- Email: [email protected]
Ultrafast unequilibrated charge transfer: A new channel in the quenching of fluorescent biological probes
Chemical Physics Letters 412 (2005) 158–163 www.elsevier.com/locate/cplett
Ultrafast unequilibrated charge transfer: A new channel in the quenching of fluorescent biological probes Chaozhi Wan, Tianbing Xia, Hans-Christian Becker 1, Ahmed H. Zewail
*
Laboratory for Molecular Sciences, Arthur Amos Noyes Laboratory of Chemical Physics, Division of Chemistry and Chemical Engineering, California Institute of Technology, Mail Code 127-72, Pasadena, CA 91125, USA Received 8 June 2005; in final form 20 June 2005 Available online 19 July 2005
Abstract The dynamics of two biological fluorescent probes, 2-aminopurine (Ap) and daunomycin, were studied using both femtosecond transient absorption and fluorescence upconversion techniques. Various Ap-containing structures were investigated in solution: free Ap, non-covalently bonded (with guanine, adenine, and tryptophan) and covalently bonded in DNA constructs (with guanine, 7deazaguanine, and adenine). The distinct difference of transient absorption and fluorescence dynamics on the ultrafast time scale, and their dependence on free energy change (DG), and the abrupt decrease of the initial fluorescence intensity suggest the efficient depopulation by charge transfer from the unequilibrated hot molecules. We provide a model for this possibly general mechanism and obtain the rate constants for charge separation, vibrational relaxation, and charge recombination. Ó 2005 Elsevier B.V. All rights reserved.
1. Introduction As a structural analog of adenine, 2-aminopurine (Ap) can form base pairs with other bases, exhibiting reasonable stability and minimal perturbations to DNA structures. Ap can be excited selectively in the presence of other bases [1], and it has high fluorescence quantum yield (0.6–0.7) [2,3], but sensitive to the environment [3–6] and surrounding structures [7]. Ap has been extensively used as a probe molecule [8] to study DNA and RNA structures and conformational dynamics [9–12], DNA hydration [13], recognition of DNA by polyamides [14] and RNA by peptides [15,16], and enzymatic activity of various DNA-binding proteins. The usefulness of Ap has prompted extensive studies of its fundamental physical properties, such as molecular *
Corresponding author. Fax: +1 626 792 8456/405 0454 . E-mail addresses: [email protected], [email protected] (A.H. Zewail). 1 Present address: Department of Physical Chemistry, Uppsala University, Box 579, SE-751 23 Uppsala, Sweden. 0009-2614/$ - see front matter Ó 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.cplett.2005.06.101
geometry, tautomerism and dipole moments [17], directions and strength of electronic transition moments [18], electronic structure [19], and state ordering and energy gaps [20]. The goal was to understand the basis for the very different luminescence properties between the two isomers, adenine and 2-aminopurine [21]. Of particular usefulness is the use of Ap as a probe for charge transfer (CT) between excited state Ap and nucleobases in various DNA assemblies, in both timeresolved [22–24] and steady-state measurements [25, 26]. Charge transfer reactions lead to shortening of the lifetime of excited Ap, from the nanosecond (ns) regime for free Ap to a wide range within the picosecond (ps) regime in DNA, depending on the nature and distance between the donor and acceptor [22]. The direction of charge transfer (electron transfer vs. hole transfer) depends on the identity of the interaction partner with Ap [23]. This non-radiative decay pathway results in a large reduction of the fluorescence quantum yield. While the time-resolved methods directly measure the decay rates, steady-state methods infer the change in rates
C. Wan et al. / Chemical Physics Letters 412 (2005) 158–163
from measurement of overall fluorescence quantum yield and assuming that the decrease is caused by the charge transfer processes. However, this inference is valid provided there are no other non-radiative channels. Recently, Larsen et al. [27,28] proposed the existence of a long-lived non-radiative state for Ap, based on the wavelength dependent anisotropy decays in transient absorption. More recently a non-radiative state was proposed to explain the observed strong fluorescence quenching of Ap dinucleotides, but maintaining that the ps charge transfer still plays an important role in the quenching [29]. In these studies either transient absorption was performed or fluorescence decays were measured by time correlated single photon counting with limited resolution of 50 ps. However, the fluorescence quenching may also occur on much shorter time scales. Also the nature of the non-fluorescent state is unknown. In this work, we have used both fluorescence upconversion and transient absorption spectroscopy with femtosecond time resolution to systematically investigate the dynamics in a series of Ap-containing structures: free Ap, covalently bonded and non-covalently bonded complexes. Attention was paid to the early dynamics on the time scale of 200 fs and longer. On this ultrafast time scale, our analyses suggest the depopulation of the unequilibrated state by a charge transfer reaction for all complexes studied; the yield of the reaction increases when DG increases. The ps charge transfer process reported before [22–24] was also observed. By extending our measurements to daunomycin (DM) complexed with (GMP and AMP) mononucleotides we observed the same femtosecond charge transfer for the DM complexes. The results show that this behavior of deactivation of excited state may be a general phenomenon: the ÔdirectÕ ultrafast charge transfer as well as the ÔindirectÕ charge transfer are two contributing channels to the large fluorescence quenching in the Ap and DM complexes. As such, measurements of quantum yield alone are not sufficient to map out behavior of rates in complexes of biological structures.
2. Experimental 2-Aminopurine (Ap), daunomycin (DM), guanosine monophosphate (GMP), adenine monophosphate (AMP), and L-tryptophan (Trp) were purchased from Sigma. DNA dimers, Ap–A and Ap–G; trimers, G–Ap–G, A–Ap–G, and Ap–Z–A; and tetramer Z–Ap–Z–A (all from 5 0 to 3 0 direction containing Ap with Z denoting 7-deaza guanine), were synthesized at Caltech Oligo Facility. Ap and Ap-containing oligos were used in about 1 mM concentrations. Ap in saturated Trp solution was obtained by soaking Trp in Ap
159
solution, and the Ap/GMP and Ap/AMP mixture solutions were prepared by dissolving appropriate amount of (300 mM) GMP and AMP powder in Ap solution. The experimental setups for both transient absorption and fluorescence up-conversion have been published elsewhere [22,23]. Briefly, a pump pulse at 325 nm was used for excitation. Then a probe pulse at 800 nm was used to mix the Ap fluorescence signal (380 nm) in a BBO crystal. The up-conversion was monitored at the wavelength 257 nm. For DM complexes, a pump pulse at 470–540 nm was used and the fluorescence signal was probed at 570–630 nm. For the transient absorption experiments, a probe pulse, from 350 to 700 nm, was used to detect the absorbance after the excitation by the pump pulse at 325 nm. All the transient absorption data shown in this work were probed at 600 nm, unless specified otherwise. The pulse width of both the pump and probe beams is 100 fs. The polarizations of the pump and probe pulses were set at the magic angle (54.7°) to avoid complications from the effect of orientation on decay behavior. Care was taken to avoid artifact generated by high power intensities, and all measurements were carried out in a 5 mm quartz cell with the sample being constantly stirred during data collection.
3. Results and discussion As shown in Fig. 1, the excited state of free Ap decays on the same nanosecond time scale for both transient absorption and upconversion signals. Such long lifetime is consistent with observations made earlier [3,18]. In a saturated solution of tryptophan (Trp, 60 mM), however, Ap has a totally different decay dynamics in transient absorption and in upconversion (Fig. 1). Ap forms complexes as evidenced from concentration dependent titration with Trp and also the chemical shift perturbation of Ap (data not shown). The upconversion features a 2 ps (12%) decay component, while transient absorption has an 11 ps (32%) decay (Table 1). The 2 ps component in upconversion describes the charge transfer between Ap and Trp [30], but, if so, why it is different from the 11 ps component in transient absorption? To confirm our observations on the Ap/Trp complex and further investigate how the process depends on the nature of the interacting partner in DG-controlled structures, we performed a series of measurements on Apcontaining covalently bonded and non-covalently bonded complexes. We measured both upconversion and transient absorption decay profiles for all of these constructs. In Fig. 2, we show the transients for Z–Ap–Z–A, G–Ap–G, and Ap–A as representatives, taken at short times; some transients at longer times are presented and others not shown. From the data in Table 1, the general trends are mainly two: the rates in-
160
C. Wan et al. / Chemical Physics Letters 412 (2005) 158–163
Normalized Fluorescence
a
1.0
A
0.8 B
0.6
A. Free Ap B. Ap/Trp
0.4 0.2 0.0 0
b
20
40 Time (ps)
60
80
1.0 A
Absorbance, ∆A
0.8 0.6 B
A. Free Ap B. Ap/Trp
0.4 0.2 0.0 0
20 40 Time (ps)
60
80
Fig. 1. (a) Upconversion decay of free Ap and Ap in saturated Trp (60 mM). The samples were excited at 325 nm, the emission monitored at 380 nm. (b) Transient absorption decays of Ap and Ap/Trp solution. The absorption was monitored at 600 nm. Table 1 Ultrafast decay time constants and their normalized pre-exponential factors for Ap complexes Ap complexes
Ultrafast decay parameters Fluorescence upconversion
Transient absorption
Loss of initial fluorescence (%)
Covalent Z–Ap–Z–A Ap–Z–A G–Ap–G Ap–G A–Ap–G Ap–A
0.7 ps (78%) 1 ps (56%) 6 ps (75%) 14 ps (51%) 15 ps (54%) 80 ps (49%)
3.6 ps (79%) 6 ps (67%) 12 ps (78%) 19 ps (54%) 27 ps (64%) 85 ps (42%)
78 78 69 49 61 50
Non-covalent Ap/Trp Ap/GMP Free Ap
2 ps (12%) 6 ps (34%) No decay
11 ps (32%) 19 ps (50%) No decay
77 66 0
Because the rise in transient absorption is 6 200 fs, we take k 1 1 to be 200 fs (see text); k 1 is on the ns scale, as discussed in text. The f deduced k 1 2 from our analyses gives values in the range of 60–200 fs. The time constants given under the fluorescence upconversion column represent the rate constants for k 1 CT , while those under transient absorption represent k 1 CR .
crease with increasing DG (Z > G > A) [22,31,32], and for larger DG the discrepancy between the apparent transient absorption and upconversion is the largest, becoming insignificant for the smallest DG case studied. We have also measured the initial fluorescence intensity normalized to the same absorbance at pump wavelength (325 nm) for these complexes. Fig. 3a shows the fluorescence decay profiles for these Apcontaining covalent constructs for a short time window. The initial fluorescence strongly depends on the identity and number of nucleotides adjacent to Ap in the DNA chain; see Table 1. The decrease again follows the general trend of Z > G > A, as neighboring nucleobases; sandwiching Ap by two nucleotides is more efficient. We also observed significant decrease of the initial fluorescence intensity with increasing concentrations of Trp, GMP, or AMP in Ap solutions (data not shown). To investigate whether this behavior is a common phenomenon for other molecules, we extended the measurements to daunomycin (DM) complexes with GMP and AMP. Fig. 3b shows the normalized fluorescence transients of the free DM and DM/GMP complex. The free DM has a flat trace after the excitation, as expected in this time window, and the DM/GMP complex shows a decay of 3 ps due to charge transfer [33]. The inset shows the percentage loss of the initial fluorescence intensity as a function of the excitation wavelength. Again the initial fluorescence loss accounts for 60% of the population in the wavelength range 490–540 nm. For the DM/ AMP complex (data not shown), it is only 10% due to its much slower charge transfer rate (0.02 ps1). From the above results we concluded that the nonradiative state is a CT state, for the following two reasons: (1) there is a correlation between the percentage loss of initial fluorescence intensity with the rate measured by upconversion (Fig. 4) and termed here kCT (see Fig. 5); and (2) the fact that the behavior is robust for two very different systems, complexes of Ap and also DM, and both systems are expected to undergo charge separation only in the complexes, with correlation for the net DG values. In order to quantify the observed trend, we considered the model depicted in Fig. 5. Following the initial photoexcitation of Ap to the S1 state, the initial population of the complexed (ÔstackedÕ) molecules undergo two ultrafast and competing processes – vibrational redistribution/relaxation with rate constant k1, and the direct CT from the hot molecules with rate constant k2. The two processes must compete on the femtosecond time scale, suggesting that the k2-process is for a (mainly) barrierless CT. Following vibrational relaxation (by k1 and k3), the relaxed S1 population undergoes the equilibrated CT reaction, with rate constant kCT (Ôindirect processÕ). Recombination occurs with the rate constant kCR. In other words, the CT
C. Wan et al. / Chemical Physics Letters 412 (2005) 158–163
a
b Transient Absorption
Upconversion
Z-Ap-Z-A
Z-Ap-Z-A
0
5
10
15
20
0
5
20
40
60
80
50
100
15
20
0 10 20 30 40 50 60 70 80
Ap-A
Ap-A
0
10
G-Ap-G
G-Ap-G
0
161
150
200
0
50
100
150
200
Time (ps) Fig. 2. Representative fluorescence upconversion (a) and transient absorption (b) decays measured for Ap-containing molecules, Z–Ap–Z–A tetramer, G–Ap–G trimer, and Ap–A dimer. Traces are shown for different time windows, depending on the rates. All decay profiles were fitted to a biexponential function except for the Z–Ap–Z–A tetramer and Ap–Z–A trimer (three-exponential function).
state manifold is reached through different pathways, and the ratio k2/k1 will determine the efficiency of the k2-process responsible for the initial loss of fluorescence on the femtosecond time scale. In Fig. 4 we plot the ratio of k2/k1 as a function of the CT rate constant kCT. We expect that the rate k1 of vibrational relaxation/redistribution to be basically constant for different Ap complexes, and thus the ratio of k2/k1 should reflect the behavior of k2 as CT rate increases. Higher rate of k2 (or k2/k1 ratio) leads to more population going into the direct CT channel, and vice versa. The question then is why do the higher k2 values ÔcorrelateÕ with the CT rate constants, kCT? In principle they are two independent processes. This can be understood within the frame work of electron transfer theory
(see Fig. 5). As DG increases, the effective barrier decreases, entirely consistent with increasing the rates for kCT and k2-processes, and with the increasing initial femtosecond drop of fluorescence. The time constants are compiled in Table 1. The upconversion experiments probe only the excited state which decays radiatively, while transient absorption can be sensitive not only to the initially-excited state but also to other non-radiative state(s). The different rates resolved in upconversion and transient absorption are accordingly the result of probing either the initial state (upconversion) and/or the CT state. But the rise of the CT state by the k2-process should be evident when probing the CT population. We have searched for the formation (rise) time of the CT state in the wavelength
162
a 100
1.0 A. Free Ap B. Ap-A C. Ap-G D. A-Ap-G E. G-Ap-G F. Ap-Z-A G. Z-Ap-Z-A
0.8
0.6
A
Loss of Initial Fluorescence (%)
Normalized Fluorescence Intensity
a
C. Wan et al. / Chemical Physics Letters 412 (2005) 158–163
B C D
0.4
E
0.2 F
80
60
40
20
0 G
0.0 –2
–1
0
1
2
3
0.0
4
b A. Free DM B. DM/GMP
1.2
1.6
4
A
0.8
3
1.0 0.8
0.6
0.4
2 k 2/k 1
Fraction loss
Normalized Fluoresence Intensity
1.0
0.8
Charge Transfer Rate (ps-1)
Time (ps) b
0.4
0.6 0.4
1
0.2 0.0
0.2
480
500 520 Wavelength (nm)
B
0.0 –10
0
540
–5
0
5
10
15
20
0.0 25
Time (ps) Fig. 3. (a) Fluorescence upconversion decays measured for the short time window of up to 4 ps for free Ap and Ap-containing DNA strands. The transients are normalized to their absorbances at the pump wavelength 325 nm. The initial fluorescence of free Ap is normalized to unity, and the intensities were obtained after deconvolution with the pulse width. (b) Fluorescence decay profiles for daunomycin (DM) and DM/GMP complex normalized to the absorbance at the excitation wavelength (515 nm). Inset. Fraction loss of initial fluorescence intensity of DM/GMP complex relative to free DM as function of excitation wavelength.
range 370 nm to 650 nm in the transient absorption experiments, and could not resolve a rise longer than 200 fs. This finding suggests that k 1 2 is 6200 fs, consistent with an efficient vibrational relaxation/redistribution (k1) in these large systems. We should note that the nanosecond component present in all these transients represent the Ap-like molecules, free or unstacked [34,35]. With the above picture in mind, we can obtain the recombination rate constants kCR, which for essentially all cases are smaller than those of the forward rate constants of CT,
0.4
0.8
1.2
1.6
Charge Transfer Rate (ps-1) Fig. 4. (a) Fraction of loss of the initial fluorescence intensity obtained from the decay profiles (Fig. 3) and plotted against the charge transfer rate kCT measured in upconversion. Data for Ap/Trp and Ap/GMP solutions are also included. The behavior represents the trend for the increase in the population yield of the direct channel (k2/(k1 + k2)). (b) Ratio of fluorescence intensities (Fig. 3) representing the loss of initial population relative to that undergoing the indirect charge transfer (k2/k1). The solid curves are for visualization purposes only.
and certainly those of the direct processes. From the transient absorption and the upconversion measurements we can estimate the absorption cross section of the CT state to be about half of that of the excited state at 600 nm. In summary, by studying the ultrafast dynamics of Ap-containing constructs and DM complexes with Trp, Z, G, and A bases, we demonstrate here that direct (unequilibrated), nearly barrierless crossing to the charge transfer state can be a major channel that competes with vibrational relaxation and indirect (equilibrated) charge separation. The presence of this pathway is facilitated by the strong interaction between the fluorophores and quenchers in the ÔstackedÕ structures of complexes. The consequence of these
C. Wan et al. / Chemical Physics Letters 412 (2005) 158–163
163
References
a k2
S1 k1
k3
CT
kCT hν
kf
kCR
S0
b S1
CT (small ∆G)
CT (large ∆G)
Fig. 5. (a) Schematic illustrating the dynamic model for excited Ap radiative and non-radiative (CT) pathways. The S0, S1, and CT represent the ground, electronically excited, and charge transfer state, respectively. The rate constant for vibrational relaxation/redistribution (k1 and k3), direct CT (k2), indirect CT (kCT), and charge recombination (kCR) are depicted. The kf represents the emission rate constant of excited state. (b) Schematics of the free energy curves of the S1 and CT states along the solvent coordinate, illustrating the influence of increasing DG on the decrease of effective barrier height and increase of the rates of CT at the two indicated energies.
Ôdark channelsÕ is important to all inferences deducing electron transfer rates from quantum yield measurements, and also to theory of electron transfer, which normally addresses the equilibrated state. In a separate Letter, we will report the significance of these new findings of Ap to probing the interfacial recognition dynamics in RNA/peptide complex structures. Acknowledgments This work was supported by a grant from the National Science Foundation. We thank Dr. Spencer Baskin for his critical comments.
[1] S.F. Mason, J. Chem. Soc. (Lond.) (1954) 2071. [2] A.N. Fletcher, J. Mol. Spectrosc. (1967) 221. [3] D.C. Ward, E. Reich, L. Stryer, J. Biol. Chem. 244 (1969) 1228. [4] J. Smagowicz, K.L. Wierzchowski, J. Lumin. 8 (1974) 210. [5] C. Santhosh, P.C. Mishra, Spectrochim. Acta 47A (1991) 1685. [6] J. Drobnik, L. Augenstein, Photochem. Photobiol. 5 (1966) 83. [7] E.L. Rachofsky, R. Osman, J.B. Ross, Biochemistry 40 (2001) 946. [8] D.P. Millar, Curr. Opin. Struct. Biol. 6 (1996) 322. [9] T.M. Nordlund, S. Andersson, L. Nilsson, R. Rigler, A. Graslund, L.W. McLaughlin, Biochemistry 28 (1989) 9095. [10] C.R. Guest, R.A. Hochstrasser, L.C. Sowers, D.P. Millar, Biochemistry 30 (1991) 3271. [11] O.F.A. Larsen, I.H. van Stokkum, B. Gobets, R. van Grondelle, H. van Amerongen, Biophys. J. 81 (2001) 1115. [12] K.B. Hall, D.J. Williams, RNA 10 (2004) 34. [13] S.K. Pal, L. Zhao, T. Xia, A.H. Zewail, Proc. Natl. Acad. Sci. USA 100 (2003) 13746. [14] R. Baliga, E.E. Baird, D.M. Herman, C. Melander, P.B. Dervan, D.M. Crothers, Biochemistry 40 (2001) 3. [15] K.A. Lacourciere, J.T. Stivers, J.P. Marino, Biochemistry 39 (2000) 5630. [16] T. Xia, A. Frankel, T.T. Takahashi, J. Ren, R.W. Roberts, Nat. Struct. Biol. 10 (2003) 812. [17] A. Broo, A. Holmen, Chem. Phys. 211 (1996) 147. [18] A. Holmen, B. Norden, B. Albinsson, J. Am. Chem. Soc. 119 (1997) 3114. [19] E.L. Rachofsky, J.B.A. Ross, M. Krauss, R. Osman, J. Phys. Chem. 105 (2001) 190. [20] J.M. Jean, K.B. Hall, J. Phys. Chem. A 104 (2000) 1930. [21] A. Broo, J. Phys. Chem. 102 (1998) 526. [22] C. Wan, T. Fiebig, O. Schiemann, J.K. Barton, A.H. Zewail, Proc. Natl. Acad. Sci. USA 97 (2000) 14052. [23] T. Fiebig, C. Wan, A.H. Zewail, Chem. Phys. Chem. 3 (2002) 781. [24] M.A. OÕNeill, H.C. Becker, C. Wan, J.K. Barton, A.H. Zewail, Angew. Chem. Int. Ed. Engl. 42 (2003) 5896. [25] S.O. Kelley, J.K. Barton, Science 283 (1999) 375. [26] M.A. OÕNeill, J.K. Barton, J. Am. Chem. Soc. 124 (2002) 13053. [27] O.F.A. Larsen, I.H.M.V. Stokkum, M.-L. Groot, J.T.M. Kennis, R.v. Grondelle, H.v. Amerongen, Chem. Phys. Lett. 371 (2003) 157. [28] O.F.A. Larsen, I.H.M.V. Stokkum, F.L.d. Weerd, M. Vengris, C.T. Aravindakumar, R.v. Grondelle, N.E. Geacintov, H.v. Amerongen, Phys. Chem. Chem. Phys. 6 (2004) 154. [29] O.J.G. Somsen, V.A. Hoek, V.H. Amerongen, Chem. Phys. Lett. 402 (2005) 61. [30] T. Xia, H.C. Becker, C. Wan, A. Frankel, R.W. Roberts, A.H. Zewail, Proc. Natl. Acad. Sci. USA 100 (2003) 8119. [31] S. Steenken, S.V. Jovanovic, J. Am. Chem. Soc. 119 (1997) 617. [32] S.O. Kelley, J.K. Barton, Chem. Biol. 5 (1998) 413. [33] X. Qu, C. Wan, H.C. Becker, D. Zhong, A.H. Zewail, Proc. Natl. Acad. Sci. USA 98 (2001) 14212. [34] P.A.W.v.d. Berg, K.A. Feenstra, A.E. Mark, H.J.C. Berendsen, A.J.W.G. Visser, J. Phys. Chem. B 106 (2002) 8858. [35] J.M. Jean, K.B. Hall, Biochemistry 43 (2004) 10277.
Ultrafast unequilibrated charge transfer: A new channel in the quenching of fluorescent biological probes Chaozhi Wan, Tianbing Xia, Hans-Christian Becker 1, Ahmed H. Zewail
*
Laboratory for Molecular Sciences, Arthur Amos Noyes Laboratory of Chemical Physics, Division of Chemistry and Chemical Engineering, California Institute of Technology, Mail Code 127-72, Pasadena, CA 91125, USA Received 8 June 2005; in final form 20 June 2005 Available online 19 July 2005
Abstract The dynamics of two biological fluorescent probes, 2-aminopurine (Ap) and daunomycin, were studied using both femtosecond transient absorption and fluorescence upconversion techniques. Various Ap-containing structures were investigated in solution: free Ap, non-covalently bonded (with guanine, adenine, and tryptophan) and covalently bonded in DNA constructs (with guanine, 7deazaguanine, and adenine). The distinct difference of transient absorption and fluorescence dynamics on the ultrafast time scale, and their dependence on free energy change (DG), and the abrupt decrease of the initial fluorescence intensity suggest the efficient depopulation by charge transfer from the unequilibrated hot molecules. We provide a model for this possibly general mechanism and obtain the rate constants for charge separation, vibrational relaxation, and charge recombination. Ó 2005 Elsevier B.V. All rights reserved.
1. Introduction As a structural analog of adenine, 2-aminopurine (Ap) can form base pairs with other bases, exhibiting reasonable stability and minimal perturbations to DNA structures. Ap can be excited selectively in the presence of other bases [1], and it has high fluorescence quantum yield (0.6–0.7) [2,3], but sensitive to the environment [3–6] and surrounding structures [7]. Ap has been extensively used as a probe molecule [8] to study DNA and RNA structures and conformational dynamics [9–12], DNA hydration [13], recognition of DNA by polyamides [14] and RNA by peptides [15,16], and enzymatic activity of various DNA-binding proteins. The usefulness of Ap has prompted extensive studies of its fundamental physical properties, such as molecular *
Corresponding author. Fax: +1 626 792 8456/405 0454 . E-mail addresses: [email protected], [email protected] (A.H. Zewail). 1 Present address: Department of Physical Chemistry, Uppsala University, Box 579, SE-751 23 Uppsala, Sweden. 0009-2614/$ - see front matter Ó 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.cplett.2005.06.101
geometry, tautomerism and dipole moments [17], directions and strength of electronic transition moments [18], electronic structure [19], and state ordering and energy gaps [20]. The goal was to understand the basis for the very different luminescence properties between the two isomers, adenine and 2-aminopurine [21]. Of particular usefulness is the use of Ap as a probe for charge transfer (CT) between excited state Ap and nucleobases in various DNA assemblies, in both timeresolved [22–24] and steady-state measurements [25, 26]. Charge transfer reactions lead to shortening of the lifetime of excited Ap, from the nanosecond (ns) regime for free Ap to a wide range within the picosecond (ps) regime in DNA, depending on the nature and distance between the donor and acceptor [22]. The direction of charge transfer (electron transfer vs. hole transfer) depends on the identity of the interaction partner with Ap [23]. This non-radiative decay pathway results in a large reduction of the fluorescence quantum yield. While the time-resolved methods directly measure the decay rates, steady-state methods infer the change in rates
C. Wan et al. / Chemical Physics Letters 412 (2005) 158–163
from measurement of overall fluorescence quantum yield and assuming that the decrease is caused by the charge transfer processes. However, this inference is valid provided there are no other non-radiative channels. Recently, Larsen et al. [27,28] proposed the existence of a long-lived non-radiative state for Ap, based on the wavelength dependent anisotropy decays in transient absorption. More recently a non-radiative state was proposed to explain the observed strong fluorescence quenching of Ap dinucleotides, but maintaining that the ps charge transfer still plays an important role in the quenching [29]. In these studies either transient absorption was performed or fluorescence decays were measured by time correlated single photon counting with limited resolution of 50 ps. However, the fluorescence quenching may also occur on much shorter time scales. Also the nature of the non-fluorescent state is unknown. In this work, we have used both fluorescence upconversion and transient absorption spectroscopy with femtosecond time resolution to systematically investigate the dynamics in a series of Ap-containing structures: free Ap, covalently bonded and non-covalently bonded complexes. Attention was paid to the early dynamics on the time scale of 200 fs and longer. On this ultrafast time scale, our analyses suggest the depopulation of the unequilibrated state by a charge transfer reaction for all complexes studied; the yield of the reaction increases when DG increases. The ps charge transfer process reported before [22–24] was also observed. By extending our measurements to daunomycin (DM) complexed with (GMP and AMP) mononucleotides we observed the same femtosecond charge transfer for the DM complexes. The results show that this behavior of deactivation of excited state may be a general phenomenon: the ÔdirectÕ ultrafast charge transfer as well as the ÔindirectÕ charge transfer are two contributing channels to the large fluorescence quenching in the Ap and DM complexes. As such, measurements of quantum yield alone are not sufficient to map out behavior of rates in complexes of biological structures.
2. Experimental 2-Aminopurine (Ap), daunomycin (DM), guanosine monophosphate (GMP), adenine monophosphate (AMP), and L-tryptophan (Trp) were purchased from Sigma. DNA dimers, Ap–A and Ap–G; trimers, G–Ap–G, A–Ap–G, and Ap–Z–A; and tetramer Z–Ap–Z–A (all from 5 0 to 3 0 direction containing Ap with Z denoting 7-deaza guanine), were synthesized at Caltech Oligo Facility. Ap and Ap-containing oligos were used in about 1 mM concentrations. Ap in saturated Trp solution was obtained by soaking Trp in Ap
159
solution, and the Ap/GMP and Ap/AMP mixture solutions were prepared by dissolving appropriate amount of (300 mM) GMP and AMP powder in Ap solution. The experimental setups for both transient absorption and fluorescence up-conversion have been published elsewhere [22,23]. Briefly, a pump pulse at 325 nm was used for excitation. Then a probe pulse at 800 nm was used to mix the Ap fluorescence signal (380 nm) in a BBO crystal. The up-conversion was monitored at the wavelength 257 nm. For DM complexes, a pump pulse at 470–540 nm was used and the fluorescence signal was probed at 570–630 nm. For the transient absorption experiments, a probe pulse, from 350 to 700 nm, was used to detect the absorbance after the excitation by the pump pulse at 325 nm. All the transient absorption data shown in this work were probed at 600 nm, unless specified otherwise. The pulse width of both the pump and probe beams is 100 fs. The polarizations of the pump and probe pulses were set at the magic angle (54.7°) to avoid complications from the effect of orientation on decay behavior. Care was taken to avoid artifact generated by high power intensities, and all measurements were carried out in a 5 mm quartz cell with the sample being constantly stirred during data collection.
3. Results and discussion As shown in Fig. 1, the excited state of free Ap decays on the same nanosecond time scale for both transient absorption and upconversion signals. Such long lifetime is consistent with observations made earlier [3,18]. In a saturated solution of tryptophan (Trp, 60 mM), however, Ap has a totally different decay dynamics in transient absorption and in upconversion (Fig. 1). Ap forms complexes as evidenced from concentration dependent titration with Trp and also the chemical shift perturbation of Ap (data not shown). The upconversion features a 2 ps (12%) decay component, while transient absorption has an 11 ps (32%) decay (Table 1). The 2 ps component in upconversion describes the charge transfer between Ap and Trp [30], but, if so, why it is different from the 11 ps component in transient absorption? To confirm our observations on the Ap/Trp complex and further investigate how the process depends on the nature of the interacting partner in DG-controlled structures, we performed a series of measurements on Apcontaining covalently bonded and non-covalently bonded complexes. We measured both upconversion and transient absorption decay profiles for all of these constructs. In Fig. 2, we show the transients for Z–Ap–Z–A, G–Ap–G, and Ap–A as representatives, taken at short times; some transients at longer times are presented and others not shown. From the data in Table 1, the general trends are mainly two: the rates in-
160
C. Wan et al. / Chemical Physics Letters 412 (2005) 158–163
Normalized Fluorescence
a
1.0
A
0.8 B
0.6
A. Free Ap B. Ap/Trp
0.4 0.2 0.0 0
b
20
40 Time (ps)
60
80
1.0 A
Absorbance, ∆A
0.8 0.6 B
A. Free Ap B. Ap/Trp
0.4 0.2 0.0 0
20 40 Time (ps)
60
80
Fig. 1. (a) Upconversion decay of free Ap and Ap in saturated Trp (60 mM). The samples were excited at 325 nm, the emission monitored at 380 nm. (b) Transient absorption decays of Ap and Ap/Trp solution. The absorption was monitored at 600 nm. Table 1 Ultrafast decay time constants and their normalized pre-exponential factors for Ap complexes Ap complexes
Ultrafast decay parameters Fluorescence upconversion
Transient absorption
Loss of initial fluorescence (%)
Covalent Z–Ap–Z–A Ap–Z–A G–Ap–G Ap–G A–Ap–G Ap–A
0.7 ps (78%) 1 ps (56%) 6 ps (75%) 14 ps (51%) 15 ps (54%) 80 ps (49%)
3.6 ps (79%) 6 ps (67%) 12 ps (78%) 19 ps (54%) 27 ps (64%) 85 ps (42%)
78 78 69 49 61 50
Non-covalent Ap/Trp Ap/GMP Free Ap
2 ps (12%) 6 ps (34%) No decay
11 ps (32%) 19 ps (50%) No decay
77 66 0
Because the rise in transient absorption is 6 200 fs, we take k 1 1 to be 200 fs (see text); k 1 is on the ns scale, as discussed in text. The f deduced k 1 2 from our analyses gives values in the range of 60–200 fs. The time constants given under the fluorescence upconversion column represent the rate constants for k 1 CT , while those under transient absorption represent k 1 CR .
crease with increasing DG (Z > G > A) [22,31,32], and for larger DG the discrepancy between the apparent transient absorption and upconversion is the largest, becoming insignificant for the smallest DG case studied. We have also measured the initial fluorescence intensity normalized to the same absorbance at pump wavelength (325 nm) for these complexes. Fig. 3a shows the fluorescence decay profiles for these Apcontaining covalent constructs for a short time window. The initial fluorescence strongly depends on the identity and number of nucleotides adjacent to Ap in the DNA chain; see Table 1. The decrease again follows the general trend of Z > G > A, as neighboring nucleobases; sandwiching Ap by two nucleotides is more efficient. We also observed significant decrease of the initial fluorescence intensity with increasing concentrations of Trp, GMP, or AMP in Ap solutions (data not shown). To investigate whether this behavior is a common phenomenon for other molecules, we extended the measurements to daunomycin (DM) complexes with GMP and AMP. Fig. 3b shows the normalized fluorescence transients of the free DM and DM/GMP complex. The free DM has a flat trace after the excitation, as expected in this time window, and the DM/GMP complex shows a decay of 3 ps due to charge transfer [33]. The inset shows the percentage loss of the initial fluorescence intensity as a function of the excitation wavelength. Again the initial fluorescence loss accounts for 60% of the population in the wavelength range 490–540 nm. For the DM/ AMP complex (data not shown), it is only 10% due to its much slower charge transfer rate (0.02 ps1). From the above results we concluded that the nonradiative state is a CT state, for the following two reasons: (1) there is a correlation between the percentage loss of initial fluorescence intensity with the rate measured by upconversion (Fig. 4) and termed here kCT (see Fig. 5); and (2) the fact that the behavior is robust for two very different systems, complexes of Ap and also DM, and both systems are expected to undergo charge separation only in the complexes, with correlation for the net DG values. In order to quantify the observed trend, we considered the model depicted in Fig. 5. Following the initial photoexcitation of Ap to the S1 state, the initial population of the complexed (ÔstackedÕ) molecules undergo two ultrafast and competing processes – vibrational redistribution/relaxation with rate constant k1, and the direct CT from the hot molecules with rate constant k2. The two processes must compete on the femtosecond time scale, suggesting that the k2-process is for a (mainly) barrierless CT. Following vibrational relaxation (by k1 and k3), the relaxed S1 population undergoes the equilibrated CT reaction, with rate constant kCT (Ôindirect processÕ). Recombination occurs with the rate constant kCR. In other words, the CT
C. Wan et al. / Chemical Physics Letters 412 (2005) 158–163
a
b Transient Absorption
Upconversion
Z-Ap-Z-A
Z-Ap-Z-A
0
5
10
15
20
0
5
20
40
60
80
50
100
15
20
0 10 20 30 40 50 60 70 80
Ap-A
Ap-A
0
10
G-Ap-G
G-Ap-G
0
161
150
200
0
50
100
150
200
Time (ps) Fig. 2. Representative fluorescence upconversion (a) and transient absorption (b) decays measured for Ap-containing molecules, Z–Ap–Z–A tetramer, G–Ap–G trimer, and Ap–A dimer. Traces are shown for different time windows, depending on the rates. All decay profiles were fitted to a biexponential function except for the Z–Ap–Z–A tetramer and Ap–Z–A trimer (three-exponential function).
state manifold is reached through different pathways, and the ratio k2/k1 will determine the efficiency of the k2-process responsible for the initial loss of fluorescence on the femtosecond time scale. In Fig. 4 we plot the ratio of k2/k1 as a function of the CT rate constant kCT. We expect that the rate k1 of vibrational relaxation/redistribution to be basically constant for different Ap complexes, and thus the ratio of k2/k1 should reflect the behavior of k2 as CT rate increases. Higher rate of k2 (or k2/k1 ratio) leads to more population going into the direct CT channel, and vice versa. The question then is why do the higher k2 values ÔcorrelateÕ with the CT rate constants, kCT? In principle they are two independent processes. This can be understood within the frame work of electron transfer theory
(see Fig. 5). As DG increases, the effective barrier decreases, entirely consistent with increasing the rates for kCT and k2-processes, and with the increasing initial femtosecond drop of fluorescence. The time constants are compiled in Table 1. The upconversion experiments probe only the excited state which decays radiatively, while transient absorption can be sensitive not only to the initially-excited state but also to other non-radiative state(s). The different rates resolved in upconversion and transient absorption are accordingly the result of probing either the initial state (upconversion) and/or the CT state. But the rise of the CT state by the k2-process should be evident when probing the CT population. We have searched for the formation (rise) time of the CT state in the wavelength
162
a 100
1.0 A. Free Ap B. Ap-A C. Ap-G D. A-Ap-G E. G-Ap-G F. Ap-Z-A G. Z-Ap-Z-A
0.8
0.6
A
Loss of Initial Fluorescence (%)
Normalized Fluorescence Intensity
a
C. Wan et al. / Chemical Physics Letters 412 (2005) 158–163
B C D
0.4
E
0.2 F
80
60
40
20
0 G
0.0 –2
–1
0
1
2
3
0.0
4
b A. Free DM B. DM/GMP
1.2
1.6
4
A
0.8
3
1.0 0.8
0.6
0.4
2 k 2/k 1
Fraction loss
Normalized Fluoresence Intensity
1.0
0.8
Charge Transfer Rate (ps-1)
Time (ps) b
0.4
0.6 0.4
1
0.2 0.0
0.2
480
500 520 Wavelength (nm)
B
0.0 –10
0
540
–5
0
5
10
15
20
0.0 25
Time (ps) Fig. 3. (a) Fluorescence upconversion decays measured for the short time window of up to 4 ps for free Ap and Ap-containing DNA strands. The transients are normalized to their absorbances at the pump wavelength 325 nm. The initial fluorescence of free Ap is normalized to unity, and the intensities were obtained after deconvolution with the pulse width. (b) Fluorescence decay profiles for daunomycin (DM) and DM/GMP complex normalized to the absorbance at the excitation wavelength (515 nm). Inset. Fraction loss of initial fluorescence intensity of DM/GMP complex relative to free DM as function of excitation wavelength.
range 370 nm to 650 nm in the transient absorption experiments, and could not resolve a rise longer than 200 fs. This finding suggests that k 1 2 is 6200 fs, consistent with an efficient vibrational relaxation/redistribution (k1) in these large systems. We should note that the nanosecond component present in all these transients represent the Ap-like molecules, free or unstacked [34,35]. With the above picture in mind, we can obtain the recombination rate constants kCR, which for essentially all cases are smaller than those of the forward rate constants of CT,
0.4
0.8
1.2
1.6
Charge Transfer Rate (ps-1) Fig. 4. (a) Fraction of loss of the initial fluorescence intensity obtained from the decay profiles (Fig. 3) and plotted against the charge transfer rate kCT measured in upconversion. Data for Ap/Trp and Ap/GMP solutions are also included. The behavior represents the trend for the increase in the population yield of the direct channel (k2/(k1 + k2)). (b) Ratio of fluorescence intensities (Fig. 3) representing the loss of initial population relative to that undergoing the indirect charge transfer (k2/k1). The solid curves are for visualization purposes only.
and certainly those of the direct processes. From the transient absorption and the upconversion measurements we can estimate the absorption cross section of the CT state to be about half of that of the excited state at 600 nm. In summary, by studying the ultrafast dynamics of Ap-containing constructs and DM complexes with Trp, Z, G, and A bases, we demonstrate here that direct (unequilibrated), nearly barrierless crossing to the charge transfer state can be a major channel that competes with vibrational relaxation and indirect (equilibrated) charge separation. The presence of this pathway is facilitated by the strong interaction between the fluorophores and quenchers in the ÔstackedÕ structures of complexes. The consequence of these
C. Wan et al. / Chemical Physics Letters 412 (2005) 158–163
163
References
a k2
S1 k1
k3
CT
kCT hν
kf
kCR
S0
b S1
CT (small ∆G)
CT (large ∆G)
Fig. 5. (a) Schematic illustrating the dynamic model for excited Ap radiative and non-radiative (CT) pathways. The S0, S1, and CT represent the ground, electronically excited, and charge transfer state, respectively. The rate constant for vibrational relaxation/redistribution (k1 and k3), direct CT (k2), indirect CT (kCT), and charge recombination (kCR) are depicted. The kf represents the emission rate constant of excited state. (b) Schematics of the free energy curves of the S1 and CT states along the solvent coordinate, illustrating the influence of increasing DG on the decrease of effective barrier height and increase of the rates of CT at the two indicated energies.
Ôdark channelsÕ is important to all inferences deducing electron transfer rates from quantum yield measurements, and also to theory of electron transfer, which normally addresses the equilibrated state. In a separate Letter, we will report the significance of these new findings of Ap to probing the interfacial recognition dynamics in RNA/peptide complex structures. Acknowledgments This work was supported by a grant from the National Science Foundation. We thank Dr. Spencer Baskin for his critical comments.
[1] S.F. Mason, J. Chem. Soc. (Lond.) (1954) 2071. [2] A.N. Fletcher, J. Mol. Spectrosc. (1967) 221. [3] D.C. Ward, E. Reich, L. Stryer, J. Biol. Chem. 244 (1969) 1228. [4] J. Smagowicz, K.L. Wierzchowski, J. Lumin. 8 (1974) 210. [5] C. Santhosh, P.C. Mishra, Spectrochim. Acta 47A (1991) 1685. [6] J. Drobnik, L. Augenstein, Photochem. Photobiol. 5 (1966) 83. [7] E.L. Rachofsky, R. Osman, J.B. Ross, Biochemistry 40 (2001) 946. [8] D.P. Millar, Curr. Opin. Struct. Biol. 6 (1996) 322. [9] T.M. Nordlund, S. Andersson, L. Nilsson, R. Rigler, A. Graslund, L.W. McLaughlin, Biochemistry 28 (1989) 9095. [10] C.R. Guest, R.A. Hochstrasser, L.C. Sowers, D.P. Millar, Biochemistry 30 (1991) 3271. [11] O.F.A. Larsen, I.H. van Stokkum, B. Gobets, R. van Grondelle, H. van Amerongen, Biophys. J. 81 (2001) 1115. [12] K.B. Hall, D.J. Williams, RNA 10 (2004) 34. [13] S.K. Pal, L. Zhao, T. Xia, A.H. Zewail, Proc. Natl. Acad. Sci. USA 100 (2003) 13746. [14] R. Baliga, E.E. Baird, D.M. Herman, C. Melander, P.B. Dervan, D.M. Crothers, Biochemistry 40 (2001) 3. [15] K.A. Lacourciere, J.T. Stivers, J.P. Marino, Biochemistry 39 (2000) 5630. [16] T. Xia, A. Frankel, T.T. Takahashi, J. Ren, R.W. Roberts, Nat. Struct. Biol. 10 (2003) 812. [17] A. Broo, A. Holmen, Chem. Phys. 211 (1996) 147. [18] A. Holmen, B. Norden, B. Albinsson, J. Am. Chem. Soc. 119 (1997) 3114. [19] E.L. Rachofsky, J.B.A. Ross, M. Krauss, R. Osman, J. Phys. Chem. 105 (2001) 190. [20] J.M. Jean, K.B. Hall, J. Phys. Chem. A 104 (2000) 1930. [21] A. Broo, J. Phys. Chem. 102 (1998) 526. [22] C. Wan, T. Fiebig, O. Schiemann, J.K. Barton, A.H. Zewail, Proc. Natl. Acad. Sci. USA 97 (2000) 14052. [23] T. Fiebig, C. Wan, A.H. Zewail, Chem. Phys. Chem. 3 (2002) 781. [24] M.A. OÕNeill, H.C. Becker, C. Wan, J.K. Barton, A.H. Zewail, Angew. Chem. Int. Ed. Engl. 42 (2003) 5896. [25] S.O. Kelley, J.K. Barton, Science 283 (1999) 375. [26] M.A. OÕNeill, J.K. Barton, J. Am. Chem. Soc. 124 (2002) 13053. [27] O.F.A. Larsen, I.H.M.V. Stokkum, M.-L. Groot, J.T.M. Kennis, R.v. Grondelle, H.v. Amerongen, Chem. Phys. Lett. 371 (2003) 157. [28] O.F.A. Larsen, I.H.M.V. Stokkum, F.L.d. Weerd, M. Vengris, C.T. Aravindakumar, R.v. Grondelle, N.E. Geacintov, H.v. Amerongen, Phys. Chem. Chem. Phys. 6 (2004) 154. [29] O.J.G. Somsen, V.A. Hoek, V.H. Amerongen, Chem. Phys. Lett. 402 (2005) 61. [30] T. Xia, H.C. Becker, C. Wan, A. Frankel, R.W. Roberts, A.H. Zewail, Proc. Natl. Acad. Sci. USA 100 (2003) 8119. [31] S. Steenken, S.V. Jovanovic, J. Am. Chem. Soc. 119 (1997) 617. [32] S.O. Kelley, J.K. Barton, Chem. Biol. 5 (1998) 413. [33] X. Qu, C. Wan, H.C. Becker, D. Zhong, A.H. Zewail, Proc. Natl. Acad. Sci. USA 98 (2001) 14212. [34] P.A.W.v.d. Berg, K.A. Feenstra, A.E. Mark, H.J.C. Berendsen, A.J.W.G. Visser, J. Phys. Chem. B 106 (2002) 8858. [35] J.M. Jean, K.B. Hall, Biochemistry 43 (2004) 10277.