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Fis-protein induces rod-like DNA bending
Chemical Physics Letters 500 (2010) 318–322
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Fis-protein induces rod-like DNA bending Chi-Cheng Fu a, Ching-Fong Lin c, Quan-Ze Gao c, Wei-Zen Yang d, Tsong-Shin Lim a, Li-Ling Yang a,e, Chi-Fu Yen f, Wei-Hau Chang f,g,h,i, Hanna S. Yuan d, Sheh-Yi Sheu b,c,⇑,1, Dah-Yen Yang a,⇑,1, Wunshain Fann a,e,⇑,1, a
Institute of Atomic and Molecular Sciences, Academia Sinica, Taipei 106, Taiwan Department of Life Sciences, Institute of Genome Sciences, National Yang-Ming University, Taipei 112, Taiwan Institute of Biomedical Informatics, National Yang-Ming University, Taipei 112, Taiwan d Institute of Molecular Biology, Academia Sinica, Taipei 115, Taiwan e Department of Physics, National Taiwan University, Taipei 106, Taiwan f Institute of Chemistry, Academia Sinica, Taipei 115, Taiwan g Genomic Research Center, Academia Sinica, Taipei 115, Taiwan h Institute of Biotechnology, National Taipei University of Technology, Taipei 106, Taiwan i Institute of Engineering Technology, National Taiwan University of Science and Technology, Taipei 106, Taiwan b c
a r t i c l e
i n f o
Article history: Received 15 June 2010 In final form 11 October 2010 Available online 13 October 2010
a b s t r a c t Fis protein can bend DNA chain with length much shorter than its persistence length. We applied singlemolecule fluorescence resonance energy transfer method to probe these conformational changes. A broad distribution of end-to-end distances correlates well with the molecular dynamics simulation. The flexibility of DNA upon Fis binding is attributed to the breakages of hydrogen bonds between base pairs. DNA kinks at specific sites, instead of continuous bending. The loosening of DNA structures might have biological implications for the functions of Fis-proteins as transcription cofactors. Ó 2010 Elsevier B.V. All rights reserved.
To synthesize RNA from DNA templates requires association of RNA polymerase (RNAP) enzyme to the target genes [1]. Sophisticated regulatory mechanisms control gene transcription which involves coordinated recruitment of multiple transcriptional factors to the promoter regions [2,3]. Typically, the transcription cofactors assemble to form the pre-initiation complexes which bind directly or indirectly to DNA and activate the RNAP. In the growth phase of Escherichia coli, one of the main factors is Fis protein which binds the C-terminal region of the a-subunit of the RNAP and simultaneously bends the upstream of promoters [1,4]. By increasing the affinity of RNAP to rrnB P1 promoter about 2.5-fold, Fis can increase transcription level by 4 to 10-fold in vivo [5]. DNA bending was also observed in supercoiled bacterial chromosomes, which are highly compacted by associating with histone-like proteins, such as Fis, IHF, HU and H-NS [6]. The persistence length (Lp) of a double-stranded DNA (dsDNA) in solution is ca. 50 nm. A DNA molecule with length of 10 nm should be very rigid, and thus is hard to bend [7]. Binding of Fis can induce a significant bending of such short DNA. However, the detailed mechanism is still unclear. Fis proteins recognize specific 15 bp core sequences and induce bending of DNA with an angle ⇑ Corresponding authors. E-mail addresses: [email protected] (S.-Y. Sheu), [email protected]. edu.tw (D.-Y. Yang). 1 These authors contributed equally to this work. Deceased. 0009-2614/$ - see front matter Ó 2010 Elsevier B.V. All rights reserved. doi:10.1016/j.cplett.2010.10.026
ranging from 50° to 90°, which varies with flanking sequences [8]. Conventionally, the bending angles are measured by the gel mobility shift assays. However, these kinds of ensemble measurements cannot differentiate two models proposed to explain protein induced DNA bending phenomena [9]. One is called ‘transient kink’ model, in which protein binds to DNA only in short periods of time. It induces a kink, and then dissociates, which restores DNA to its original length. The transient binding accounts for the fluctuation of DNA conformations. The other is called ‘flexible hinge’ model, in which protein binds to DNA strongly, induces a partial denaturation of DNA, and thus creates flexible complex structures. In this Letter, a single pair fluorescence resonance energy transfer (spFRET) method [10] combined with all-atom molecular dynamics (MD) simulation was used to study the mechanism of Fis protein induced DNA bending. The spFRET technique is applied to study the bending of ‘rod-like’ DNA due to protein interactions at single molecule level. The advantage of this approach is that one can directly obtain the length distribution of DNA instead of its average value. This is of particular relevance to biological systems because most bio-molecules are flexible [11]. In this experiment, the Fis protein with Pro26Ala mutation was prepared as previously described [12]. As shown in Figure 1a, the DA_DNA is designed as a specific 26 bp dsDNA (50 -GATTT TGCAT AAAAA ACAGA CTACA T-30 ) tailed with biotinylated singlestranded DNA (ssDNA, six base, yellow). Both two 50 ends of the duplex are labeled with a donor (Alexa Fluor 546) and an acceptor (Alexa Fluor 647) fluorescent dye molecule respectively. The
C.-C. Fu et al. / Chemical Physics Letters 500 (2010) 318–322
Figure 1. Snapshots of the initial state (a) and the intermediate state at the minimum 50 –50 distance of dsDNA (b) for Fis–DNA complex structure along the MD trajectory.
donor-to-acceptor distance R is 11.6 nm, which includes the length of dsDNA (8.8 nm) and two linkers of dyes (1.4 nm for each). The tail of ssDNA allows dsDNA about 1 nm away from surface to minimize the surface effect. Note that the D_DNA, a DNA labeled with
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donor dye alone, is to the same as DA_DNA except acceptor label. The DNA samples were diluted by Fis binding buffer (with or without 2 lM Fis) at concentrations of 20 pM or 100 nM for the singlemolecule or ensemble measurements, respectively [13]. Because the equilibrium dissociation constant for Fis with DNA is 9.3 1010 M [5], majority of DNA molecules will bind with Fis for all samples. The FRET efficiency EFRET is related to R and the Förster radius R0, satisfying the EFRET = [1 + (R/R0)6]1 relationship [14]. Our time-resolved FRET measurements are based on the measurements of the averaged lifetime of the donor in the presence hsDAi and in the absence hsDi of the acceptor to obtain EFRET = 1(hsDi/hsDAi). As shown in Figure 2a, the lifetime of D_DNA and DA_DNA are 3.8 ns and 3.5 ns which correspond to EFRET = 0.079. Using the total length R as the input, R0 of this donor–acceptor pair was determined to be 7.6 nm, consistent with the value specified by the vendor (Molecular Probes). Note that this is one of the largest R0 available. After incubation with Fis, the measured lifetime decreases to 3.2 ns which corresponded to a change of R to 10 nm. For single molecular spFRET measurements [14], the biotinylated DNA molecules were immobilized on a glass plate by standard biotin–streptavidin interaction in a fluid channel. The distributions of their conformations obtained from measuring many molecules can be compared with the time-dependent conformational fluctuations simulated by MD (see below). The fluorescence intensities of both donor ID and acceptor IA were measured simultaneously by a confocal microscopy equipped with a double port. We can obtain the FRET efficiency in terms of EFRET = IA/(ID + IA). The upper panel of Figure 2b shows that the EFRET histogram of DA_DNA sample is well fitted with two Gaussian distributions. Apparently, the first distribution (blue), centered at zero, can be attributed to DNA molecules containing donors but inactive acceptors. This was confirmed by measuring the D_DNA sample (data not shown). The second distribution (green) is centered at 0.072 with width of 0.045. Its center corresponds to a mean distance of R = 11.64 nm, which agrees with the length of our designed DNA; its width corresponds to the R ranging from 10.64 nm to 13.81 nm, as shown in Figure 3c (green dash line). Notice that even the energy transfer distribution, Figure 2b, is symmetric; the corresponding distance
Figure 2. (a) The ensemble time-resolved FRET results of D_DNA, DA_DNA and DA_DNA + Fis. (b) The EFRET histograms of DA_DNA (upper panel) and DA_DNA + Fis (lower panel) measured by the single molecular method. The inset of (b) shows typical single spFRET molecule time traces.
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Figure 3. Time evolution (upper panel) of 30 -to-50 (a) and 50 -to-50 (b) distance of dsDNA binding with Fis by MD simulation and the corresponding histogram (lower panel). The chains A and B of dsDNA are colored in black and red (a). The 50 -to-50 distances of dsDNA in Fis–DNA complex and dsDNA alone are colored in orange and green (b). (c) The donor–acceptor distance R distributions of DA_DNA (green dash lines) and DA_DNA binding with Fis (red dash line). The orange line is the convolution of green dash line with the distribution function shown in the lower panel of (b). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
distribution, Figure 3c, is asymmetric. This is due to the fact that the average distance, 11.64 nm, is much longer than R0, 7.6 nm. Thus the measurements operated at the tail of the transfer curve. After incubating with Fis, an additional third distribution attributed to the Fis–DNA complex (red) appeared in the lower panel of Figure 2b. The first two distributions (blue and green) are caused by the DA_DNA without Fis binding, as referred to the similar distributions seen in the DA_DNA only measurements. The third distribution has a mean value at 0.12 with width of 0.08. The center energy transfer value corresponds to R = 10.59 nm, which agrees well with the results of ensemble time-resolved measurement. This agreement suggests that the third distribution should represent the conformation distribution of DNA–Fis complexes in solution. The width of this distribution corresponded to the R between 9.58 and 12.91 nm, indicating the higher flexibility of the DNA–Fis complex as shown in Figure 3c (red dash line). Two typical single molecule time traces were shown in the inset of Figure 2b. In the upper panel, the intensity of accepter drops to the background following the photobleach of donor dye. Concurrent
increase of the intensity of donor and decrease of the intensity of acceptor is shown on the lower panel. MD simulations, including both bio-molecules as well as the water molecules, were carried out with the GROMACS code and AMBER force field, using periodic boundary condition and a cubic cell [15]. Homology modeling was used to construct the Fis–DNA initial structure. In order to model the initial Fis–DNA binding site, the Fis protein was taken from PDB structure with code 1ETY. A straight B-form DNA chain is generated by INSIGHTII. For homology comparison, we use the known Fis–DNA complex structures, with PDB code 3JR9, as template. Finally, the Fis–DNA structure was repaired for missing residues and performed by the position restrained iterations of minimizations. The helix-turn-helix DNAbinding motif was anchored within the major grooves. As shown in Figure 1a, the stacking of DNA base and its phosphate backbone make DNA a rigid and negatively charged molecule. Fis protein with positively charged (colored in blue) helix-turn-helix binding motif facilitates a specific binding with core sequences of DNA through electrostatic interactions [16]. Upon Fis docking, the posi-
C.-C. Fu et al. / Chemical Physics Letters 500 (2010) 318–322
tive charged residues on its two sides impose attractive forces to draw the flanking DNA. To simulate the global flexibility of DNA, we investigated the fluctuation of 30 -to-50 distances of the dsDNA chain (ssDNA is ne-
321
glected) over an extended period of 10 ns. In the upper panel of Figure 3a, after 2 ns of Fis binding, the length of dsDNA chain significantly decreases. This indicates that the bending direction is toward the protein side chain as shown in Figure 1b. The
Figure 4. The time evolution of DNA sequence dependent roll and opening induced by Fis (a), dsDNA alone (b) and the difference between Fis–DNA complex and dsDNA alone (c). The significant roll and opening are observed around two Fis binding sites (indicated in red underlines). Notable the roll and opening processes occur on the flanks are remarked by white dots. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
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C.-C. Fu et al. / Chemical Physics Letters 500 (2010) 318–322
corresponding histogram of length fluctuation is shown in the lower panel of Figure 3a. In order to compare with single molecule experiment performed in equilibrium, only data after 2 ns were used to construct the histogram in both Figure 3a and b. It is because in the simulation a DNA molecule takes about 2 ns to achieve equilibrium after Fis docking. Our statistical histograms show that the distributions of the two strands represented by the black and red colors were centered at ca. 8.8 nm. In addition, the time evolution of the distance between two DNA 50 ends (upper panel) and corresponding histogram (lower panel) are shown in Figure 3b. A broader Gaussian distribution was observed, suggesting the loosening of the bent DNA structure without Fis protein. To directly compare the simulation with our experimental results, two additional factors need to be considered. Firstly, adding the linker length of two dyes, the Gaussian function from simulation is shifted by ca. 2.8 nm. Secondly, the shifted Gaussian function is convoluted with the instrument response function, i.e. the distance distribution of free DA_DNA. As shown in Figure 3c, our final calculated distribution (orange) was plotted together with the experimental results. The simulation results agree very well with those of the single-molecule measurements. In Figure 4, the sequence-dependent dynamics of DNA conformation is described by step and base-pair parameters relative to the initial condition. Here, the most significant change is the rolling motion [17], with positive (or negative) value as the angle between base-pairs opens toward the minor (or major) groove, as shown in the inset of Figure 4a. The time evolution of the roll and opening angles for the Fis–DNA complex, DNA alone and the difference between the complex and DNA alone are shown in Figure 4a–c, separately. In Figure 4c, about 10–70° positive rolling angle gathers around two Fis binding sites (chain A 5–6 bp and 13–18 bp) and one flanking sequence (chain A 2–5 bp), while about 0–40° negative angle distributes on the core sequence (chain A 7–11 bp). Apparent discontinuous and large rolling angles make DNA deformed from stretched B-form to a kinked bending (rather than smooth bending) at about 60° angles as shown in Figure 1b. Note that according to the worm-like chain model, it is more favorable to be bent smoothly [7]. Another obvious change of the DNA chain is its opening motion [17], with positive (or negative) value as the angle between two bases opening toward the minor (or major) groove, as shown in the inset of Figure 4a. Some changes of the opening angle are above ±40° near the binding sites as remarked by white dots in Figure 4. Additionally, at the same regions, a dramatically base-pair stretching change as large as 0.5 Å was observed (data not shown), which are close to a quarter of a length of hydrogen bond in water. Both opening and stretching parameters are large enough to destabilize the hydrogen bonds between base-pair and partially unwind DNA. It indicates that the Fis binding breaks DNA duplex violently especially near the flank, as shown in Figure 1b. There are two contacts regions between Fis and DNA. One is practically fixed during the bending process, while the other slides along the DNA chain. This causes DNA to bend, and the twist motions of DNA helix allows the DNA base pair step motions. It appears that there exist two different kinds of motions induced by the protein–DNA binding processes. One is the bending or unzipping proceeding along the DNA helical axis, and the other is mainly the base pair opening process, which is in the direction orthogonal to the helical axis. According to the worm-like chain model [18], the Lp of polymer chain is proportional to the inverse of its flexibility, and can be obtained via its end-to-end distance RF and the contour length Lc (8.8 nm, in this study): RF2 = 2Lp[Lc + Lp(exp(Lc/Lp))] 2Lp2. As the results, after Fis binding RF reduces to 7.7 nm. Therefore, Lp becomes 10 nm, five times shorter than that of free DNA. This phe-
nomenon is more adequately described by the ‘flexible hinge’ model because Fis is able to increase the flexibility of the rod-like DNA without dissociating from the DNA. By partially rolling and opening DNA base pairs, this is due to Fis proteins create some flexible points on DNA molecules around their binding sites and the fluctuation amplitude of the roll angle of the core sequence part is ranged from 20° to 40° (Figure 4c) and the opening angle is ranged from +80° to 40°. So the fluctuation amplitude is quite large. Our studies suggest that DNA bending together with loosening is generated by Fis clipping with rolling DNA and partially unwinding its flank sides [19]. Recently, the force that Fis generated to induce long chain lambda phage DNA condensation has been measured by the magnetic tweezers approach [20]. Their results suggest that Fis may play an important role in stabilizing the loop structure by reducing the Lp of DNA to 20 nm, which is consist with our results. The scanning force microscopy (SFM) studies of long chain DNA also showed widely distributed bending angles from 50° to 120° [21]. Flexible DNA bending angles allows RNAP to easily dock nearby cofactors and then wrap DNA on its surface to stabilize RNA transcription. In addition, protein induced DNA kinking together with opening motion makes it possible for buried bases accessible to biomolecules such as RNAP and other cofactors. Highly varied degree of bending might also play important roles in bacterial chromatin structural dynamics [6]. It allows proteins to facilitate the formation of different high order protein–DNA complexes that may require different DNA angles [22]. Here, combining single molecule experiments with MD simulations provide a useful way to understand these processes at the molecular level [23]. Acknowledgments SYS and DYY gratefully acknowledge financial support of the National Science Council of Taiwan Grants: NSC-97-2113-M-010002-MY2 and NSC-97-2113-M-001-021-MY2, respectively. The authors are grateful to the National Center for High-Performance Computing of Taiwan. References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17] [18] [19] [20] [21] [22] [23]
D.F. Browning, S.J.W. Busby, Nat. Rev. Microbiol. 2 (2004) 57. J. Elf, G.W. Li, X.S. Xie, Science 316 (2007) 1191. Y.M. Wang, R.H. Austin, E.C. Cox, Phys. Rev. Lett. 97 (2006) 48302. H.S. Yuan, S.E. Finkel, J.A. Feng, M. Kaczor-Grzeskowiak, R.C. Johnson, R.E. Dickerson, Proc. Natl. Acad. Sci. USA 88 (1991) 9558. A.J. Bokal 4th, W. Ross, R.L. Gourse, J. Mol. Biol. 245 (1995) 197. R.T. Dame, Mol. Microbiol. 56 (2005) 858. C. Bustamante, Z. Bryant, S.B. Smith, Nature (London) 421 (2003) 423. C.Q. Pan, S.E. Finkel, S.E. Cramton, J.A. Feng, D.S. Sigman, R.C. Johnson, J. Mol. Biol. 264 (1996) 675. M.J. McCauley, M.C. Williams, Biopolymers 85 (2007) 154. S. Weiss, Science 283 (1999) 1676. H.P. Lu, Acc. Chem. Res. 38 (2005) 557. W.Z. Yang, T.P. Ko, L. Corselli, R.C. Johnson, H.S. Yuan, Protein Sci. 7 (1998) 1875. S.M. McLeod, J.M. Xu, R.C. Johnson, J. Bacteriol. 182 (2000) 4180. T. Ha, Methods 25 (2001) 78; W.E. Moerner, D.P. Fromm, Rev. Sci. Instrum. 74 (2003) 3597. D.A. Case et al., J. Comput. Chem. 26 (2005) 1668. J.K. Strauss, L.J. Maher 3rd, Science 266 (1994) 1829. R.E. Dickerson et al., J. Mol. Biol. 205 (1989) 787. I. Teraoka, Polymer Solutions: An Introduction to Physical Properties, Wiley, New York, 2002. J. Yan, J.F. Marko, Phys. Rev. Lett. 93 (2004) 108108. D. Skoko, J. Yan, R.C. Johnson, J.F. Marko, Phys. Rev. Lett. 95 (2005) 208101. J. Zhang, Y. Zeuner, A. Kleefeld, G. Unden, A. Janshoff, ChemBioChem 5 (2004) 1286. J. van Noort, S. Verbrugge, N. Goosen, C. Dekker, T.T. Dame, Proc. Natl. Acad. Sci. USA 101 (2004) 6969. M. Sotomayor, K. Schulten, Science 316 (2007) 1144.
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Chemical Physics Letters journal homepage: www.elsevier.com/locate/cplett
Fis-protein induces rod-like DNA bending Chi-Cheng Fu a, Ching-Fong Lin c, Quan-Ze Gao c, Wei-Zen Yang d, Tsong-Shin Lim a, Li-Ling Yang a,e, Chi-Fu Yen f, Wei-Hau Chang f,g,h,i, Hanna S. Yuan d, Sheh-Yi Sheu b,c,⇑,1, Dah-Yen Yang a,⇑,1, Wunshain Fann a,e,⇑,1, a
Institute of Atomic and Molecular Sciences, Academia Sinica, Taipei 106, Taiwan Department of Life Sciences, Institute of Genome Sciences, National Yang-Ming University, Taipei 112, Taiwan Institute of Biomedical Informatics, National Yang-Ming University, Taipei 112, Taiwan d Institute of Molecular Biology, Academia Sinica, Taipei 115, Taiwan e Department of Physics, National Taiwan University, Taipei 106, Taiwan f Institute of Chemistry, Academia Sinica, Taipei 115, Taiwan g Genomic Research Center, Academia Sinica, Taipei 115, Taiwan h Institute of Biotechnology, National Taipei University of Technology, Taipei 106, Taiwan i Institute of Engineering Technology, National Taiwan University of Science and Technology, Taipei 106, Taiwan b c
a r t i c l e
i n f o
Article history: Received 15 June 2010 In final form 11 October 2010 Available online 13 October 2010
a b s t r a c t Fis protein can bend DNA chain with length much shorter than its persistence length. We applied singlemolecule fluorescence resonance energy transfer method to probe these conformational changes. A broad distribution of end-to-end distances correlates well with the molecular dynamics simulation. The flexibility of DNA upon Fis binding is attributed to the breakages of hydrogen bonds between base pairs. DNA kinks at specific sites, instead of continuous bending. The loosening of DNA structures might have biological implications for the functions of Fis-proteins as transcription cofactors. Ó 2010 Elsevier B.V. All rights reserved.
To synthesize RNA from DNA templates requires association of RNA polymerase (RNAP) enzyme to the target genes [1]. Sophisticated regulatory mechanisms control gene transcription which involves coordinated recruitment of multiple transcriptional factors to the promoter regions [2,3]. Typically, the transcription cofactors assemble to form the pre-initiation complexes which bind directly or indirectly to DNA and activate the RNAP. In the growth phase of Escherichia coli, one of the main factors is Fis protein which binds the C-terminal region of the a-subunit of the RNAP and simultaneously bends the upstream of promoters [1,4]. By increasing the affinity of RNAP to rrnB P1 promoter about 2.5-fold, Fis can increase transcription level by 4 to 10-fold in vivo [5]. DNA bending was also observed in supercoiled bacterial chromosomes, which are highly compacted by associating with histone-like proteins, such as Fis, IHF, HU and H-NS [6]. The persistence length (Lp) of a double-stranded DNA (dsDNA) in solution is ca. 50 nm. A DNA molecule with length of 10 nm should be very rigid, and thus is hard to bend [7]. Binding of Fis can induce a significant bending of such short DNA. However, the detailed mechanism is still unclear. Fis proteins recognize specific 15 bp core sequences and induce bending of DNA with an angle ⇑ Corresponding authors. E-mail addresses: [email protected] (S.-Y. Sheu), [email protected]. edu.tw (D.-Y. Yang). 1 These authors contributed equally to this work. Deceased. 0009-2614/$ - see front matter Ó 2010 Elsevier B.V. All rights reserved. doi:10.1016/j.cplett.2010.10.026
ranging from 50° to 90°, which varies with flanking sequences [8]. Conventionally, the bending angles are measured by the gel mobility shift assays. However, these kinds of ensemble measurements cannot differentiate two models proposed to explain protein induced DNA bending phenomena [9]. One is called ‘transient kink’ model, in which protein binds to DNA only in short periods of time. It induces a kink, and then dissociates, which restores DNA to its original length. The transient binding accounts for the fluctuation of DNA conformations. The other is called ‘flexible hinge’ model, in which protein binds to DNA strongly, induces a partial denaturation of DNA, and thus creates flexible complex structures. In this Letter, a single pair fluorescence resonance energy transfer (spFRET) method [10] combined with all-atom molecular dynamics (MD) simulation was used to study the mechanism of Fis protein induced DNA bending. The spFRET technique is applied to study the bending of ‘rod-like’ DNA due to protein interactions at single molecule level. The advantage of this approach is that one can directly obtain the length distribution of DNA instead of its average value. This is of particular relevance to biological systems because most bio-molecules are flexible [11]. In this experiment, the Fis protein with Pro26Ala mutation was prepared as previously described [12]. As shown in Figure 1a, the DA_DNA is designed as a specific 26 bp dsDNA (50 -GATTT TGCAT AAAAA ACAGA CTACA T-30 ) tailed with biotinylated singlestranded DNA (ssDNA, six base, yellow). Both two 50 ends of the duplex are labeled with a donor (Alexa Fluor 546) and an acceptor (Alexa Fluor 647) fluorescent dye molecule respectively. The
C.-C. Fu et al. / Chemical Physics Letters 500 (2010) 318–322
Figure 1. Snapshots of the initial state (a) and the intermediate state at the minimum 50 –50 distance of dsDNA (b) for Fis–DNA complex structure along the MD trajectory.
donor-to-acceptor distance R is 11.6 nm, which includes the length of dsDNA (8.8 nm) and two linkers of dyes (1.4 nm for each). The tail of ssDNA allows dsDNA about 1 nm away from surface to minimize the surface effect. Note that the D_DNA, a DNA labeled with
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donor dye alone, is to the same as DA_DNA except acceptor label. The DNA samples were diluted by Fis binding buffer (with or without 2 lM Fis) at concentrations of 20 pM or 100 nM for the singlemolecule or ensemble measurements, respectively [13]. Because the equilibrium dissociation constant for Fis with DNA is 9.3 1010 M [5], majority of DNA molecules will bind with Fis for all samples. The FRET efficiency EFRET is related to R and the Förster radius R0, satisfying the EFRET = [1 + (R/R0)6]1 relationship [14]. Our time-resolved FRET measurements are based on the measurements of the averaged lifetime of the donor in the presence hsDAi and in the absence hsDi of the acceptor to obtain EFRET = 1(hsDi/hsDAi). As shown in Figure 2a, the lifetime of D_DNA and DA_DNA are 3.8 ns and 3.5 ns which correspond to EFRET = 0.079. Using the total length R as the input, R0 of this donor–acceptor pair was determined to be 7.6 nm, consistent with the value specified by the vendor (Molecular Probes). Note that this is one of the largest R0 available. After incubation with Fis, the measured lifetime decreases to 3.2 ns which corresponded to a change of R to 10 nm. For single molecular spFRET measurements [14], the biotinylated DNA molecules were immobilized on a glass plate by standard biotin–streptavidin interaction in a fluid channel. The distributions of their conformations obtained from measuring many molecules can be compared with the time-dependent conformational fluctuations simulated by MD (see below). The fluorescence intensities of both donor ID and acceptor IA were measured simultaneously by a confocal microscopy equipped with a double port. We can obtain the FRET efficiency in terms of EFRET = IA/(ID + IA). The upper panel of Figure 2b shows that the EFRET histogram of DA_DNA sample is well fitted with two Gaussian distributions. Apparently, the first distribution (blue), centered at zero, can be attributed to DNA molecules containing donors but inactive acceptors. This was confirmed by measuring the D_DNA sample (data not shown). The second distribution (green) is centered at 0.072 with width of 0.045. Its center corresponds to a mean distance of R = 11.64 nm, which agrees with the length of our designed DNA; its width corresponds to the R ranging from 10.64 nm to 13.81 nm, as shown in Figure 3c (green dash line). Notice that even the energy transfer distribution, Figure 2b, is symmetric; the corresponding distance
Figure 2. (a) The ensemble time-resolved FRET results of D_DNA, DA_DNA and DA_DNA + Fis. (b) The EFRET histograms of DA_DNA (upper panel) and DA_DNA + Fis (lower panel) measured by the single molecular method. The inset of (b) shows typical single spFRET molecule time traces.
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Figure 3. Time evolution (upper panel) of 30 -to-50 (a) and 50 -to-50 (b) distance of dsDNA binding with Fis by MD simulation and the corresponding histogram (lower panel). The chains A and B of dsDNA are colored in black and red (a). The 50 -to-50 distances of dsDNA in Fis–DNA complex and dsDNA alone are colored in orange and green (b). (c) The donor–acceptor distance R distributions of DA_DNA (green dash lines) and DA_DNA binding with Fis (red dash line). The orange line is the convolution of green dash line with the distribution function shown in the lower panel of (b). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
distribution, Figure 3c, is asymmetric. This is due to the fact that the average distance, 11.64 nm, is much longer than R0, 7.6 nm. Thus the measurements operated at the tail of the transfer curve. After incubating with Fis, an additional third distribution attributed to the Fis–DNA complex (red) appeared in the lower panel of Figure 2b. The first two distributions (blue and green) are caused by the DA_DNA without Fis binding, as referred to the similar distributions seen in the DA_DNA only measurements. The third distribution has a mean value at 0.12 with width of 0.08. The center energy transfer value corresponds to R = 10.59 nm, which agrees well with the results of ensemble time-resolved measurement. This agreement suggests that the third distribution should represent the conformation distribution of DNA–Fis complexes in solution. The width of this distribution corresponded to the R between 9.58 and 12.91 nm, indicating the higher flexibility of the DNA–Fis complex as shown in Figure 3c (red dash line). Two typical single molecule time traces were shown in the inset of Figure 2b. In the upper panel, the intensity of accepter drops to the background following the photobleach of donor dye. Concurrent
increase of the intensity of donor and decrease of the intensity of acceptor is shown on the lower panel. MD simulations, including both bio-molecules as well as the water molecules, were carried out with the GROMACS code and AMBER force field, using periodic boundary condition and a cubic cell [15]. Homology modeling was used to construct the Fis–DNA initial structure. In order to model the initial Fis–DNA binding site, the Fis protein was taken from PDB structure with code 1ETY. A straight B-form DNA chain is generated by INSIGHTII. For homology comparison, we use the known Fis–DNA complex structures, with PDB code 3JR9, as template. Finally, the Fis–DNA structure was repaired for missing residues and performed by the position restrained iterations of minimizations. The helix-turn-helix DNAbinding motif was anchored within the major grooves. As shown in Figure 1a, the stacking of DNA base and its phosphate backbone make DNA a rigid and negatively charged molecule. Fis protein with positively charged (colored in blue) helix-turn-helix binding motif facilitates a specific binding with core sequences of DNA through electrostatic interactions [16]. Upon Fis docking, the posi-
C.-C. Fu et al. / Chemical Physics Letters 500 (2010) 318–322
tive charged residues on its two sides impose attractive forces to draw the flanking DNA. To simulate the global flexibility of DNA, we investigated the fluctuation of 30 -to-50 distances of the dsDNA chain (ssDNA is ne-
321
glected) over an extended period of 10 ns. In the upper panel of Figure 3a, after 2 ns of Fis binding, the length of dsDNA chain significantly decreases. This indicates that the bending direction is toward the protein side chain as shown in Figure 1b. The
Figure 4. The time evolution of DNA sequence dependent roll and opening induced by Fis (a), dsDNA alone (b) and the difference between Fis–DNA complex and dsDNA alone (c). The significant roll and opening are observed around two Fis binding sites (indicated in red underlines). Notable the roll and opening processes occur on the flanks are remarked by white dots. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
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corresponding histogram of length fluctuation is shown in the lower panel of Figure 3a. In order to compare with single molecule experiment performed in equilibrium, only data after 2 ns were used to construct the histogram in both Figure 3a and b. It is because in the simulation a DNA molecule takes about 2 ns to achieve equilibrium after Fis docking. Our statistical histograms show that the distributions of the two strands represented by the black and red colors were centered at ca. 8.8 nm. In addition, the time evolution of the distance between two DNA 50 ends (upper panel) and corresponding histogram (lower panel) are shown in Figure 3b. A broader Gaussian distribution was observed, suggesting the loosening of the bent DNA structure without Fis protein. To directly compare the simulation with our experimental results, two additional factors need to be considered. Firstly, adding the linker length of two dyes, the Gaussian function from simulation is shifted by ca. 2.8 nm. Secondly, the shifted Gaussian function is convoluted with the instrument response function, i.e. the distance distribution of free DA_DNA. As shown in Figure 3c, our final calculated distribution (orange) was plotted together with the experimental results. The simulation results agree very well with those of the single-molecule measurements. In Figure 4, the sequence-dependent dynamics of DNA conformation is described by step and base-pair parameters relative to the initial condition. Here, the most significant change is the rolling motion [17], with positive (or negative) value as the angle between base-pairs opens toward the minor (or major) groove, as shown in the inset of Figure 4a. The time evolution of the roll and opening angles for the Fis–DNA complex, DNA alone and the difference between the complex and DNA alone are shown in Figure 4a–c, separately. In Figure 4c, about 10–70° positive rolling angle gathers around two Fis binding sites (chain A 5–6 bp and 13–18 bp) and one flanking sequence (chain A 2–5 bp), while about 0–40° negative angle distributes on the core sequence (chain A 7–11 bp). Apparent discontinuous and large rolling angles make DNA deformed from stretched B-form to a kinked bending (rather than smooth bending) at about 60° angles as shown in Figure 1b. Note that according to the worm-like chain model, it is more favorable to be bent smoothly [7]. Another obvious change of the DNA chain is its opening motion [17], with positive (or negative) value as the angle between two bases opening toward the minor (or major) groove, as shown in the inset of Figure 4a. Some changes of the opening angle are above ±40° near the binding sites as remarked by white dots in Figure 4. Additionally, at the same regions, a dramatically base-pair stretching change as large as 0.5 Å was observed (data not shown), which are close to a quarter of a length of hydrogen bond in water. Both opening and stretching parameters are large enough to destabilize the hydrogen bonds between base-pair and partially unwind DNA. It indicates that the Fis binding breaks DNA duplex violently especially near the flank, as shown in Figure 1b. There are two contacts regions between Fis and DNA. One is practically fixed during the bending process, while the other slides along the DNA chain. This causes DNA to bend, and the twist motions of DNA helix allows the DNA base pair step motions. It appears that there exist two different kinds of motions induced by the protein–DNA binding processes. One is the bending or unzipping proceeding along the DNA helical axis, and the other is mainly the base pair opening process, which is in the direction orthogonal to the helical axis. According to the worm-like chain model [18], the Lp of polymer chain is proportional to the inverse of its flexibility, and can be obtained via its end-to-end distance RF and the contour length Lc (8.8 nm, in this study): RF2 = 2Lp[Lc + Lp(exp(Lc/Lp))] 2Lp2. As the results, after Fis binding RF reduces to 7.7 nm. Therefore, Lp becomes 10 nm, five times shorter than that of free DNA. This phe-
nomenon is more adequately described by the ‘flexible hinge’ model because Fis is able to increase the flexibility of the rod-like DNA without dissociating from the DNA. By partially rolling and opening DNA base pairs, this is due to Fis proteins create some flexible points on DNA molecules around their binding sites and the fluctuation amplitude of the roll angle of the core sequence part is ranged from 20° to 40° (Figure 4c) and the opening angle is ranged from +80° to 40°. So the fluctuation amplitude is quite large. Our studies suggest that DNA bending together with loosening is generated by Fis clipping with rolling DNA and partially unwinding its flank sides [19]. Recently, the force that Fis generated to induce long chain lambda phage DNA condensation has been measured by the magnetic tweezers approach [20]. Their results suggest that Fis may play an important role in stabilizing the loop structure by reducing the Lp of DNA to 20 nm, which is consist with our results. The scanning force microscopy (SFM) studies of long chain DNA also showed widely distributed bending angles from 50° to 120° [21]. Flexible DNA bending angles allows RNAP to easily dock nearby cofactors and then wrap DNA on its surface to stabilize RNA transcription. In addition, protein induced DNA kinking together with opening motion makes it possible for buried bases accessible to biomolecules such as RNAP and other cofactors. Highly varied degree of bending might also play important roles in bacterial chromatin structural dynamics [6]. It allows proteins to facilitate the formation of different high order protein–DNA complexes that may require different DNA angles [22]. Here, combining single molecule experiments with MD simulations provide a useful way to understand these processes at the molecular level [23]. Acknowledgments SYS and DYY gratefully acknowledge financial support of the National Science Council of Taiwan Grants: NSC-97-2113-M-010002-MY2 and NSC-97-2113-M-001-021-MY2, respectively. The authors are grateful to the National Center for High-Performance Computing of Taiwan. References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17] [18] [19] [20] [21] [22] [23]
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