmol photons 9 m-2 s-1. Lower actinic light intensities are not high enough to sustain NPQ (Kalituho et al. 2007), whereas higher light intensities can cause significant photodamage (van Wijk et al. 1993). When the fluorescence lifetimes are measured throughout the induction and relaxation of NPQ, it is possible to see the fluorescence lifetimes as a function of time during or after exposure to actinic light. The series of measurements can therefore be visualized in a two-dimensional plot of the fluorescence intensity as a function of time following excitation and as a function of the illumination time. Plots for leaves illuminated with 500 and 1200 lmol photons 9 m-2 s-1 light are shown in Fig. 4a and b, respectively. The normalized fluorescence intensity is calculated from the exponential decays used to fit each fluorescence lifetime measurement, and is displayed on a log scale. The horizontal axis is the timescale of the fluorescence decay in nanoseconds. The vertical axis is the timescale of light acclimation in minutes. The actinic light is on for 10 min, as shown by the white bar on the left in Fig. 4. The different dynamics for the induction of NPQ under two different illumination intensities can be seen by comparing the fluorescence intensity in the 2D plots. The leaves illuminated with 500 lmol photons 9 m-2 s-1 light have an initial shortening of the fluorescence lifetime during the first minute of illumination. Then, over the next 2 min, the decay lengthens. In minutes 3–5, the fluorescence lifetime shortens again, before stabilizing during the last 4 min of illumination. The dynamics of the induction of qE are different for the leaves illuminated with 1200 lmol photons 9 m-2 s-1 light. Similar to the 500 lmol photons 9 m-2 s-1 illumination leaves there is an initial
shortening of the fluorescence lifetime during the first minute of light acclimation. However, after this time, the fluorescence intensity gradually increases at long decay times (C4 ns) while it decreases at short decay times (B3 ns). The dynamics of the relaxation of NPQ for the leaves illuminated with 500 and 1200 lmol photons 9 m-2 s-1 light are very similar. Both experience an increasingly gradual lengthening of the fluorescence decays. Integrating over the fluorescence decay axis results in the amplitude-weighted average fluorescence lifetime, shown in Fig. 5. Because the average fluorescence lifetime is proportional to the fluorescence yield, the dynamics of the average lifetime should resemble the dynamics of the fluorescence yield under conditions of closed PSII reaction centers measured with PAM fluorometers. However, as noted earlier, unlike chlorophyll fluorescence yield measurements, the amplitude-weighted average fluorescence lifetime is not influenced by non-quenching processes such as chloroplast shielding and photobleaching. To assess the extent to which non-quenching processes influence the amount of NPQ measured by PAM fluorometers, we compared the NPQ dynamics from the fluorescence lifetime measurements with the NPQ dynamics as measured through PAM fluorometry. Because the actinic light color has been shown to influence chloroplast shielding, and in turn, the chlorophyll fluorescence yield, two different PAM fluorometers were used, each with a different color of actinic light. The Hansatech FMS fluorometer uses a white actinic light (Mouget and Tremblin 2002), while the Walz Dual PAM fluorometer uses a red actinic light (Heinz Walz GmbH 2006). Both PAM fluorometers detect at similar wavelengths, [705 nm for the white light PAM, and [700 nm for the red light PAM. The two different PAM
Fig. 4 Multiple measurements of the chlorophyll fluorescence lifetime for leaves illuminated with a 500 lmol photons 9 m-2 s-1 and b 1200 lmol photons 9 m-2 s-1 are represented in two-dimensional plots. The maximum fluorescence for each measurement is normalized to 1 and intensity is shown on a logarithmic scale. The average
decay time in nanoseconds is shown on the horizontal axis. The vertical axis is the time in minutes for the leaves’ acclimation to light. The actinic light turns on at 0 min, and is on for 10 min, at which point it is turned off for the remaining 10 min. The bars at the left of a indicate when the actinic light is on (white) and off (black)
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quantum yields of each component of NPQ when reaction center are closed (Ahn et al. 2009). Because this term is linearly related to the fluorescence yield, it more accurately tracks the changing excitation dissipation dynamics as NPQ turns on and off. The NPQ parameter from the PAM fluorescence measurements was calculated by comparing the fluorescence yield in the dark when PSII reaction centers are closed, Fm and the fluorescence yield after light exposure when PSII 0 reaction centers are closed, Fm : 0
Fm Fm NPQ ¼ 0 Fm
Fig. 5 The average fluorescence lifetime for leaves illuminated with 500 lmol photons 9 m-2 s-1 (gray) and 1200 lmol photons 9 m-2 s-1 (black). The bars at the top of the figure indicate when the actinic light is on (white) and off (black)
ð3Þ
The NPQ curves from the fluorescence lifetime measurements were calculated using an analogous equation [see Supplement by Sylak-Glassman et al. (2014) for derivation]: savg;dark savg;light NPQs ¼ ð4Þ savg;light
fluorometers were both set to an actinic light intensity of 1000 lmol photons 9 m-2 s-1. These illumination and detection parameters differ slightly from the fluorescence lifetime apparatus, which detects fluorescence centered at 684 nm and uses a white actinic light with an intensity of 1200 lmol photons 9 m-2 s-1, though this difference in intensity is not expected to have a significant impact on the amount of qE (Bailey et al. 2004). In addition to calculating the standard NPQ parameter, we also calculated the approximate quantum yield of NPQ with PSII reaction centers closed, UNPQ,RCC. This term was chosen because it also reports the amount of qE but, unlike the NPQ parameter, is linearly related to the fluorescence yield of the light-adapted leaf and is equal to the sum of the
A comparison of NPQ versus time for the fluorescence lifetime data and the two PAM fluorometers is shown in Fig. 6a. For the values of UNPQ,RCC from the PAM fluorescence measurements, the following equation was used:
Fig. 6 a NPQ and b UNPQ,RCC from fluorescence lifetime measurements (black), a red (620 nm) actinic light PAM (Walz Dual PAM) (red), and a white actinic light PAM (Hansatech FMS) (green). The fluorescence lifetime measurements have a white actinic light with an intensity of 1200 lmol photons 9 m-2 s-1 and the PAM measurements have an
actinic light intensity of 1000 lmol photons 9 m-2 s-1. The data points are shown as circles. The smoothed data are shown as lines. The bars at the top of the figures indicate when the actinic light is on (white) and off (black). Error bars corresponding to one standard deviation are shown for the NPQ PAM measurements
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/NPQ;RCC
Fm Fm ¼ Fm
ð5Þ
The calculation of UNPQ,RCC for the fluorescence lifetime measurements was adapted from the fluorescence yield measurements in the same manner as the NPQ parameter: savg;dark savg;light /NPQ;RCC;s ¼ ð6Þ savg;dark
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A plot of UNPQ,RCC,s versus time for the fluorescence lifetime data and the two PAM measurements is shown in Fig. 6b. The NPQ dynamics shown in Fig. 6a during the induction of qE are similar for the fluorescence lifetime measurement and the red actinic light PAM measurement. The two reach similar maximum values of NPQ as well. The slightly higher amount of NPQ from the fluorescence lifetime measurements compared to the white actinic light PAM measurement may be due to the difference in detection wavelength (Lambrev et al. 2010). The NPQ curve from the white actinic light PAM measurement displays an additional rise component and reaches a maximum value greater than 3, whereas the other two techniques reach a maximum value just under 2. However, the relaxation of NPQ is nearly identical for both the fluorescence lifetime measurement and the white actinic light PAM measurement, whereas the relaxation kinetics from the red actinic light PAM measurements are faster. The similarity between the amount of qE measured by red actinic light PAM fluorometers and the fluorescence lifetime measurements suggests that the chlorophyll fluorescence yield NPQ parameter from red actinic light PAM fluorometers is uncontaminated by nonquenching processes such as photobleaching or chloroplast avoidance. As suggested by Cazzaniga and co-workers, the extra NPQ seen in the white actinic light PAM measurements does not appear to be due to qE (Cazzaniga et al. 2013). While both the white actinic light PAM and the fluorescence lifetime measurements show a slower relaxation of qE than the red actinic light PAM, they most likely have different origins. The slower relaxation in white light PAM measurements has been shown to be due to a reversal of chloroplast shielding (Cazzaniga et al. 2013). The slower relaxation in fluorescence lifetime measurements must be due to a relaxation of qE processes, and could be due to a different response to the 420 nm laser light used in measurements compared to the saturating pulses of red or white light used in PAM fluorometers. The NPQ-related dynamics and values of all three techniques are much more similar when UNPQ,RCC is plotted instead of NPQ (Fig. 6b). The induction dynamics seen in the UNPQ,RCC curves for the fluorescence lifetime measurement and the red actinic light PAM measurement are still very similar, but the difference between these two techniques and the white actinic light PAM measurement is less drastic. Similar to the NPQ relaxation kinetics, the relaxation of UNPQ,RCC is nearly identical for both the fluorescence lifetime measurement and the white actinic light PAM measurement, with the red actinic light PAM measurement showing faster relaxation. Comparing each technique’s plot of UNPQ,RCC and NPQ shows that the interpretation of the amount of qE can vary drastically depending on which parameter is used in calculations.
Concluding remarks By measuring fluorescence lifetimes while NPQ turns on and off, it is possible to ensure that changes attributed to NPQ are due to the changes in the chlorophyll-excited state relaxation dynamics rather than photobleaching or chloroplast movement. Having access to the whole fluorescence decay curve in addition to the average decay time and the fluorescence yield provides more information that can enable mechanistic inferences to be drawn. Using similar measurements to those described here on wild type and mutant A. thaliana leaves, we were able to distinguish the roles of the protein PsbS and the xanthophyll zeaxanthin in NPQ (Sylak-Glassman et al. 2014). In parallel, Bennett, Amarnath, and Fleming have developed a structure-based model for excitation energy transfer and qE in substantial (200 nm 9 200 nm) model membrane patches (publication forthcoming). In combination with the fluorescence lifetime snapshot technique described in this paper, the potential exists for a substantially improved understanding of photosynthetic light harvesting and its regulation. Acknowledgments This material is based upon work supported by the U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences, Chemical Sciences, Geosciences, and Biosciences Division. E. J. S.-G. was partially supported by a National Science Foundation Graduate Research Fellowship.
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EMERGING TECHNIQUES
Characterizing non-photochemical quenching in leaves through fluorescence lifetime snapshots Emily J. Sylak-Glassman1,2,3 • Julia Zaks2,4,5 • Kapil Amarnath1,2,6 Michelle Leuenberger1,2 • Graham R. Fleming1,2,4
•
Received: 19 November 2014 / Accepted: 16 February 2015 Ó Springer Science+Business Media Dordrecht 2015
Abstract We describe a technique to measure the fluorescence decay profiles of intact leaves during adaptation to high light and subsequent relaxation to dark conditions. We show how to ensure that photosystem II reaction centers are closed and compare data for wild type Arabidopsis thaliana with conventional pulse-amplitude modulated (PAM) fluorescence measurements. Unlike PAM measurements, the lifetime measurements are not sensitive to photobleaching or chloroplast shielding, and the form of the fluorescence decay provides additional information to test quantitative models of excitation dynamics in intact leaves.
Electronic supplementary material The online version of this article (doi:10.1007/s11120-015-0104-2) contains supplementary material, which is available to authorized users. & Graham R. Fleming [email protected] 1
Department of Chemistry, University of California Berkeley, Berkeley, CA 94720, USA
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Physical Biosciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA 94720, USA
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Present Address: IDA Science and Technology Policy Institute, 1899 Pennsylvania Avenue Suite 520, Washington, DC 20006, USA
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Graduate Program in Applied Science and Technology, University of California Berkeley, Berkeley, CA 94720, USA
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Present Address: Agilent Technologies, 5301 Stevens Creek Blvd, Santa Clara, CA 95051, USA
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Present Address: FAS Center for Systems Biology, Harvard University, 52 Oxford Street, Rm. 440, Cambridge, MA 02138, USA
Keywords Arabidopsis thaliana Chlorophyll Fluorescence lifetime Fluorescence yield Nonphotochemical quenching
Introduction Plants dynamically regulate the photosynthetic apparatus in response to fluctuations in light intensity (Blankenship 2002). To minimize damage (Barber 1995; Melis 1999) from excess light plants utilize a set of mechanisms collectively known as non-photochemical quenching (NPQ) to dissipate singlet excitations safely as heat (Demmig-Adams and Adams 1996). The most rapid component of NPQ is known as energy-dependent quenching (qE). An improved understanding of NPQ in general and qE in particular is of importance as mutant plants impaired in NPQ have decreased fitness (Ku¨lheim et al. 2002) and because altering the dynamics of NPQ may improve photosynthetic efficiency (Zhu et al. 2010). The amount of NPQ is generally measured through changes in the chlorophyll fluorescence yield, most often using a pulse-amplitude modulated (PAM) fluorometer. In a typical experiment, a dark-adapted leaf is monitored while it is exposed first to excess light to turn on NPQ and then to darkness as NPQ relaxes (Cazzaniga et al. 2013). While changes in the chlorophyll fluorescence yield are interpreted as the result of changes in the amount of NPQ, these fluorescence yield changes can have multiple causes. For instance, photobleaching and chloroplast movement can also decrease the fluorescence yield, thus complicating the interpretation of PAM data (Baker 2008). In contrast, the time-resolved fluorescence decay of chlorophyll is not sensitive to either photobleaching or chloroplast movement. The form of the fluorescence decay is sensitive to the
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number of quenchers and their qE rates and holds the potential for greater insight into the entire NPQ phenomenon. Because the amount of qE turns on and off in seconds to minutes, the fluorescence decays must be collected in time increments (‘‘snapshots’’) commensurate with the intrinsic timescales on which qE turns on and off. Amarnath et al. solved this problem by building a set of computer-controlled shutters to measure the fluorescence decay of Chlamydomonas reinhardtii for 80 ms every 200 ms, ensuring that the decay profile was effectively constant during the measurement (Amarnath et al. 2012). Repeated measurements were used to produce adequate statistics. In this paper, we describe the extension of this fluorescence lifetime snapshot approach to detached leaves of Arabidopsis thaliana (A. thaliana) plants.
Methods Measurements were made on medium-sized A. thaliana leaves detached from plants before the stage of bolting. Plants were grown in a growth chamber with 100 lmol photons 9 m-2 s-1 light. Measurements were carried out using a similar procedure to that described by Sylak-Glassman et al. (2014). Plants were each dark-adapted for 30 min prior to detaching a leaf. To prevent any plant from having its light cycle disturbed, measurements were performed on multiple plants during each day of data collection so that any given plant would not be dark-acclimated for more than 30 min in a 1.5 h period. Between dark adaptation times, plants were placed in a 100 lmol photons 9 m-2 s-1 light chamber for re-light acclimation. Each leaf was selected from the middle of the rosette to control for leaf age and size. After leaf selection, the leaf was immediately placed in a home-built holder (shown in Online Resource 1). To keep the leaf from overheating or dehydrating, the holder keeps most of the leaf open to air, and allows the leaf to take up water. Similar to the set-up described by Amarnath et al., the light exposure of the leaf and detector were controlled by shutters programmed in LabVIEW. The set-up is shown in Fig. 1a. The excitation pulse hits the leaf at 70° to the adaxial leaf surface (Fig. 1b). Excitation is into the chlorophyll a (chl a) Soret band at 420 nm. The average power of the laser at the sample was 5 mW with a pulse energy of 66 pJ. This laser power corresponds to an approximate intensity of 5000 lmol photons 9 m-2 s-1, which is enough to close reaction centers (Schansker et al. 2006) and is similar to the intensity used in PAM fluorometers (Brooks and Niyogi 2011; Cazzaniga et al. 2013; Porcar-Castell et al. 2008). An actinic light (Schott KL1500) was used to turn on qE. The leaf’s exposure to the actinic light was controlled
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by a shutter. The actinic light intensity was set at either 500 lmol photons 9 m-2 s-1 or 1200 lmol photons 9 m-2 s-1 for experiments. The light was collimated by a lens, forming a 1 cm diameter illumination spot size at a 45° angle to the adaxial side of the leaf. Detection was at a 45° angle to the adaxial side of the leaf and at a 65° angle to the laser. Fluorescence was collimated with a 5 cm lens and passed through a polarizer set to the magic angle (54.7°). The light was then focused into a monochromator (HORIBA Jobin–Yvon; H-20) with a 10 cm lens. The detection wavelength was centered at 684, 715, or 720 nm. 684 nm was chosen to look at fluorescence emanating from the Qy band of chlorophyll a, and 715 and 720 nm were chosen to mimic the spectrum of light collected by PAM fluorometers. The light from the monochromator was collimated with a 5 cm lens before being sent to the detector (Hamamatsu; R3809U). The instrument response function was 35–39 ps FWHM. Following Amarnath et al., the sequence of shutters opening and closing was designed in MATLAB and executed in LabVIEW (Amarnath et al. 2012). A typical measurement sequence consisted of an initial measurement taken on a dark-adapted leaf. After 2 min, the leaves were illuminated with actinic light. After 10 min of illumination, the actinic light was blocked, and the leaf was allowed to relax for 10 min. Data were collected at intervals ranging from 10 to 60 s during the light acclimation period and subsequent relaxation period in dark. Unless otherwise specified, identical data collection routines were performed on multiple leaves from multiple plants (6 leaves total from 3 different plants). The process for data summing, aligning, and fitting is identical to that described by Sylak-Glassman et al. (2014). The data from each leaf were aligned according to the maximum of a cross-correlation [xcorAlign.m (Carmichael 2010)]. The curves were fit to a sum of four exponentials: three decay functions and one fast rise (1–4 ps) (Picoquant Fluofit Pro 4.5). A comparison of the fits resulting from fitting the data to three decay functions versus three decay functions and a fast rise are shown in Online Resource 2. Following data fitting, curves were reconstructed by plotting the fluorescence intensity calculated by the exponential decays: P Ai et=si FðtÞ ¼ i P ; ð1Þ i Ai where Ai are the amplitudes and si are the fluorescence lifetime components. Amplitude-weighted average lifetimes of the leaves, savg, were calculated using the three exponential decays as follows: P i Ai s i : ð2Þ savg ¼ P i Ai
Photosynth Res Fig. 1 A top-view schematic of the apparatus is shown in the top (a), and the detection geometry is shown below (b). A diode laser pumps an ultrafast Ti:Sapphire oscillator. The resulting 840 nm light is frequency doubled to 420 nm. A portion of the beam is sent through a shutter to the leaf in its holder (green line). An actinic light is also sent through a shutter such that both beams can impinge on the leaf. Fluorescence is collected at the magic angle. The illumination and detection geometry is shown in more detail in b. The leaf is held at a 45° angle to the actinic light and the detector. The angle between the laser and actinic light is 25°
The standard deviations of the values of savg were calculated by performing bootstrapping on the lifetimes. Bootstrapping involves selecting a subset of points in the fluorescence decay and determining the fluorescence lifetime components required to fit the subset of points. Repeating this process one thousand times gives an estimate of the error in the fluorescence lifetime fitting parameters. The amplitudes were bootstrapped separately to determine the error in the amplitudes of each component.
Results We confirmed that detached leaves exhibit the same qE characteristics as leaves attached to an intact plant provided that the detached leaf is able to take up water during the measurement (Fig. 2a and Fig. 2b). The amount of qE is significantly reduced for detached leaves without access to water (Fig. 2c) In order to selectively track changes in the fluorescence decay profile due to NPQ rather than photochemistry, it is necessary to measure the fluorescence when the reaction centers are closed (Stirbet 2011). This corresponds to the longest fluorescence lifetime, and we determined via multiple fluorescence decay measurements on leaves while they are exposed to the laser for different lengths of time. In data collection, the total exposure time was 1 s total divided into five 0.2 s bins. This is shown schematically in
Fig. 3a. Figure 3b shows how the fluorescence lifetime changes over seven overlapping 0.2 s bins for leaves measured at 715 nm for wild type A. thaliana. Five leaves were summed for each 0.2 s bin, and the average fluorescence lifetime reached a peak at 0.4 s of laser exposure before the lifetime measurement. Once the bin with the longest lifetime is identified, it is used for further data analysis, and the other bins are discarded. The high concentration of chloroplasts in leaves leads to re-absorption and subsequent re-emission of fluorescence. This has a large impact on the leaf’s emission spectrum, which shows a peak at 735 nm that is absent in measurements on isolated chloroplasts (Moise and Moya 2004). It has been hypothesized that the re-emitted fluorescence prevents the accurate measurement of fluorescence lifetimes (Johnson and Ruban 2009). While relative changes in the fluorescence lifetime during qE would still be instructive, for the measurements to be used to parameterize models, the fluorescence lifetimes must be accurate. To verify the role of chlorophyll concentration on the measured fluorescence lifetime, the fluorescence lifetimes were compared between the spinach leaves and solutions of spinach chloroplasts with varying concentrations at two different wavelengths, 684 and 720 nm. These wavelengths were chosen to correspond to fluorescence from the Qy band of chlorophyll (684 nm) and to a wavelength range closer to that measured in a PAM fluorometer (720 nm). It is expected that the average fluorescence lifetimes at 684
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Fig. 2 The fluorescence decays for 3 leaves with different sample holder conditions and measured after 3 different illumination conditions are shown in (a, b, and c). The fluorescence decays for dark-adapted leaves are shown in a. The fluorescence lifetimes for the same leaves after 10 min of illumination are shown in b. The same leaves after 60 min of relaxation in the dark are shown in c. In each of
Fig. 3 Multiple fluorescence measurements are required to determine the maximum average lifetime. A schematic for how a single measurement period is subdivided is shown in a. The measurement period is 1 s long, during which time the leaf is exposed to the laser and fluorescence is detected. The measurement period is subdivided into 5 steps, each with a 0.2 s duration. The time increment with the maximum average lifetime (shown in gray, with the label peak savg), is the only step that is retained. The duration of laser exposure prior to the step with the maximum average fluorescence lifetime is referred to as the saturation time, because it is the time required to close all the PSII reaction centers. Measurements of savg summed for 5 darkadapted leaves are shown in b, measured at 715 nm. Exposure to the laser begins at 0 s. The horizontal bars indicate the time span over which the measurement was made. The vertical bars indicate the standard deviation in the average fluorescence lifetime value. The saturation time for this measurement is 0.3 s
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the images, the fluorescence decays for the leaf attached to a plant are shown in black, the fluorescence decays for the leaf detached from the plant with the stem immersed in water are shown in gray, and the fluorescence decays for the leaf detached from the plant without water are shown in red
and 720 nm will differ because of differing fractions of fluorescence from PSII and PSI antennas (Borisov and Il’Ina 1973; Moise and Moya 2004), each of which exhibits different chlorophyll energy transfer dynamics. The average fluorescence lifetimes are shown in Table 1. The spinach leaf and chloroplast solutions with a concentration of chlorophyll of 0.025 and 0.125 lg/lL all had nearly equivalent fluorescence lifetimes at 684 nm. At 720 nm, the spinach leaf has a fluorescence lifetime most similar to the lowest concentration of chloroplasts, 0.0025 lg/lL. Given that a typical spinach leaf has a chlorophyll concentration of approximately 2.7 lg/lL (Mehler 1951; Porra et al. 1989) and an average fluorescence lifetime similar to that of the chloroplast solutions, the average fluorescence lifetime does not show a concentration dependence over 3–4 orders of magnitude of chlorophyll concentration. The very dilute sample of chloroplasts (0.0025 lg/lL) displayed a slightly shorter average fluorescence lifetime. The consistency of the fluorescence lifetimes between the isolated chloroplasts and the leaves indicates that the fluorescence lifetimes that we measure for leaves are uncontaminated by non-primary fluorescence. To examine the effect of the actinic light intensity on the measured fluorescence lifetime, two actinic light intensities Table 1 Average fluorescence lifetimes of a spinach leaf and of spinach chloroplasts, measured at 684 and 720 nm [Chl] (lg/lL)
savg (ns) at 684 nm
savg (ns) at 720 nm
0.0025
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0.47 ± 0.03
0.025
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0.64 ± 0.03
0.125
1.44 ± 0.08
0.64 ± 0.03
Leaf
1.44 ± 0.07
0.51 ± 0.02
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were selected that are typically used in NPQ measurements: 500 and 1200 lmol photons 9 m-2 s-1. Lower actinic light intensities are not high enough to sustain NPQ (Kalituho et al. 2007), whereas higher light intensities can cause significant photodamage (van Wijk et al. 1993). When the fluorescence lifetimes are measured throughout the induction and relaxation of NPQ, it is possible to see the fluorescence lifetimes as a function of time during or after exposure to actinic light. The series of measurements can therefore be visualized in a two-dimensional plot of the fluorescence intensity as a function of time following excitation and as a function of the illumination time. Plots for leaves illuminated with 500 and 1200 lmol photons 9 m-2 s-1 light are shown in Fig. 4a and b, respectively. The normalized fluorescence intensity is calculated from the exponential decays used to fit each fluorescence lifetime measurement, and is displayed on a log scale. The horizontal axis is the timescale of the fluorescence decay in nanoseconds. The vertical axis is the timescale of light acclimation in minutes. The actinic light is on for 10 min, as shown by the white bar on the left in Fig. 4. The different dynamics for the induction of NPQ under two different illumination intensities can be seen by comparing the fluorescence intensity in the 2D plots. The leaves illuminated with 500 lmol photons 9 m-2 s-1 light have an initial shortening of the fluorescence lifetime during the first minute of illumination. Then, over the next 2 min, the decay lengthens. In minutes 3–5, the fluorescence lifetime shortens again, before stabilizing during the last 4 min of illumination. The dynamics of the induction of qE are different for the leaves illuminated with 1200 lmol photons 9 m-2 s-1 light. Similar to the 500 lmol photons 9 m-2 s-1 illumination leaves there is an initial
shortening of the fluorescence lifetime during the first minute of light acclimation. However, after this time, the fluorescence intensity gradually increases at long decay times (C4 ns) while it decreases at short decay times (B3 ns). The dynamics of the relaxation of NPQ for the leaves illuminated with 500 and 1200 lmol photons 9 m-2 s-1 light are very similar. Both experience an increasingly gradual lengthening of the fluorescence decays. Integrating over the fluorescence decay axis results in the amplitude-weighted average fluorescence lifetime, shown in Fig. 5. Because the average fluorescence lifetime is proportional to the fluorescence yield, the dynamics of the average lifetime should resemble the dynamics of the fluorescence yield under conditions of closed PSII reaction centers measured with PAM fluorometers. However, as noted earlier, unlike chlorophyll fluorescence yield measurements, the amplitude-weighted average fluorescence lifetime is not influenced by non-quenching processes such as chloroplast shielding and photobleaching. To assess the extent to which non-quenching processes influence the amount of NPQ measured by PAM fluorometers, we compared the NPQ dynamics from the fluorescence lifetime measurements with the NPQ dynamics as measured through PAM fluorometry. Because the actinic light color has been shown to influence chloroplast shielding, and in turn, the chlorophyll fluorescence yield, two different PAM fluorometers were used, each with a different color of actinic light. The Hansatech FMS fluorometer uses a white actinic light (Mouget and Tremblin 2002), while the Walz Dual PAM fluorometer uses a red actinic light (Heinz Walz GmbH 2006). Both PAM fluorometers detect at similar wavelengths, [705 nm for the white light PAM, and [700 nm for the red light PAM. The two different PAM
Fig. 4 Multiple measurements of the chlorophyll fluorescence lifetime for leaves illuminated with a 500 lmol photons 9 m-2 s-1 and b 1200 lmol photons 9 m-2 s-1 are represented in two-dimensional plots. The maximum fluorescence for each measurement is normalized to 1 and intensity is shown on a logarithmic scale. The average
decay time in nanoseconds is shown on the horizontal axis. The vertical axis is the time in minutes for the leaves’ acclimation to light. The actinic light turns on at 0 min, and is on for 10 min, at which point it is turned off for the remaining 10 min. The bars at the left of a indicate when the actinic light is on (white) and off (black)
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quantum yields of each component of NPQ when reaction center are closed (Ahn et al. 2009). Because this term is linearly related to the fluorescence yield, it more accurately tracks the changing excitation dissipation dynamics as NPQ turns on and off. The NPQ parameter from the PAM fluorescence measurements was calculated by comparing the fluorescence yield in the dark when PSII reaction centers are closed, Fm and the fluorescence yield after light exposure when PSII 0 reaction centers are closed, Fm : 0
Fm Fm NPQ ¼ 0 Fm
Fig. 5 The average fluorescence lifetime for leaves illuminated with 500 lmol photons 9 m-2 s-1 (gray) and 1200 lmol photons 9 m-2 s-1 (black). The bars at the top of the figure indicate when the actinic light is on (white) and off (black)
ð3Þ
The NPQ curves from the fluorescence lifetime measurements were calculated using an analogous equation [see Supplement by Sylak-Glassman et al. (2014) for derivation]: savg;dark savg;light NPQs ¼ ð4Þ savg;light
fluorometers were both set to an actinic light intensity of 1000 lmol photons 9 m-2 s-1. These illumination and detection parameters differ slightly from the fluorescence lifetime apparatus, which detects fluorescence centered at 684 nm and uses a white actinic light with an intensity of 1200 lmol photons 9 m-2 s-1, though this difference in intensity is not expected to have a significant impact on the amount of qE (Bailey et al. 2004). In addition to calculating the standard NPQ parameter, we also calculated the approximate quantum yield of NPQ with PSII reaction centers closed, UNPQ,RCC. This term was chosen because it also reports the amount of qE but, unlike the NPQ parameter, is linearly related to the fluorescence yield of the light-adapted leaf and is equal to the sum of the
A comparison of NPQ versus time for the fluorescence lifetime data and the two PAM fluorometers is shown in Fig. 6a. For the values of UNPQ,RCC from the PAM fluorescence measurements, the following equation was used:
Fig. 6 a NPQ and b UNPQ,RCC from fluorescence lifetime measurements (black), a red (620 nm) actinic light PAM (Walz Dual PAM) (red), and a white actinic light PAM (Hansatech FMS) (green). The fluorescence lifetime measurements have a white actinic light with an intensity of 1200 lmol photons 9 m-2 s-1 and the PAM measurements have an
actinic light intensity of 1000 lmol photons 9 m-2 s-1. The data points are shown as circles. The smoothed data are shown as lines. The bars at the top of the figures indicate when the actinic light is on (white) and off (black). Error bars corresponding to one standard deviation are shown for the NPQ PAM measurements
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/NPQ;RCC
Fm Fm ¼ Fm
ð5Þ
The calculation of UNPQ,RCC for the fluorescence lifetime measurements was adapted from the fluorescence yield measurements in the same manner as the NPQ parameter: savg;dark savg;light /NPQ;RCC;s ¼ ð6Þ savg;dark
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A plot of UNPQ,RCC,s versus time for the fluorescence lifetime data and the two PAM measurements is shown in Fig. 6b. The NPQ dynamics shown in Fig. 6a during the induction of qE are similar for the fluorescence lifetime measurement and the red actinic light PAM measurement. The two reach similar maximum values of NPQ as well. The slightly higher amount of NPQ from the fluorescence lifetime measurements compared to the white actinic light PAM measurement may be due to the difference in detection wavelength (Lambrev et al. 2010). The NPQ curve from the white actinic light PAM measurement displays an additional rise component and reaches a maximum value greater than 3, whereas the other two techniques reach a maximum value just under 2. However, the relaxation of NPQ is nearly identical for both the fluorescence lifetime measurement and the white actinic light PAM measurement, whereas the relaxation kinetics from the red actinic light PAM measurements are faster. The similarity between the amount of qE measured by red actinic light PAM fluorometers and the fluorescence lifetime measurements suggests that the chlorophyll fluorescence yield NPQ parameter from red actinic light PAM fluorometers is uncontaminated by nonquenching processes such as photobleaching or chloroplast avoidance. As suggested by Cazzaniga and co-workers, the extra NPQ seen in the white actinic light PAM measurements does not appear to be due to qE (Cazzaniga et al. 2013). While both the white actinic light PAM and the fluorescence lifetime measurements show a slower relaxation of qE than the red actinic light PAM, they most likely have different origins. The slower relaxation in white light PAM measurements has been shown to be due to a reversal of chloroplast shielding (Cazzaniga et al. 2013). The slower relaxation in fluorescence lifetime measurements must be due to a relaxation of qE processes, and could be due to a different response to the 420 nm laser light used in measurements compared to the saturating pulses of red or white light used in PAM fluorometers. The NPQ-related dynamics and values of all three techniques are much more similar when UNPQ,RCC is plotted instead of NPQ (Fig. 6b). The induction dynamics seen in the UNPQ,RCC curves for the fluorescence lifetime measurement and the red actinic light PAM measurement are still very similar, but the difference between these two techniques and the white actinic light PAM measurement is less drastic. Similar to the NPQ relaxation kinetics, the relaxation of UNPQ,RCC is nearly identical for both the fluorescence lifetime measurement and the white actinic light PAM measurement, with the red actinic light PAM measurement showing faster relaxation. Comparing each technique’s plot of UNPQ,RCC and NPQ shows that the interpretation of the amount of qE can vary drastically depending on which parameter is used in calculations.
Concluding remarks By measuring fluorescence lifetimes while NPQ turns on and off, it is possible to ensure that changes attributed to NPQ are due to the changes in the chlorophyll-excited state relaxation dynamics rather than photobleaching or chloroplast movement. Having access to the whole fluorescence decay curve in addition to the average decay time and the fluorescence yield provides more information that can enable mechanistic inferences to be drawn. Using similar measurements to those described here on wild type and mutant A. thaliana leaves, we were able to distinguish the roles of the protein PsbS and the xanthophyll zeaxanthin in NPQ (Sylak-Glassman et al. 2014). In parallel, Bennett, Amarnath, and Fleming have developed a structure-based model for excitation energy transfer and qE in substantial (200 nm 9 200 nm) model membrane patches (publication forthcoming). In combination with the fluorescence lifetime snapshot technique described in this paper, the potential exists for a substantially improved understanding of photosynthetic light harvesting and its regulation. Acknowledgments This material is based upon work supported by the U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences, Chemical Sciences, Geosciences, and Biosciences Division. E. J. S.-G. was partially supported by a National Science Foundation Graduate Research Fellowship.
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