Excitation−Emission Polarization Spectroscopy of Single Light

Apr 12, 2011 - A Berek compensator (BC) is placed in the excitation path to compensate ..... complexes with variable intensity reveals a few qualitati...
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ExcitationEmission Polarization Spectroscopy of Single Light Harvesting Complexes Sumera Tubasum,*,† Richard J. Cogdell,‡ Ivan G. Scheblykin,† and T~onu Pullerits† † ‡

Department of Chemical Physics, Lund University, P.O. Box 124, SE-22 100, Lund, Sweden Glasgow Biomedical Research Centre, University of Glasgow, G12 8QQ, United Kingdom ABSTRACT: Excitation and emission polarization dependence of fluorescence intensity of single LH2 complexes from Rhodopseudomonas acidophila 10050 and Rhodobacter sphaeroides is reported. The results are presented as two-dimensional polarization plots and interpreted in terms of tilted light harvesting complexes indicating that sample preparation leads to partially oriented LH2 cylinders. An alternative explanation of the observation can be structural deformation. Fluorescence intensity of the complexes has four qualitatively distinct excitationemission polarization dependencies. The differences in excitation polarization dependence are interpreted as due to the tilt of the complexes, whereas the emission polarization behavior is mainly determined by spectral inhomogeneity of the emitting B850 ring. Some complexes show abrupt reversible variations of the total emission intensity together with changes of the polarization properties which cannot be described by the simplest model of tilted LH2s with spectral disorder.

1. INTRODUCTION For optimal and flexible solar energy conversion, nature has divided the primary photosynthetic machinery into light harvesting antenna and reaction center (RC) pigment protein complexes.1 In the antenna complexes, energy of sunlight is absorbed and delivered in the form of electronic excitation to the RC where the energy is employed to drive unidirectional electron transfer along an electron donoracceptor chain. Energy of the separated charges is then used for following slower reactions. Important milestones on the way toward molecular level understanding of the primary light harvesting were resolving of the atomic structure of the so-called FennaMatthewsOlson antenna complex2 and of the bacterial reaction center.3 By now many structures of the photosynthetic pigmentprotein complexes are known.1 One of the complexes, the peripheral light harvesting antenna (LH2) of the photosynthetic purple bacterium Rhodopseudomonas (Rps.) acidophila, was resolved in 1995.4 LH2 is the main object of the current study. The LH2 from Rps. acidophila is a ring of nine pairs of Rhelixes. Each pair of helixes binds three bacteriochlorophyll a (BChl) and one (or possibly 2) carotenoid rhodopin glucoside molecules.4 The BChls form two rings of 9 and 18 molecules. The ring of 9 absorbs at around 800 nm, and the other ring at 850 nm. They are called B800 and B850, respectively. The BChls of the B850 ring are closely packed leading to significant intermolecular excitonic interactions which dominate the spectroscopy and excitation dynamics of the band.57 In the B800 ring the corresponding interactions are much weaker, and for interpreting the spectroscopic observations exciton theory is typically not needed.8 However, even in B800, novel electronic coherent two-dimensional (2D) spectroscopy9 has indicated some level of intermolecular electronic coherence. r 2011 American Chemical Society

Excitonic interaction between chromophores is not the only reason for the spectral shifts in pigment protein complexes.10 It has been shown that the neighborhood of a pigment can cause significant shifts via hydrogen bonding,11 axial ligations,12 structural distortions of the chromophore,13 local electric fields,14,15 and dispersion interactions.16 Compared to the excited state lifetime, the structural changes which are related to these shifts are slow, and because of the ensemble averaging, the shifts lead to static inhomogeneous broadening. The spectra also have a dynamic component due to nuclear motions causing additional broadening of the spectra.17 Strong coupling between electronic and nuclear degrees of freedom can lead to polaron formation as seen in LH2 at low temperatures.18 In a conventional spectroscopic experiment, a very large number of complexes are simultaneously measured. In order to avoid loss of information due to the ensemble averaging, singlemolecule spectroscopy (SMS) has become increasingly popular.19 A number of groups have performed experiments with single light harvesting complexes. Many different characteristics of single LH2s were studied like room temperature photobleaching and fluorescence lifetime,20 fluorescence spectral diffusion,21 low temperature fluorescence spectra,22 and fluorescence excitation spectra.23 New knowledge obtained via SMS studies of LH2 complexes was reviewed by Cogdell et al.24 Some SMS studies were interpreted in terms of elliptical distortions of LH2 rings.23,25 Calculated individual excitonic spectra of a ring, ellipse and ellipse with a gap, were thoroughly discussed in the context of SMS studies of bacterial light harvesting systems.26 Elliptical Received: August 9, 2010 Revised: March 10, 2011 Published: April 12, 2011 4963

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LH2 shape was also suggested based on the SAXS studies.27 At the same time simulations of single LH2 fluorescence did not require any assumptions about structural distortions.28 In the current contribution we use novel 2D polarization single-molecule imaging29 to study individual LH2 complexes. In the following section we describe the sample preparation technique and briefly introduce SMS setup. We proceed with explaining various ways of representing and analyzing experimental data. Finally the results are presented and discussed in the context of known structural and spectroscopic data.

2. MATERIALS AND METHODS 2.1. Sample Preparation. A working solution of pM concentration is prepared by diluting a stock solutions of LH2 isolated from either Rps. acidophila or Rhodobacter (Rb.) sphaeroides (method of their isolation is described in ref 30) with 25 mM Tris-HCl buffer solution (pH = 8, 0.1% LDAO). The buffer solution is made oxygen-free by adding an oxygen scavenging system to the solution consisting of D-(þ)-glucose, glucose oxidase, and catalase (all from Sigma Aldrich) in prescribed ratios.28 For matrix formation, a solution of purified poly(vinyl alcohol) (Clariant, Mw = 30 000, 1.5% solution in buffer) is mixed with the working sample solution in 1:1 volume ratio. A microscope coverslip (0.17 mm) is cleaned by repeated ultrasonification and treatment with UV light. A 50 μL drop of the polymerLH2 working solution mixture is smeared over the coverslip and is spin coated at high speed of 3000 rpm to get uniform monolayer. The thickness of the film measured with a Veeco DekTak 6 M stylus profiler is ∼60 nm. 2.2. Experimental Setup. Experimental setup is described in Figure 1. A CW Ti:Sapphire laser (Spectra Physics 3900S) is pumped by a Spectra Physics Millennia Pro. The laser beam is spectrally cleaned up with a narrow band-pass filter at 800 nm. A wire grid polarizer (P) provides a vertically linearly polarized excitation beam. The polarization plane of excitation is varied by a rotating half-wave plate. The beam is then passed through a wide field inverted microscope (Olympus IX 71) and is focused at the sample plane by an objective lens (60, NA = 1.25, oil immersion). A Berek compensator (BC) is placed in the excitation path to compensate for the loss of polarization due to optical elements before the sample plane. The emission from sample is collected by the same objective lens and is separated from the excitation path by a dichroic mirror (DM). The emission is then passed through two long-pass filters, LP 830 nm and LP823 nm. A home-built analyzer containing a set of a rotating wire grid polarizer (R) and a fixed reflecting mirror (T) is used to split the emission into two beams with mutually perpendicular polarization as follows.29 The rotating polarizer (R) reflects the emission component IR perpendicular to its polarizing plane, while the parallel component IT is transmitted through the polarizer and is then reflected by the mirror (T) behind. Both components are independently imaged by an EMCCD (PhotonMax 512, 512  512 pix; 1 pix =16 μm). Data collection is synchronized with rotation of the half-wave plate and the analyzer by light emitting diodes (LEDs) connected to the rotating motors. We show typical fluorescence intensity traces of a single LH2 complex from two orthogonal channels together with synchronizing LED signals (Figure 1).

3. DATA COLLECTION AND ANALYSIS We excite B800 BChls and detect the emission from B850 BChls. A series of images of fluorescence intensity is collected by

Figure 1. Experimental setup. Excitation beam (red solid line) with rotating polarization and emission (black dotted line), which is split into two components perpendicularly polarized with respect to each other. These two components are individually imaged by a CCD. P = polarizer, F1 = excitation filter (800 nm), F2 = long-pass filter (823 nm), DM = dichroic mirror, BC = Berek compensator, SF = synchronizing flash, R = polarizer, T = mirror, and OI = oil immersion objective lens. Inset shows intensities of two mutually perpendicular emission components detected from two different spots on the CCD which correspond to the same LH2. Fast intensity modulation is due to the excitation polarization rotation, whereas the slow modulation is due to the detection rotation. Synchronization signals from the LED are also shown. The absorption spectra of LH2 solution for two bacterial species along with the laser excitation spectrum are shown in the left upper corner.

simultaneously rotating the excitation and the detection polarization planes. The frequency of rotation is configured to get on average 152 excitation modulations and 18 detection modulations within 100 s data acquisition time. A dye solution (IR-797 Chloride, Sigma Aldrich) is used to characterize the isotropicity of the experimental setup and correct for the setup-induced excitation and detection anisotropy. The LH2 complexes vary in their fluorescence intensities as excitation intensity at the sample plane is not constant. With an excitation spot of ∼30 μm in diameter, the average excitation power is 3.2 kW/cm2. For polarization data analysis, LH2 complexes are identified and their fluorescence intensities traced out with the help of homemade software developed by Mirzov et al.29 Plotting and fitting of the polarization data have been done with Matlab. 3.1. Photostability. Carotenoids are protecting the BChls from photoxidation by quenching their triplet state and singlet oxygen.31 Excitation intensities in SMS experiments are high— on average a complex is excited once per ∼20 ns. This means that LH2s in our experiment are susceptible to photooxidation. Indeed, if the sample is in open air, all the complexes show rapid bleaching within a time frame of less than 50 s, as shown in Figure 2a. From earlier work of C. Tietz,22 it is concluded that photostability achieved at cryogenic temperature is the result of reduction in oxygen diffusion. As a first step in sample stabilization, we use oxygen scavengers. In this case, if the sample is in open air during the measurements, rapid but this time mainly reversible bleaching takes place. Besides, the majority of the studied LH2s show clear blinking in these conditions; see Figure 2b. Clear improvement in sample stability is achieved if N2 atmosphere is used for experiment, Figure 2c. Still many LH2s show bleaching within 100 s in these conditions. As 4964

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The Journal of Physical Chemistry B expected, the sample with oxygen scavengers kept in N2 gas is found to be the most stable leading to almost constant signals as shown in Figure 2d. A few complexes prepared in this way bleached within 100 s. All polarization measurements have been carried out with the most optimized (Figure 2d) strategy. The complexes which entered dark state during the measurement time were removed from the analyses. About 15% of the complexes showed reversible variations in fluorescence intensity. Such complexes were included in the analyses. LH2s from Rb. sphaeroides were found somewhat less bright and less stable compared to Rps. acidophila 10050. 3.2. Polarization. Fluorescence intensity F(jex, jem) measured as a function of independently varying excitation jex and emission polarization angles jem can be represented as 2D plots shown in Figure 3a. The horizontal axis corresponds to jex while a vertical axis is jem. Figure 3a shows the measured 2D plots (frames) during the data collection time of ∼100 s, their averaged 2D plots, and the fitting plots of a LH2 complex. It takes ∼8 s to record a full 2D plot frame. The first row of plots is the total fluorescence intensity (I) plots which is the sum of

Figure 2. Characteristic examples of fluorescence intensity transients of single LH2s from Rps. acidophila 10050 with four different sample preparations. Pure sample (a), oxygen scavengers (b), N2 gas atmosphere (c), both oxygen scavengers and N2 atm (d).

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transmitted (IT) and reflected (IR) channels. The third row of plots is intensity from transmitted channel (IT), while the middle row shows the normalized intensity (Inorm= IT/I) plots. The total fluorescence intensity trace for the analyzed molecule is also shown in Figure 3b. Each part of such fluorescence trajectories can be directly related to the certain frames in Figure 3a. 3.3. Model and Fitting. In a multichromophoric system with excitation energy transfer, like LH2, detected fluorescence intensity can be expressed as a function of excitation and detection polarization and wavelength: Fðjex , jem , λex , λem Þ ¼

N

~ ij ð dBj 3 eB Þ2 Aj ðλex Þ ð dBi 3 eBem Þ2 Ei ðλem ÞG ∑ ex i, j ¼ 1

ð1Þ

Here N is the number of chromophores, dBi is the transition dipole moment of the chromophore i, and eBex and eBem are the unit vectors along excitation and detection polarization, respectively. Ei(λem) is the emission spectrum of the chromophore i at wavelength λem, and Aj(λex) is the absorption spectrum of the ~ ij is the time chromophores j at the excitation R ¥wavelength λex 3 G integral of the Green function 0 Gij(t) dt. The Green function gives the probability that the initial excitation at the chromophore j at t = 0 resides at the chromophore i at the time moment ~ ij is the average probability that t.32 Thereby the transfer matrix G the excitation, which initially was created at the chromophore j, is at chromophore i. The emission intensity from a chromophore is proportional to the probability that the chromophore is in the excited state. In the case of strong exciton coupling, instead of chromophores, the exciton states have to be used; otherwise, the same formalism is applicable. In conventional experiment with bulk sample, eq 1 is to be averaged over all possible orientations of LH2s. In SMS a single LH2 is detected and no averaging is needed. However, the specific orientation of the LH2 with respect to the laboratory axes is not known. In order to analyze the experimental 2D polarization plots, we use a model developed by Mirzov et al.29 This is the minimal model needed to describe the 2D polarization plots of

Figure 3. Time-dependent polarization plots of a single LH2 (Rps. acidophila 10050). The emission intensity is represented as 2D maps where the horizontal and vertical axes correspond to the excitation and detection polarization angles, respectively. (a) Three rows of 2D polarization plots for total intensity (I = IT þ IR), normalized intensity (Inorm), and transmitted intensity (IT). The ÆAvæ represents the average of the frames in the corresponding rows. The last column is the fitting of the average plots to the model (see text). (b) Time profile of the total fluorescence intensity (I) which is used to construct the 2D plots. 4965

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multichromophoric system with excitation transfer. The general eq 1 with N transition dipoles is reduced to a simplified system with three symmetric dipoles in a plane perpendicular to the optical axis of the setup. A detailed description of the model can be found in ref 29. Briefly, 2D polarization plots are expressed as Fðjex , jem Þ ¼

3

~ ij 3 Ij ð! ð! μ i 3 eBem Þ2 G μ j 3 eBex Þ2 ∑ i, j ¼ 1

ð2Þ

! μ i is the unit vector pointing in the direction of the ith transition. Ij is the cross section of the transition j. In order to reduce the amount of the free parameters, it is assumed that I2 = I3 and dipoles 2 and 3 are symmetric with respect to the first dipole. The angle between the dipoles 1 and 2 is called the spread angle γ, and it defines the mutual orientation of all three dipoles. The transfer matrix conserves excitation thereby adding condi~ ij = 1.This restriction reduces the number of tion the ∑iG independent transfer matrix elements to 6. As experimentally shown in ref 29, this is a minimal model for describing a general 2D SMS polarization measurement of multichromophoric system with excitation transfer. The model has been earlier used for conjugated polymers29 and for biomolecular complexes33 and is applied here to fit the experimental 2D polarization plots as shown in Figure 3. The experimental data and the model parameters can be used to derive a set of quantitative characteristics of the system and the ~ ij energy transfer. Nondiagonal elements of the transfer matrix G lead to a change in the polarization angle of the observed fluorescence. Consequently one can construct a parameter which corresponds to average absolute polarization rotation angle due to excitation transfer29 hjRrot ji ¼

3

∑ Rij GBij Ij i, j ¼ 1

ð3Þ

where Rij is the angle between dipoles Ii and Ij; |Rij| e 90. By averaging I over jex, one obtains conventional onedimensional (1D) excitation polarization curves with a functional form IðjÞ ¼ I0 f1 þ Mex cos½2ðj  j0 Þg

ð4Þ

where I0 is a constant, Mex is polarization modulation depth, and j0 is the phase. If Imax and Imin are the maximum and minimum of the 1D polarization curves, the modulation depth can be obtained as Mex ¼

Imax  Imin Imax þ Imin

ð5Þ

The parameters M and j0 can be also found for emission using either IT or the outcome of the fitting. The emission and excitation parameters are generally different. The difference of excitation and emission j0 is called polarization phase shift Δj0.

4. RESULTS AND DISCUSSION 4.1. Polarization Modulation Depth Distribution Function of Circular Aggregate. In the following discussion we relate the

known LH2 structure and statistics of measured excitation and emission modulation depths. In the discussion we assume that all 9 B800 BChls of a specific LH2 have the same absorption spectrum (high temperature) and that efficient B800 to B850 transfer takes place. The excitation laser beam (coaxial with the

Figure 4. LH2 with definition of tilt angle θ. Theoretical distributions assume random orientation of LH2.

detection) defines the laboratory axis z (optical axis), whereas xy is the microscope sample plane. We take an ideal circular aggregate, like B800 ring, oriented in such a way that the circle plane coincides with the laboratory xy-plane. Such LH2 is isotropic with respect to excitation polarization, and it would not show any emission intensity modulation while excitation (detection) polarization is rotated. Apparently this is not the case in our SMS study (see Figure 3). An obvious explanation can be that the LH2s in our experiment are randomly oriented. If the ring is tilted as in Figure 4, then it has an elliptic projection to the xyplane. The projection of the transition dipole moments of B800 BChls in this plane follows the same elliptic trend. If excitation polarization is rotated, then, because of the elliptical projection, the amount of excitation, and thereby the emission intensity, is modulated. At the same time, the maximum signal should not depend on the tilt. If the polarization vector is perpendicular to the longer axis of the ellipse, the amount of absorbed light is proportional to the cos2 θ, where θ is the tilt angle; see Figure 4. By using the eq 5 we can express the modulation depth as Mex ¼

1  cos2 ðθÞ 1 þ cos2 ðθÞ

ð6Þ

If we know modulation depth, we can calculate from eq 6 the corresponding tilt angle as pffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi θ ¼ arcos ð1  Mex Þ=ð1 þ Mex Þ ð7Þ We have measured the excitation and emission modulation depths for 95 Rps. acidophila 10500 and 26 Rb. sphaeroides LH2s and used eq 7 to find the corresponding tilt angles within the above model. The modulation depth and tilt angle distribution functions are presented for Rps. acidophila in Figure 5. In the following we will find the expected tilt angle distribution function P(θ) in the case of random orientation of LH2s. 4966

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Figure 5. Distribution functions of experimental and theoretical tilt angles P(θ) (a) and modulation depths P(M) (b) of Rps. acidophila 10500.

Table 1. Variance σ2 and Mean μ of Mex and Mem of Rps. acidophila 10500 and Rb. sphaeroides σ2ex/μex

σ2em/μem

Rps. acidophila 10500

0.05/0.61

0.05/0.72

Rb. sphaeroides

0.03/0.51

0.03/0.54

We draw the unit vector starting from the origin of the Cartesian laboratory coordinate system parallel to the principal axis of the LH2. All possible LH2 configurations of the same θ are represented by allowing the principal axis of the LH2 to rotate around the z-axis keeping the tilt angle θ unchanged. The unit vector draws a circle with the radius sin θ. The density of the LH2 orientations with the tilt angle θ is proportional to the circumference of this circle. Thereby, we obtain PðθÞ ¼ sin θ

ð8Þ

This gives the following modulation depth distribution function: 1 PðMex Þ ¼ pffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi 2 ð1 þ M Þ 1  Mex ex

ð9Þ

The theoretical random tilt angle and M (eqs 8and 9) together with the corresponding experimental data are shown in the Figures 5a and 5b, respectively. Apparently, the theoretical distributions based on random orientation of LH2s do not fit our experimental results. The experimental distributions are obtained from eq 7. They are quite similar for excitation and emission. The mean for excitation-based distribution is at about 0.6 rad (∼35). Table 1 gives the variances σ2 and the means μ of the Mex and Mem of the two studied species. The experimental tilt angle distributions are somewhat affected by the high NA of the microscope objective. This can lead to lower values of observed modulation depths since the isotropic z-component of polarization may start playing a role. The control experiments with lower NA objective (0.65) indicated that the effect is not very significant. An LH2 has a height of ∼6 nm and a diameter of ∼8 nm. While spin-coated film is drying, shear forces are expected to orient such cylinders upright. This is the most likely explanation why LH2s are not randomly oriented. An alternative explanation for the observed polarization dependencies is the possible deformation of the LH2. At this point we cannot exclude the possibility that because of the shear forces the LH2s are of elliptical shape,23 which would lead to analogous behavior as observed here. In order to distinguish

Figure 6. Scatter points Mem versus Mex for 95 Rps. acidophila 10050 (a) and 26 Rb. sphaeroides (b) antenna complexes. (a) is tentatively divided into four regions representing qualitatively differently behaving LH2s (see text).

between these possibilities, we are planning experiments with more controlled sample preparation methods.34 4.2. Excitation and Emission Modulation Depth Correlation. Even though the excitation and emission modulation depth distributions are very similar, the modulation depths of a certain LH2 can differ significantly. In Figure 6 we have presented all measured LH2s as scatter points Mem versus Mex. Clearly, the points do not follow the diagonal line. The correlation coefficient between Mem and Mex R ¼ 2

∑i ðMexi  μex ÞðMemi  μem Þ σex σem

ð10Þ

was found to be 0.35 for Rps. acidophila and 0.12 for Rb. sphaeroides. Relatively low correlation between Memand Mex indicates that the excitation and emission polarization properties have different origin. 4.3. 2D Polarization Plots of LH2s. We have selected four extreme cases to illustrate different outcomes of the SMS 2D polarization analysis (see Figure 7). The selected LH2s are labeled by different symbols in Figure 6a and represent four tentative areas in the figure. The LH2 (a) (diamond in Figure 6a) has negligible Mex and Mem. This is the expected behavior of an LH2 with θ = 0 (see Figure 4). In this case the random structures in the 2D plot are due to the noise. We point out that the color coding of the 2D plots is stretched out between minimum and maximum of the plot values. For (a) this means that despite large changes in the colors the 2D plot represents a relatively flat surface with modest random fluctuations. The situation is very different for (b) (square in Figure 6a). This LH2 has the largest Mex times Mem among the investigated complexes. The total intensity I plot shows a prominent ridge at jex = 80. Similarly, the normalized transmitted intensity plot has a ridge at jem = 75. This type of behavior is expected from an LH2 which is tilted by θ = 65. The example (c) (triangle in Figure 6a) has negligible Mex while Mem is relatively large. Accordingly, the I plot does not have any regular structure, and it is mainly due to noise. Inorm, however, has a ridge at jem = 170. This behavior can correspond to an LH2 with θ ≈ 0 but where most of the emission comes from a single B850 exciton state leading to significantly polarized emission. Such a state can be a result of strong disorder of B850 BChls. Contrary to the previous one, (d) has a large Mex = 0.37 while Mem is small. The high Mex value corresponds again to a 4967

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Figure 7. Characteristic examples of 2D polarization plots for four different regions of Figure 6a. Each panel presents the total intensity plot I and the normalized intensity plot Inorm together with the corresponding modulated time profiles. Panels correspond to the symbols of Figure 6a as follows: (a) (, (b) 9, (c) 2, and (d) b.

pronounced ridge in the I plot. Even though the Mem is small, the structure in the Inorm plot can be clearly distinguished. In the context of tilted light harvesting complexes, Mex = 0.37 corresponds to θ = 20. At the same time emission is almost isotropic. This can be the result of emission from more than one state which together by chance compensates the tilt-related anisotropy in the system. Such a coincidence cannot occur very often, and, indeed, we have found only very few LH2s which show such behavior. Based on these examples we can divide the (Mex, Mem) plot (Figure 6a) into four regions. The borderlines between the regions are somewhat arbitrary. Particularly the regions represented by complexes (a) and (b) gradually change from one to another. However the extreme cases of Figure 7 have important qualitative differences. In Figure 8 we have illustrated these cases via absorbing B800 ring with a tilt angle leading to differences in the Mex and two perpendicular diploes representing emitting exciton states of the B850. In (a) and (b) the tilt at the LH2 and emitting dipoles are well correlated, whereas in (c) and (d) there is no correlation. The lack of correlation is mainly due to the disorder in the system. Because of a random realization of disorder, the exciton structure may become such that the emission of a LH2 is considerably anisotropic. Since the disorder and the tilt are not correlated in any way, the emission polarization orientation can differ significantly from the orientation of the main absorption. With an excitation rate of 1 excitation per 20 ns, triplet population can build up in the carotenoid pool since the carotenoid triplet lifetime is in the microsecond region.35,36 This opens an

Figure 8. Interpretation of Figures 6 and 7 in terms of tilted absorbing B800 ring and two perpendicular emitting B850 dipoles representing strongly allowed main emitting states. The symbols are as in Figure 6a. The panels of the Figure 7 correspond to the symbols as (a) (, (b) 9, (c) 2, and (d) b.

extra dissipation channel by singlettriplet annihilation.37 Since the quenching triplet state is well localized, it may generate certain asymmetry in the B850 excited state population which can lead to anisotropic emission. However, since excitation dynamics in B850 occurs in subpicosecond time scale,7,38 it is unlikely 4968

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Figure 9. Excitation polarization dependence of the total emission intensity I together with excitation and emission modulation depths for four complexes showing abrupt intensity changes.

that this type of asymmetric quenching can explain the observed polarization effects because of the rapid equilibration. 4.4. Time Dependence. It takes about 8 s to record the data for one frame in Figure 3. Clearly, there are differences between frames, but they remain within the level of noise. None of the observed complexes has shown any clear time dependence of the polarization angle of maximum intensity of the I plots. This means that the total fluorescence intensity curves follow the ideal cosine excitation polarization angle dependence in eq 4 without any change of phase. Even the complexes which demonstrated clear abrupt emission intensity jumps did not show any discontinuity of the phase. In the context of tilt-dependent excitation modulation, this means that the tilt angle and orientation of the

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LH2s do not change. For our matrix-embedded samples, this is an expected behavior. Bopp et al.25 in their excitation SMS polarization sweep experiment observed changes in the angle of excitation polarization where the emission intensity is maximal. In a few LH2s the changes occurred in the time range of seconds. In that work the LH2s were immobilized on mica surface which allows for some flexibility of the tilt, hence the changes in the excitation anisotropy. Analogously, maximum excitation polarization angle of a fluorophore covalently bound to a single-strand DNA was shown to have a small probability to change.39 Such changes were related to the possible changes of fluorophore orientation. In multichromophoric systems, phase jumps in excitation polarization can also occur because of bleaching of a part of the system or changes in excitation transfer pathways.40,41 The majority of the complexes had stable modulated intensity time profiles like in Figure 3b. In about 15% of the complexes we observed reversible intensity changes. Most of the changes were abrupt within the time resolution of the experiment, whereas three complexes showed gradual changes in the signal intensity. A closer look at the excitation modulation patterns of the complexes with variable intensity reveals a few qualitatively different behaviors shown in Figure 9. In the complex (a) the intensity jump is such that the modulation depth does not change. The change is most likely due to a quencher close to the B850 ring which reduces the total emission intensity approximately by half. After about 30 s the quencher disappears. In the complex (b) the intensity changes occur without noticeable changes in the modulation amplitude which means that the modulation depth changes quite a lot. The effect is particularly large for emission modulation depth. Complex (c) has the same behavior when the intensity jumps down while the modulation amplitude increases noticeably when the intensity is recovered. As a result the change in the excitation modulation depth is quite pronounced, whereas Mex does not vary so much. (d) shows very obvious intensity jumps without noticeable changes in the amplitude of the modulation. Again, the outcome is a clear modulation depth variation. In (d) emission and excitation modulation depths closely follow each other. The changes observed for complexes (b), (c), and (d) cannot be explained in the context of simple tilted ring model. Clearly, the isotropic part of absorption is changed in those complexes leaving the anisotropic part largely unchanged. A possible origin of such behavior is spectral inhomogeneity of the B800 band. The observation can also originate from mixing of the B800 transitions with B850 ring excitons which are resonant with B800 and can partially lose their forbidden character due to the mixing. Such mixing would be a direct analogue to the B800 to B850 energy transfer description which takes into account strongly coupled multichromophoric nature of the accepting B850 ring.42

5. CONCLUDING REMARKS We report 2D polarization single-molecule imaging results of LH2 complexes from two purple bacteria. Based on excitation polarization results, we conclude that cylindrical LH2 complexes are preferably upright oriented in our sample. The orientation of the complexes is most probably due to the shear forces during drying of the sample film. Correlation between excitation and emission polarization effects was found to be very low, indicating that they have different origins. The emission polarization behavior is explained in terms of the disordered exciton model of the B850 ring. A minor part of the LH2 complexes showed 4969

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’ AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected].

’ ACKNOWLEDGMENT We thank Oleg Mirzov, Daniel Thomsson, Hongzhen Lin, and Yuxi Tian for valuable discussions. We thank David Adolph and nmC@LU for technical help with film thickness measurements. This work was financially supported by the Swedish Research Council, KWA Foundation, and STEM. S.T. is indebted to Swedish Institute and Higher Education Commission of Pakistan for financial assistance. ’ REFERENCES (1) Blankenship, R. E. Molecular Mechanism of Photosynthesis; Blackwell Science: Oxford, U.K., 2002. (2) Fenna, R. E.; Matthews, B. W. Nature 1975, 258, 573–577. (3) Deisenhofer, J.; Epp, O.; Miki, K.; Huber, R.; Michel, H. Nature 1985, 318, 618–624. (4) McDermott, G.; Prince, S. M.; Freer, A. A.; HawthornthwaiteLawless, A. M.; Papiz, M. Z.; Cogdell, R. J; Isaacs, N. W. Nature 1995, 374, 517–521. (5) Pullerits, T.; Sundstr€om, V. Acc. Chem. Res. 1996, 29, 381–389. (6) Monshouwer, R.; Abrahamsson, M.; van Mourik, F.; van Grondelle, R. J. Phys. Chem. B 1997, 101, 7241–7248. (7) Trinkunas, G.; Herek, J. L.; Polivka, T.; Sundstr€om, V.; Pullerits, T. Phys. Rev. Lett. 2001, 86, 4167–4170. (8) Hess, S.; Åkesson, E.; Cogdell, R. J.; Pullerits, T.; Sundstr€om, V. Biophys. J. 1995, 69, 2211–2225. (9) Zigmantas, D.; Read, E. L.; Mancal, T.; Brixner, T.; Gardiner, A. T.; Cogdell, R. J.; Fleming, G. R. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 12672–12677. (10) Timpmann, K.; Ellervee, E.; Pullerits, T.; Ruus, R.; Sundstr€ om, V.; Freiberg, A. J. Phys. Chem. B 2001, 105, 8436–8444. (11) Hess, S.; Visscher, K. J.; Pullerits, T.; Sundstr€om, V.; Fowler, G. J. S.; Hunter, C. N. Biochemistry 1994, 33, 8300–8305. (12) He, Z.; Sundstr€om, V.; Pullerits, T. J. Phys. Chem. B 2002, 106, 11606–11612. (13) Gudowska-Nowak, E.; Newton, M. D.; Fajer, J. J. Phys. Chem. 1990, 94, 5795–5801. (14) Kjellberg, P.; He, Z.; Pullerits, T. J. Phys. Chem. B 2003, 107, 13737–13742. (15) Herek, J. L.; Polivka, T.; Pullerits, T.; Fowler, G. J. S.; Hunter, C. N.; Sundstr€om, V. Biochemistry 1998, 37, 7057–7061. (16) Pullerits, T.; Mirzov, O.; Scheblykin, I. G. J. Phys. Chem. B 2005, 109, 19099–19107. (17) Pullerits, T.; van Mourik, F.; Monshouwer, R.; Visschers, R. W.; van Grondelle, R. J. Lumin. 1994, 58, 168–171. (18) Polivka, T.; Pullerits, T.; Herek, J. L.; Sundstr€om, V. J. Phys. Chem. B 2000, 104, 1088–1096. (19) Moerner, W. E.; Orrit, M. Science 1999, 283, 1670–1676. (20) Bopp, M. A.; Jia, Y.; Li, L.; Cogdell, R. J.; Hochstrasser, R. M. Proc. Natl. Acad. Sci. U.S.A. 1997, 94, 10630–10635. (21) Rutkauskas, D.; Novoderezhkin, V.; Cogdell, R. J.; van Grondelle, R. Biochemistry 2004, 43, 4431–4438. (22) Tietz, C.; Chekhlov, O.; Dr€abenstedt, A.; Schuster, J.; Wrachtrup, J. J. Phys. Chem. B 1999, 103, 6328–6333.

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