Báo cáo khoa học: Fluorescence studies of the replication initiator protein RepA in complex with operator and iteron sequences and free in solution pdf

Fluorescence studies of the replication initiator protein RepA in complex with operator and iteron sequences and free in solution ´ Rutger E M Diederix1,2, Cristina Davila1,2, Rafael Giraldo2 and M Pilar Lillo1 ´ ´ ´ Departamento de Biofısica, Instituto de Quımica Fısica ‘Rocasolano’, CSIC, Madrid, Spain ´ ´ Departamento de Microbiologıa Molecular, Centro de Investigaciones Biologicas, CSIC, Madrid, Spain Keywords anisotropy; DNA replication; fluorescence; hydrodynamics; RepA Correspondence ´ M P Lillo, Departamento de Biofısica, ´ ´ Instituto de Quımica Fısica ‘Rocasolano’, CSIC, Serrano 119, 28006 Madrid, Spain Fax: +34 91 564 2431 Tel: +34 91 561 9400, ext 1027 E-mail: pilar.lillo@iqfr.csic.es (Received 26 June 2008, revised August 2008, acccepted September 2008) doi:10.1111/j.1742-4658.2008.06669.x RepA, the replication initiator protein from the Pseudomonas plasmid pPS10, regulates plasmid replication and copy number It is capable of autorepression, in which case it binds as a dimer to the inverted repeat operator sequence preceding its own gene RepA initiates plasmid replication by binding as a monomer to a series of four adjacent iterons, which contain the same half-repeat as found in the operator sequence RepA contains two domains, one of which binds specifically to the half-repeat The other is the dimerization domain, which is involved in protein–protein interactions in the dimeric RepA–operon complex, but which actually binds DNA in the monomeric RepA–iteron complex Here, detailed fluorescence studies on RepA and an N-(iodoacetyl)aminoethyl-8-naphthylamine-1-sulfonic acidlabeled single-cysteine mutant of RepA (Cys160) are described Using timeresolved fluorescence depolarization measurements, the global rotational correlation times of RepA free in solution and bound to the operator and to two distinct iteron dsDNA oligonucleotides were determined These provide indications that, in addition to the monomeric RepA–iteron complex, a stable dimeric RepAiteron complex can also exist Further, Forster resoă nance energy transfer between Trp94, located in the dimerization domain, and N-(iodoacetyl)aminoethyl-8-naphthylamine-1-sulfonic acid-Cys160, located on the DNA-binding domain, is observed and used to estimate the distance between the two fluorophores This distance may serve as an indicator of the orientation between both domains in the unbound protein and RepA bound to the various cognate DNA sequences No major change in distance is observed and this is taken as evidence for little to no re-orientation of both domains upon complex formation RepA is the DNA replication initiator protein of the Pseudomonas plasmid pPS10 It is representative of a family of plasmid replication initiators active in many Gram-negative bacteria, including the initiators from plasmids such as pSC101, F and R6K [1] The operator region preceding the repA gene contains a partially palindromic sequence (inverted repeat, IR) to which RepA can bind, which acts as an autogenous repressor of transcription [2] The plasmid also carries an origin of replication, containing a sequence with four contiguous tandem repeats (direct repeats, DR; termed iterons) of the same bp sequence found inverted in the operator region of RepA RepA thus has dual DNA-binding activity: it can bind as a dimer to its operator region, in which case it functions in transcription repression; and it can bind in a highly cooperative fashion to the four directly repeated iterons, in which case it functions in replication initiation [3] Abbreviations (I)AEDANS, N-(iodoacetyl)aminoethyl-8-naphthylamine-1-sulfonic acid; FRET, Forster resonance energy transfer ă FEBS Journal 275 (2008) 5393–5407 ª 2008 The Authors Journal compilation ª 2008 FEBS 5393 Fluorescence studies of RepA R E M Diederix et al Interestingly, in the latter case, the protein binds as a monomer [2–6] Free in solution, the protein is essentially dimeric, but it dissociates and binds as a monomer in the presence of even a single iteron sequence [2,3] The mechanism by which this occurs is unclear, but it involves considerable conformational changes in RepA [3,4] judged by comparison of crystal structures of (truncated) RepA dimer [5] and the monomeric RepA– iteron complex that was modeled on the complex structure of the close homolog RepE from the F plasmid [2,7–9] For the latter protein, the crystal structures of both the monomer–iteron and dimer–operator complexes are available, indicating secondary structural changes in the linker connecting the dimerization and DNA-binding domains, and rearrangement of the relative orientation of the two domains [7,9] The conformational change upon iteron binding may expose a recognition site for protein–protein interaction, enabling coupling of recently replicated origins from different plasmid molecules [10,11] This so-called handcuffing is thought to be the mechanism for replication inhibition in iteron-containing plasmids [12] Following our series of biophysical studies of RepA [3–6], we report hydrodynamic and structural studies on RepA and its complexes with operator and single iteron sequences Global rotational correlation times were determined by fluorescence anisotropy decay experiments using the extrinsic fluorophore N-(iodoacetyl)aminoethyl-8-naphthylamine-1-sulfonic acid (AEDANS), specifically coupled to Cys160 in the single-cysteine mutant C160–RepA The AEDANS probe was also used as a Forster resonance energy ă transfer (FRET) acceptor to monitor putative interdomain movements in RepA upon binding the various DNA sequences We show that, despite the extensive structural rearrangement that is known to occur upon monomerization and DNA binding to the iteron sequence [3–6], an appreciable change in the interdomain organization is not actually observed Finally, it appears that monomerization does not occur efficiently in very short oligonucleotides that contain few bases more than the iteron sequence, and RepA binds as a dimer instead Results Labeling and characterization of C160–RepA C160–RepA is a double-mutant of His6-tagged wildtype RepA [4] in which two wild-type Cys residues (C29 and C106) have been changed to Ser The single remaining Cys160 is located on the C-terminal DNA5394 binding domain of RepA, also called the WH2 domain, which specifically recognizes the operator and iteron sequences [1,2] Most C160–RepA is expressed in inclusion bodies, and the His6-tagged protein was purified from solubilized inclusion bodies using Ni(II)affinity chromatography under denaturing conditions As shown previously [3,4], the His6-tag does not interfere with protein function or structure, and it was not removed after purification Refolding of C160–RepA is by rapid 20-fold dilution in buffer (0.15 m (NH4)2SO4, 15 mm Na-acetate, 0.03 mm EDTA, 3% glycerol, pH 6.0) Almost all the protein is recovered and is present as a single, soluble species Refolded C160– RepA is dimeric, as judged by size-exclusion chromatography, where it elutes at exactly the same volume as wild-type RepA (not shown) Labeling of the single Cys of native C160–RepA with IAEDANS gives very low yields (< 5%) The yield can be improved significantly (to 50%) by performing the labeling reaction under conditions where the protein is unfolded, i.e in the presence of 5.6 m guanidinium hydrochloride Presumably, this poor reactivity is related to the low solubility of the native protein (up to 10–20 lm) Under denaturing conditions, RepA can easily be concentrated 10- to 100-fold, thus favoring the bimolecular labeling reaction greatly under the conditions of $ 15-fold excess IAEDANS The CD spectrum of unlabeled or AEDANS-labeled C160–RepA is indistinguishable from that of wild-type RepA at °C (Fig 1A), indicating that the secondary structure is not affected by the mutation or by AEDANS labeling Thermal denaturation analysis of the protein variants suggests a lower stability of the mutant (Fig 1B) The C160–RepA variants show a lower melting temperature than wild-type RepA (60 versus 67 °C for wild-type RepA), and the thermal transition of unlabeled C160–RepA has a substantially lower slope (reduced co-operativity) than wild-type RepA and the labeled variant However, room temperature is well below the melting transition, and as the experiments described here have been performed at or below this temperature, it can safely be assumed that the mutant protein is fully folded This is supported by the observation that the fluorescence emission spectrum of the unique Trp residue (W94), a sensitive indicator of the folding state of the dimerization domain of RepA [4], is unchanged in the mutant with respect to that of wild-type RepA (Fig 1C) Finally, AEDANS C160–RepA and wild-type RepA show identical binding to the operator sequence (Fig 1D), confirming that mutation and labeling not affect the function, and by implication therefore also the structure, of RepA FEBS Journal 275 (2008) 5393–5407 ª 2008 The Authors Journal compilation ª 2008 FEBS R E M Diederix et al A B Fluorescence studies of RepA Fig (A) Near- and far-UV CD spectra of wild-type RepA (solid line) and C160–RepA both unlabeled (dashed line) and AEDANSlabeled (dash-dots) The spectra were recorded at °C with $ 3.5 lM wild-type and unlabeled C160–RepA, and lM AEDANS C160–RepA The buffer spectrum is subtracted and the spectra have been transformed to mean residual ellipticity units (B) Thermal denaturation curves for wild-type RepA (solid lines) and C160– RepA unlabeled (dashed line) and AEDANS-labeled (dash-dots) The temperature dependence of the ellipticity at 220 nm is shown, normalized to help compare the different proteins (C) Fluorescence emission spectra (kex = 295 nm) of wild-type RepA (solid line), C160–RepA both unlabeled (dashed line) and AEDANS labeled (dash-dots), recorded at 23.5 °C with $ lM protein and with intensities normalized with respect to their emission maximum at 327 nm (D) Binding of wild-type RepA ( ) and AEDANS C160– RepA (s) to 10 nm Alexa568-labeled 1IR, monitored by Alexa568 fluorescence anisotropy (kex = 535 nm, kem = 605 nm) Data for both proteins were fitted (see Eqns and 5) together (solid line) to a : RepA : 1IR binding equilibrium using the quadratic equation This yielded Kd = ± nm, compatible with previous reports [3] Experiments were carried out in 0.15 M (NH4)2SO4, 15 mM NH4acetate, 0.03 mM EDTA, 3% glycerol; pH 6.0 C A B D Figure 2A shows the emission spectrum of AEDANS C160–RepA excited at 295 and 375 nm, respectively When excited at 295 nm, fluorescence contributions from both AEDANS and W94 are visible Figure 2B shows the excitation spectrum of the acceptor (kem = 480 nm) There is a clear contribution from W94 visible as a shoulder at 280–290 nm, which is assignable to FRET from W94 to C160-AEDANS Fig (A) Fluorescence emission spectra of AEDANS-labeled C160–RepA, excited at 295 nm (solid line) and 375 nm (dashed line) The spectra are normalized with respect to the emission intensity at 484 nm (B) Excitation spectrum of AEDANS C160– RepA, measured at 480 nm The arrow indicates the contribution of Trp fluorescence The spectra were recorded at 23.5 °C, in 0.15 M (NH4)2SO4, 15 mM NH4-acetate, 0.03 mM EDTA, 3% glycerol; pH 6.0 [RepA] was $ lM FEBS Journal 275 (2008) 5393–5407 ª 2008 The Authors Journal compilation ª 2008 FEBS 5395 Fluorescence studies of RepA R E M Diederix et al Binding of C160–RepA to operator and iteron sequences followed by AEDANS fluorescence The fluorescence of AEDANS–C160 was studied as a function of DNA concentration for the operator and two distinct iteron sequences (described in Table 1) RepA binding to 1IR and 1DR has been studied in detail previously [3,6] When increasing amounts of 1IR, 1DR or 1DR-short are added to AEDANS C160–RepA, no effect is seen on the shape or intensity of the ‘pure’ AEDANS fluorescence, i.e the emission spectrum excited at 375 nm (not shown) There is, however, a clear increase in the fluorescence anisotropy for each of the sequences (Fig 3D–F), indicating a decrease in the rotational mobility of AEDANS C160– RepA The anisotropy increase is slightly different for each of the three sequences, and relates to an increased global rotational correlation time for the AEDANS probe caused by C160–RepA binding to DNA (see below) Addition of DNA also induces a change in the shape of the excitation spectra This is caused by a decrease in the Trp contribution to AEDANS fluorescence, as illustrated by the difference spectra between free and bound RepA, which are typical of Trp (Fig 3A–C) The increase in directly excited AEDANS anisotropy matches very well with the decrease in W94 contribution to AEDANS fluorescence for each of the three Table Sequence of the oligonucleotides used in this study IR (operator, half sites in bold), 1DR (single iteron underlined, with the half site also present in the operator in bold, purported DnaA box dashed underlined), 1DR-short (single iteron underlined, with the half site also present in the operator in bold) Name Length (bp) Sequence 1IR 1DR 1DR-short 39 45 30 GAACAAGGACAGGGCATTGACTTGTCCCTGTCCCTTAAT ATACCCGGGTTTAAAGGGGACAGATTCAGGCTGTTATCCACACCC GCCCGGGTTTAAAGGGGACAGATTCAGGCC A D B E C F 5396 Fig Excitation spectra (kem = 480 nm) of AEDANS C160–RepA with increasing concentrations of 1IR, 1DR and 1DR-short (A, B and C, respectively), causing changes in the direction of the arrows The spectra are inner filter corrected and normalized to the intensity at 340 nm Difference spectra between free and DNA-bound RepA are shown as dashed lines RepA was 1.25 lM and 0, 0.2, 0.4, 0.6 and lM 1IR (A), 0, 0.4, 1.2, 2.4 and lM 1DR (B), and 0, 0.8, 1.8, 3.2 and lM 1DR-short (C) (D) Fluorescence intensity (kex = 280 nm, kem = 480 nm), corrected and normalized as in (A) ( ), and AEDANS fluorescence anisotropy (kex = 375 nm, kem = 480 nm) of AEDANS C160–RepA as a function of [1IR] (s) Data were fit using the quadratic binding equation (see Eqns 3–4) (E) and (F) as (D), except they refer to titrations with 1DR and 1DR-short, respectively Experiments were performed at 23.5 °C, in 0.15 M (NH4)2SO4, 15 mM NH4-acetate, 0.03 mM EDTA, 3% glycerol, pH 6.0 FEBS Journal 275 (2008) 5393–5407 ª 2008 The Authors Journal compilation ª 2008 FEBS R E M Diederix et al Fluorescence studies of RepA tested oligonucleotides (Fig 3D–F) The change in fluorescence and anisotropy were fit simultaneously for each titration In the fits, the protein concentration was left free, to serve as an indicator of stoichiometry In the case of 1IR, the fit resulted in a binding stoichiometry of : 1, i.e dimer binding The reactant concentrations were too high to obtain relevant information on the binding affinity For binding to 1DR, the best fit yielded a binding stoichiometry of $ : 1, i.e monomer binding, with a Kd between 0.2 and 0.6 lm With 1DR-short, a reliable estimate for the stoichiometry of binding could not be made Assuming binding as monomer or as dimer, respectively, the dissociation constants obtained were 2.1 ± 0.2 and 2.9 ± 0.2 lm, without an apparent difference in goodness of fit However, in a separate experiment involving inter-monomeric homoFRET (see below) the binding stoichiometry was confirmed as dimeric RepA to the 1IR and IDR-short sequences, and monomeric RepA to 1DR The binding affinity under these conditions is thus 2.9 lm FRET between Trp94 and the AEDANS As mentioned above, DNA binding induces an apparent decrease in FRET efficiency between W94 and AEDANS–C160 Along with this decrease, there is also a considerable degree of quenching of W94 fluorescence This residue has a relatively high quantum yield for Trp [13] that is strongly quenched upon binding to its cognate DNA sequences (see Table 2) This quenching is unrelated to FRET, as it also occurs with unlabeled RepA Furthermore, it does not decrease the lifetime of W94 fluorescence significantly (see Table S1), Table Fluorescence and FRET parameters of the W94– AEDANS–C160 pair and resulting average inter-probe distances, in free RepA and RepA bound to various cognate DNA sequences FRET efficiency was determined using Eqn (1), and assuming   eW 94 eAEDANS = and eAEDANS eAEDANS = 0.17 (see Experimental 280 nm 340 nm 280 nm 340 nm procedures) The apparent quantum yield of W94 (FW94) was determined both for wild-type RepA and unlabeled C160–RepA The degree of quenching, i.e the ratio of FW94 in free and DNA-bound RepA was used to determine the fraction of fluorescent donor (d+ in Eqn 1) Species FW94 (± 0.02) d+ (± 0.08) FRET efficiency (± 0.15) / ˚ R(23) (A) (± 7)b free RepA + 1IR + 1DR + 1DR-short 0.29 0.14 0.21 0.16a 1.00 0.48 0.72 0.55a 0.7 0.8 0.6 0.8a 22 20 23 20 a Values based on extrapolations from binding curves and as such ˚ not experimentally confirmed b Using R0(23) = 25 ± A / indicating that it is static in nature We not have an unequivocal interpretation of the origin of the static quenching However, judging from the binding stoichiometry together with the shape of the binding curves (Fig 3), it is safe to conclude that the quenching does not affect the RepA–DNA binding equilibria, and thus that dark state(s) of W94 are present in the RepA– DNA complexes Because the fraction of non-fluorescent donor molecules does not contribute to the Trp fi AEDANS energy transfer process, a correction of the FRET efficiencies for the presence of nonfluorescent W94 is required (see Eqn 1, Experimental procedures) After doing so, it appears that the difference in FRET efficiency between free RepA and its DNA complexes is actually relatively minor (see Table 2) Accordingly, the resulting distance calculated between W94 and AEDANS–C160 does not display large variations between the different species However, there are a number of caveats that should be taken into account First, there are several tyrosine residues in RepA As the fluorescence was excited at 280 nm, there is the possibility that some of the five tyrosines present in the W94-containing N-terminal domain of RepA also contribute to the experimental FRET efficiency, by Tyr fi Trp energy transfer As the distance between W94 and the nearest Tyr residue ˚ is $ 15 A [5], this contribution is negligible, however This conclusion is well supported by the apparent lack of contribution of Tyr to the excitation spectrum of acceptor AEDANS indicated in the excitation difference spectra seen in Fig 3AC Second, the Forster ă and donor-acceptor distances determined here, relate to the R0 value determined assuming hj2 i = ⁄ 3, R0(2/3) (see Experimental procedures) This value was ˚ calculated to be 25 ± A The value of hj2 i is not known exactly, leading to additional uncertainty The maximum and minimum limits of the value of hj2 i for the W94 ⁄ AEDANS–C160 couple in RepA were estimated as described previously [14,15], from the depolarization factors determined from time-resolved fluorescence anisotropy recorded for wild-type RepA W94 and AEDANS C160–RepA (see below, and Table S1) It appears that the factor hj2 i for RepA– DNA complexes would have a value between 0.06 and 3.51, which in turn yields an uncertainty in the absolute distance between 0.7 and 1.3 times the value of R(2/3), presented in Table Nevertheless, the R(2/3) value in the complex with 1DR is in excellent agreement with the distance measured between the Cb atoms of both residues in the structural model of RepA [2] based on the monomer– iteron complex structure of the homologous RepE protein [7] W94 and C160 are each located on one of the FEBS Journal 275 (2008) 5393–5407 ª 2008 The Authors Journal compilation ª 2008 FEBS 5397 Fluorescence studies of RepA R E M Diederix et al two different domains of RepA, and therefore changes in distance between both residues can be interpreted in terms of domain movements Because no relevant change is observed, it can be concluded that no significant reorientation takes place between the two domains of RepA upon binding to the operator or iteron DNA or as a result of the monomerization of RepA that accompanies binding to 1DR We can not currently exclude a rotation centered about C160, as this will also not affect the distance between both residues Also, note that, in theory, inter-monomeric FRET may occur in the case of RepA dimers This is unlikely however, considering the distance between both W94 residues ˚ ($ 20 A) and that both DNA binding domains containing the AEDANS probes are located roughly on opposing ends of the dimerization domains [5] Time-resolved fluorescence depolarization and rotational correlation times of RepA and its DNA complexes Time-resolved depolarization measurements were performed to obtain information on global and local A C 5398 dynamics of the AEDANS and W94 probes in free and DNA-bound RepA The decay of the total fluorescence intensity was recorded, as well as the decays of its vertically and horizontally polarized components The anisotropy decay of the fluorophore can be described in terms of its slow and fast components, i.e of global and local re-orientational motions, respectively This was carried out for both W94 in wild-type RepA and AEDANS-labeled C160–RepA AEDANS has a much longer fluorescence lifetime than Trp, allowing a much greater level of confidence in the determination of correlation times pertaining to the global rotational motion Nevertheless, the global rotational information obtained from Trp fluorescence anisotropy decays (see Fig S1 and Table S1) shows a trend in agreement with the data from the AEDANS experiments Furthermore, despite the relatively poor photon-counting statistics, the local dynamics of W94 have been characterized from the Trp decays In Fig 4, anisotropy decays (kem = 480 nm) are shown for the different AEDANS C160–RepA species, together with best fits assuming a bi-exponential function for r(t) (see Experimental procedures) The B D Fig Anisotropy decays R(t) (kex = 375 nm, kem = 480 nm) of AEDANS C160–RepA free in solution (A) and bound to 1IR (B), 1DR (C) and 1DR-short (D) Experiments were performed at 23.5 °C in 0.15 M (NH4)2SO4, 15 mM NH4-acetate, 0.03 mM EDTA, 3% glycerol, pH 6.0 Experimental data (s) were reconstructed from the fluorescence decays that were polarized parallel and perpendicular to the polarization plane of the excitation beam, after subtracting their respective dark counts Fits to the data are shown as solid gray lines AEDANS C160–RepA was $ lM in each experiment and with 2.5 lM 1IR, lM 1DR and 12 lM 1DR-short, respectively Weighted residuals for the fits between experimental and calculated R(t) are shown below the curves FEBS Journal 275 (2008) 5393–5407 ª 2008 The Authors Journal compilation ª 2008 FEBS R E M Diederix et al Fluorescence studies of RepA Table Fluorescence lifetimes, time-resolved and steady-state anisotropy parameters for AEDANS–C160 in free RepA and RepA bound to various cognate DNA sequences.a Sample hrss i ± 0.002 hsic (ns) ± 0.4 b1 ± 0.05 /1 (ns) ±3 b2 ±0.05 /2 (ns) ± 10 /2 calc (ns) (max–min) Free RepA + 1IR + 1DR + 1DR-short 0.209 0.239b 0.237b 0.238b 13.1 12.2 13.4 13.5 0.234 0.156 0.160 0.150 4.5 2.7 4.2 7.3 0.766 0.844 0.840 0.850 56 109 97 98 (42–89)d (61–131)d (43–138)d (35–59)e Estimated errors at the 67% confidence level [30] b Steady-state anisotropy from fits to the data in Fig c kex = 375 nm, kem = 480 nm; r0 (from the fits) = 0.31 ± 0.015); T = 23.5 °C d Minimum and maximum calculated rotational correlation times assuming a prolate ellipsoid shape, and using shape factors from frictional ratios previously [3] determined using sedimentation velocity experiments e Minimum and maximum calculated rotational correlation times calculated using the HYDROPRO program [17] using as input homology models of the RepA– 1DR and 1DR-short complexes, respectively, based on the crystal structure [7] of monomeric RepE in complex with iteron DNA a analogous decays with kem = 530 nm, with corresponding best fits and tabulated parameters, are supplied in Fig S2 and Table S1 The AEDANS data confirm the presence of discrete complexes under the conditions of the experiment, and that binding is complete, in agreement with the binding curves (Fig 3), except for the case of the complex with 1DR-short, which under these conditions should contain $ 20% free RepA As expected, the global rotational correlation time, /2, increases upon binding of RepA to its cognate DNA Apart from the RepA–1DR-short complex, the observed values easily fall within the range reasonably expected from molecules of this size and shape (Table 3) The expected values for free RepA and the dimeric RepA– 1IR complex were calculated on basis of hydrodynamic shapes and volumes corresponding to prior [3] sedimentation velocity measurements as shown in Fig Both can be characterized as rigid elongated shapes For the monomer RepA–1DR complex, the structure modeled on the homologous mRepE–DNA crystal structure [7] was used directly to calculate the expected global rotational correlation time The calculated values for the RepA–1DR-short complex pertain to a monomer, i.e the modeled structure as above, but with a truncated oligonucleotide having 30 bp instead of the 45 bp of 1DR This purported complex of monomeric RepA with 1DR-short is not shown, but it is easily imagined that this complex is quite spherical and that it should have a relatively short global rotational correlation time This is clearly not what is observed Note that because the orientation of the AEDANS probe in the complex is not known, we provide a range of calculated values, i.e the minimum and maximum of the correlation times corresponding to the complex (see Experimental procedures) Nevertheless, even given this significant uncertainty, the measured value of the complex with 1DR-short clearly Fig Prolate ellipsoids equivalent to (non-hydrated) free RepA (upper) and RepA–1IR complex (lower), with axial ratio and volumes corresponding to frictional ratios determined from prior sedimentation velocity analysis (3) and molecular mass (23) respectively The modeled structure of monomeric RepA–1DR is shown in two orientations (center) For clarity, the purported structure of RepA monomer with 1DR-short is not shown The length of 1DR-short only allows for five nucleotides (half a helical turn) to protrude from either end of the protein–DNA interface exceeds the maximum value that was calculated for a hypothetical complex involving RepA monomer By contrast, a correlation time around 100 ns fits well with a complex involving dimeric RepA and a 30 bp oligonucleotide It should further be noted that the presence of 20% free RepA in the case of the 1DRshort complex will lead to a slight underestimation of the rotational correlation time There appears to be linear correlation between oligonucleotide size (zero for free RepA) and experimental correlation time for the complexes involving dimeric RepA (Table 3) Only the complex between 1DR and RepA does not fit this FEBS Journal 275 (2008) 5393–5407 ª 2008 The Authors Journal compilation ª 2008 FEBS 5399 Fluorescence studies of RepA R E M Diederix et al correlation, in line with the fact that it is the only complex involving monomeric RepA Finally, we note that the range of global rotational correlation times calculated for the dimer–operator complex of the F plasmid RepE protein, which is highly homologous to RepA and of which the crystal structure is known [9], is shorter (57–83 nucleotides) than observed here for the RepA–1IR complex This could mean that there are significant differences between the RepE– and RepA–operator complexes, which are possibly related to the different spacing between the half repeats in both operator DNA sequences [9] A B Oligomerization state of free and complexed RepA determined by homoFRET In order to understand the oligomerization state of RepA in the different DNA complexes better, homoFRET experiments were carried out Herein, use is made of C160–RepA specifically labeled with Atto532 In a double-labeled sample, FRET is expected to occur between the two Atto532 moieties whenever the interprobe distance is not greater than $ 1.5 times the ˚ Forster distance The calculated R0(2/3) = 55 A for ă Atto532Atto532 homoFRET, and thus energy transfer is expected to occur in double-labeled RepA dimers Thus, no FRET is expected when RepA is monomeric, or in single-labeled Atto532–RepA dimers HomoFRET between the fluorophores is detected through depolarization of their emission, but note that this occurs only if they not happen to be aligned more-or-less parallel to each other in the dimer C160–RepA samples labeled with 60% Atto532 (i.e with 43% of Atto532 residing on double-labeled RepA dimers) show clearly different excitation anisotropy spectra from C160–RepA samples labeled with only 10% Atto532, i.e with very little double-labeled RepA dimers (< 5%) This is shown in Fig 6A, where there is an evident decrease in anisotropy for the sample containing the double-labeled C160–RepA dimers, which is less pronounced at longer excitation wavelengths (red-edge excitation) The enhanced fluorescence depolarization in the double-labeled dimers with respect to the single-labeled samples is a clear indication of homoFRET in the double-labeled samples [16] The increase in steady-state anisotropy observed upon decreasing the degree of Atto532-labeling from 60% to 10% is also observed when excess 1DR is added to 60% labeled Atto532 C160–RepA, but not upon the addition of excess 1IR and 1DR-short (Fig 6B) This means that addition of 1DR abolishes the homoFRET, by inducing RepA monomerization In fact, the addi5400 Fig (A) Excitation anisotropy spectra of Atto532–C160 RepA labeled to different degrees (solid line: 60% label, dashed line: 10%) [RepA] is 0.5 lM in either case, and conditions are 0.5 M (NH4)2SO4, 50 mM NH4-acetate pH 6.0, 30 lM EDTA, 10% glycerol, T = °C (B) Average changes in steady state fluorescence anisotropy between 60% Atto532–C160 RepA and, from left to right, 10% Atto532–C160 RepA, 60% Atto532–C160 RepA in the presence of lM 1IR, 1–4 lM 1DR-long and 8–16 lM 1DR-short Conditions: 0.15 M (NH4)2SO4, 15 mM NH4-acetate pH 6.0, 10 lM EDTA, 3% glycerol, T = °C In the experiments with DNA, [RepA] = 15 nm tion of 1IR and 1DR-short causes a small decrease in anisotropy which may be related to enhanced homoFRET caused by slight rearrangement of the monomers in the RepA dimers or by minor aggregation Thus, RepA is dimeric free in solution and when bound to its operator sequence, but also when bound to 1DR-short In the presence of excess 1DR, monomerization of RepA takes place Discussion One of the striking properties of RepA is that it is able to recognize two types of DNA sequence, either the operator – with inverted repeats – or the iteron, in which the same bp sequence half-site found in the operator is specifically recognized Upon binding to the operator, RepA remains dimeric; it thus retains its symmetry matching the inverted repeats of the oligonucleotide When this symmetry is absent, i.e for the FEBS Journal 275 (2008) 5393–5407 ª 2008 The Authors Journal compilation ª 2008 FEBS R E M Diederix et al iteron sequence, RepA binds as a monomer instead of a dimer When operator or iteron DNA is added to AEDANS C160–RepA, discrete complexes are formed (Fig 3), characterized by higher AEDANS fluorescence anisotropy values and decreased apparent TrpAEDANS FRET (see below) RepA binds operator DNA (1IR) with a clear stoichiometry of : 1, i.e the protein binds as a dimer With the iteron sequence 1DR, which includes an additional stretch of bases (see Table 1), a stoichiometry of : is found, i.e monomer binding When the number of bases flanking the iteron sequence is considerably shorter, as with 1DR-short, the binding affinity is significantly decreased (2.9 lm), and nears that of non-specific DNA [6] Still, a discrete complex is formed in this case, as corroborated by fluorescence anisotropy decay measurements Fluorescence anisotropy decay analysis is a potent method to obtain information on the local and global dynamics of species in solution Here, it is used to characterize the discrete species discussed above For free RepA and RepA in complex with 1IR or 1DR, experiments were performed with AEDANS The analysis, summarized in Table 3, yields global rotational correlation times for free RepA and the complex with 1IR corresponding to species involving dimeric RepA, as expected In the case of the complex with 1DR, a fair correlation is also found between the experimental and calculated global rotational correlation times For the latter, the hydropro program was employed, which is able to extract hydrodynamic parameters using the protein’s atomic co-ordinates [17] A homology model based on the RepE–iteron structure was used as input Note that the bending angle of the 1DR as observed by EMSA (52°) is significantly larger than in the crystal structure (20°) which was used for the homology model [6,7] Furthermore, the crystal structure has a much shorter DNA oligonucleotide than the 1DR sequence: the latter is $ 3–4 times longer than the protein itself and may thus form a source of significant flexibility, difficult to account for in model building However, using the same structure as a basis to construct a potential complex between 1DR-short and monomeric RepA is not realistic The observed global rotational correlation time for the RepA–1DR-short complex cannot conceivably be justified assuming a complex similar to the RepE–iteron complex However, the purported dimeric RepA–1DR-short complex fits very well into the linear correlation between oligonucleotide size and experimental correlation time for the complexes involving dimeric RepA The complex Fluorescence studies of RepA between 1DR and RepA does not fit this correlation, in line with the fact that it involves monomeric RepA It is thus tempting to assume that dimeric RepA is actually involved in binding the 1DR-short sequence, despite the fact that it contains the full 22 bp iteron This last conclusion is corroborated by the observation that inter-monomeric homoFRET is observable with 1DR-short, but not 1DR (Fig 6) That dimerbinding to iterons is, in principle, possible has previously been shown by us According to EMSAs carried out under crowded conditions, a fraction of RepA dimers was observed to bind to the 1DR sequence [6] This fraction is obviously much larger in the case of 1DR-short, and the extra bases on the longer, monomer-binding, oligonucleotide 1DR seem to play a role in aiding monomerization The presence of bases downstream of the iteron sequence has also previously been shown to promote binding of Rep to pSC101 [18] The related replication initiator protein p from R6K plasmid is a well-documented case where not only monomers, but also dimers, are known to bind to the iteron [19] Interestingly, dimers of p protein occupy a much shorter stretch of the iteron sequence than monomers; whereas almost the entire 22 bp iteron sequence is occupied by the p monomers, only half of this – notably including the specific bp recognition sequence (repeat) – is occupied when dimeric p protein is bound [19] This may occur here as well As there is only one half of the inverted repeat of the operator sequence present in 1DR-short, it is likely that only one of two WH2 DNA-binding domains in RepA dimers is involved in binding This also makes sense energetically, the RepA dimer binds operator DNA with Kd = 0.7 nm i.e DG = )21.2 kJỈmol)1 [6] Subtracting from this a penalty of $ 7.8 kJỈmol)1 for the DNA bending [20] induced by dimer binding (61°), a free energy of ()21.2 to 7.8) ⁄ = )14.5 kJỈmol)1 is expected for binding of a single DNA-binding domain without the need to force DNA bending This translates to Kd = 1.3 lm, which is reasonably close to the value of 2.9 lm observed here for 1DR-short It is clear that in vitro, the effect of decreased length of the iteron flanking sequence is to weaken the iteronbinding affinity of monomeric RepA so that, at high [RepA], dimer binding occurs In vivo, this effect may be comparable in the sense that monomer-binding affinity is attenuated by the length or identity of the flanking sequence It is well established that Rep protein dimers not act as initiators in plasmid replication [21] A positive effect on monomer-binding affinity thus provides a way of selecting against dimer binding, favoring monomer binding and thus FEBS Journal 275 (2008) 5393–5407 ª 2008 The Authors Journal compilation ª 2008 FEBS 5401 Fluorescence studies of RepA R E M Diederix et al initiation It should be mentioned that the four iterons in pPS10 are contiguous, thus limiting the degree to which the flanking sequences may contribute to binding In other replicons, however, there are spacer sequences between the iterons, which in addition may have some sequence variability [22] It would be interesting to see whether our findings for RepA can be extrapolated to other Rep proteins An attractive feature of using AEDANS as an extrinsic label is that, besides its use to analyze the global rotational correlation times of macromolecules, it is useful as a FRET acceptor for intrinsic Trp residues RepA fortunately has only one Trp, making this use of the AEDANS probe more meaningful and helping interpretation of the FRET in terms of distances between the two fluorophores Moreover, W94 and C160 are located on the dimerization and DNA-binding domains of RepA, respectively, allowing us to interpret any observed changes in FRET in terms of relative movements between the two domains It emerges that the average estimated distance ˚ between the C160–AEDANS and W94 is $ 22 A in the free RepA dimer, and this distance decreases by a few angstroms upon binding either the 1IR, or 1DRshort oligonucleotides and increases slightly upon monomerization and binding to 1DR (see Table 2) The average distance observed in the complex with 1DR is in very good agreement with the value measured between the Cb atoms of residues W94 and C160 in the homology model of RepA, supporting the estimated value It is interesting that within the error, the distance between the AEDANS and indole moieties apparently does not change significantly between unbound RepA and RepA bound to either 1IR (as a dimer with both DNA-binding domains involved in binding), or 1DR-short (as a dimer, but presumably with only one domain involved), or indeed when bound as a monomer to 1DR This suggests that binding to both inverted half-repeats, as in the operator sequence, does not trigger large conformational rearrangements with respect to the free dimeric protein or to the dimer purportedly bound via one DNA-binding domain (1DR-short) Although significant structural rearrangements of RepA occur upon monomerization [3–5], these not appear to grossly alter the relative orientation of the two domains with respect to each other Naturally, it should be noted that manifold relative orientations of the two domains may exist, satisfying the observed distance, but which are still significantly different We are currently working towards a more comprehensive understanding of interdomain orientations using FRET 5402 Experimental procedures Cloning, expression and purification of wild-type RepA and C160–RepA In all cases, the concentration of protein is expressed in monomer units What is referred to as wild-type RepA is the His6-tagged variant of RepA, which was expressed and purified as described previously [4] This protein is indistinguishable from that without His-tag, except that it has a higher solubility [3,4] It was therefore used without subsequent removal of the tag C160–RepA also has the His6-tag and is a single-cysteine variant of wild-type RepA in which two of the three wild-type Cys residues (C29, C106) have been successively replaced by Ser using the PCR-based QuickChange Kit (Stratagene, Cedar Creek, CA, USA) Mutations were verified by sequencing C160–RepA was expressed as wild-type RepA [4] but almost all C160–RepA was present in the form of insoluble aggregates The protein was isolated by solubilization of the inclusion bodies and purification by Ni(II)-affinity chromatography under denaturing conditions, similarly as described previously [4] This results in pure protein, exhibiting a single band on SDS ⁄ PAGE After purification, the protein was reduced by addition of mm 2-mercaptoethanol and exchanged to unfolding buffer (5.6 m guanidinium hydrochloride, 0.56 m (NH4)2SO4, 0.2 m NH4-acetate, 0.2 mm EDTA, 1.2% Chaps, pH 6.0) Immediate refolding is achieved by fast 20-fold dilution in 0.15 m (NH4)2SO4, 15 mm NH4-acetate, 0.03 mm EDTA, 3% glycerol, pH 6.0 A small amount of precipitate generated by the refolding procedure was spun down at 14 000 g for 20 The latter buffer was used both for storage ()80 °C) and experiments Protein labeling C160–RepA was labeled with IAEDANS (Molecular Probes, Leiden, The Netherlands) under denaturing conditions, as follows C160–RepA was concentrated to $ 150 lm in unfolding buffer by ultrafiltration (10 kDa cut-off) A small amount of m Tris ⁄ HCl (pH 8.5) was added to increase the pH to 7.2, and Tris(2-carboxyethyl) phosphine to keep the single Cys reduced (1 mm) The end volume was 1.6 mL IAEDANS was dissolved (40 mm) in unfolding buffer and quickly mixed with the reduced protein to a final concentration of mm The reaction was allowed to proceed for h at room temperature, and then 12.3 mg of glutathione was added to quench the reaction The reaction mixture was exchanged for fresh unfolding buffer by extensive ultrafiltration The labeling efficiency was close to 50%, as judged from UV ⁄ Vis spectroscopy AEDANS C160–RepA was refolded in the same way as unlabeled RepA Similarly, C160–RepA was labeled with maleimide Atto532 (Atto-Tec, Siegen, Germany), with 60% labeling efficiency Here, the degree of labeling in the folded FEBS Journal 275 (2008) 5393–5407 ª 2008 The Authors Journal compilation ª 2008 FEBS R E M Diederix et al protein was important and varied from 60% to 10% by mixing the appropriate amounts of Atto532-labeled and unlabeled C160–RepA before refolding Correct refolding of Atto532 C160–RepA was confirmed by determining its binding efficiency and stoichiometry to Alexa647-labeled 1IR, using FRET (not shown) DNA purification and labeling 1DR (Table 1) was prepared as described previously [3–6] The 1IR and 1DR-short duplexes were prepared by annealing their constituent complementary strands (Sigma-Genosys, Cambridge, UK) in equimolar amounts The duplexes were purified using a MA7 column (BioRad, Hercules, CA, USA) followed by desalting using C18 Sep-PaK columns (Waters, Milford, MA, USA) Also, 5¢-amine modified variants of the 1IR and 1DR-short oligonucleotides were first reacted with NHS-Alexa568 and Alexa647 (Molecular Probes) respectively, according to the manufacturer’s instructions Unreacted Alexa568 or Alexa647, and DNA, were removed by chromatography (MA7), and the oligonucleotides were subsequently annealed with their complementary strands, and purified as above UV ⁄ Vis and CD spectroscopy CD spectra and melting curves of wild-type RepA and AEDANS-labeled, as well as unlabeled, C160–RepA were recorded as described elsewhere [3] Protein concentrations were 2.5–5 lm, and the optical path length was 0.1 cm Room temperature UV ⁄ Vis spectra were recorded using a Cary 3E UV ⁄ Vis spectrophotometer with cm path-length cuvettes Size-exclusion chromatography Gel-filtration assays were performed at room temperature, with a Superdex HR 10 ⁄ 30 column (Amersham Biosciences, Freiburg, Germany) Sample volumes were 50 lL and protein concentrations $ lm Steady-state fluorescence spectroscopy Fluorescence measurements were performed with an ISS PC1 or an SLM 8000D photon counting spectrofluorimeter at or 23.5 °C using · mm path-length quartz cuvettes (Starna, Hainault, UK) Spectra were recorded using magic angle conditions (Glan-Taylor polarizers, excitation polarizer vertical, emission polarizer 54.7º to vertical) Bandwidths were nm (excitation) and 10 nm (emission) Steady-state emission anisotropies of fluorescent probes were measured using Glan-Taylor polarizers, as described previously [6] For AEDANS C160–RepA, FRET between the single Trp residue, (W94, the donor) and AEDANS (the acceptor) Fluorescence studies of RepA was quantified by the (ratio)A approach [23,24] using excitation spectra measured at an emission wavelength where W94 fluorescence does not contribute (480 nm) This permits several simplifications, among them disregard of uncertainties in the degree of labeling Because of significant static quenching of the W94 donor upon complex formation, a correction was necessary to account for the fraction (d+) of fluorescent donor remaining The FRET efficiency was calculated as follows:  ! !  AEDANS  F280 nm eAEDANS e À 280 nm 340 nm 1ị Eẳ ỵ d F340 nm eW94nm eAEDANS 280 340 nm The AEDANS fluorescence intensity for 340 nm excitation (F340 nm) arises from direct excitation of the acceptor It is independent of FRET and depends only on the acceptor concentration The extinction coefficients of W94 [25] and AEDANS [26] are assumed constant as a function of labeling and ⁄ or protein complexation state Because of relatively low signal intensity for 295 nm excitation caused by the poor solubility of RepA, we measured it for 280 nm excitation instead The disadvantage of this is that Tyr absorbs at 280 nm, and thus may, in principle, contribute to the measured intensities This would occur, because no Tyr emission is observed, via FRET to W94 followed by W94 fi AEDANS energy transfer The FRET efficiencies determined using Eqn (1) thus strictly represent an upper limit for the W94 fi AEDANS process However, the ˚ closest Tyr–W94 distance is $ 15 A, based on the crystal structure [5], and thus any contribution of Tyr was not taken into account in the efficiency calculations The W94 fluorescence decay could be analyzed assuming three discrete lifetime components with $ 80% of the fluorescence from one of these species (s $ ns) (see below and Table S1) The decay can thus be approximated as monoexponential, allowing use of simple FRET theory ˚ The Forster radius, R0, in A, for the W94AEDANS ă pair was calculated as follows: R0 ¼ 0:211ðnÀ4 Á UD Á j2 Á JÞ1=6 ð2Þ Here, n is the refractive index, with a value of 1.4 The orientation factor j2 depends on the relative orientation of the donor emission and acceptor absorption transition moments with respect to the donor–acceptor vector As a first approximation, a value of ⁄ for j2 was taken in this work, equivalent to assuming rapid isotropic averaging [27] J is the overlap integral between Trp emission and AEDANS absorbance spectra (expressed as nm4Ỉm)1cm)1) FD is the fluorescence quantum yield of W94 (FW94) This was determined for unlabeled C160–RepA by calibration with a NATA solution in 10 mm sodium phosphate, pH 7.0, using FNATA = 0.14 [28] The apparent value for FW94 in the various DNA complexes was derived from the value of free RepA by comparing the fluorescence intensity FEBS Journal 275 (2008) 5393–5407 ª 2008 The Authors Journal compilation ª 2008 FEBS 5403 Fluorescence studies of RepA R E M Diederix et al (kex = 295 nm) of free and bound unlabeled C160–RepA and wild-type RepA, with the protein concentration constant The ratio of the apparent FW94 of bound and free RepA was then used to estimate the fraction of fluorescent donor remaining upon DNA complexation (d+ in Eqn 1) For the R0 calculation we used the quantum yield determined for free C160–RepA (see above) Titrations were performed with increasing amounts of DNA added to AEDANS C160–RepA, or with increasing amounts of wild-type RepA or unlabeled C160–RepA added to Alexa568–1IR Fresh solutions were prepared for each data point, and equilibrated 10 before measuring Binding curves were fit to the quadratic expression given in Eqn (3) for the amount of RepA–DNA complex, with [RepA] divided by the expected stoichiometry n The amount of bound protein or DNA is related to the instrument signal (AEDANS C160–RepA anisotropy and the fluorescence intensity ratio for excitation at 280 and 340 nm F280 nm ⁄ F340 nm) via the corresponding signals Sbound and Sfree (Eqn 4) or the equivalent for bound and free Alexa568–1IR (Eqn 5) ½RepA À DNA  q 3aị  ẳ 0:5 c À c2 À  ½RepAŠT n  ẵDNAT  c ẳ Kd ỵ ẵRepAT n ỵ ẵDNAT signal AEDANS ! ẵRepA DNA  Sbound ẳ ẵRepAT n ! ẵRepA DNA  Sfree ỵ ẵRepAT n signalAlexa568 ẳ  ẵRepA DNA Sbound ẵDNAT   ẵRepA DNA Sfree ỵ ẵDNAT 3bị 4ị  Rtị ẳ 5ị The Forster distance R0(2/3) for Atto532Atto532 homoă FRET was calculated using Eqn (2), with FD (FAtto532) = 0.9 and calculating the overlap integral between Atto532 emission and excitation using spectral data provided by the manufacturer Time-resolved fluorescence spectroscopy Time-resolved fluorescence intensity and depolarization measurements were made using the time-correlated single- 5404 photon counting technique, using the set-up described previously [29] For Trp fluorescence depolarization measurements (kex = 297 nm, kem = 345 nm), the excitation light source was a Ti : sapphire picosecond laser (Tsunami, Spectra Physics, Mountain View, CA, USA), pumped with a W Nd : YVO4 diode laser (Millennia, Spectra Physics), and associated with a third harmonic generator The pulses had 1–2 ps width and a repetition rate of 0.8–4 MHz, with an average power of 20 lW reaching the cuvette The temperature of the sample was thermostated at °C for increased sample stability, required because of long acquisition times The timing calibration was 6.1 ps per channel, with 4096 data channels Typical polarized decay curves had $ 4000–10 000 counts in the peak (1–2 · 106 total photons) The quality of photon counting statistics in the Trp experiments was limited by the sample stability For AEDANS measurements (T = 23.5 °C; kex = 375 nm, kem = 480 and 530 nm), the excitation light source was an LDH-P-C-375 diode laser head (PicoQuant, Berlin, Germany), operating at 375 nm with a MHz repetition rate, with on average 75 lW power reaching the cuvette The fluorescence was collected in the plane perpendicular to the vertical orientation of the linearly polarized excitation, and at 90º to the excitation beam and focused on a monochromator (f = 100 mm, 16 nm bandwidth) through a cut-off filter, a Polaroid HNP’B polarizer and a quartz depolarizer (Acton Research Corp., Acton, MA, USA) The total fluorescence intensity decay, I(t) = Im(t), was measured with the emission polarizer set at 54.7°, relative to the vertically polarized excitation beam The two emission components, polarized parallel IVV(t) and perpendicular IVH(t) to the plane of polarization of the excitation beam, were recorded sequentially by alternating the orientation of the emission polarizer every 10–20 The timing calibration was 48.8 ps per channel, with 4096 data channels Typical polarized decay curves had $ 6000–40 000 counts in the peak (2–11 · 106 total photons).The experimental anisotropy decay R(t) is related to the experimental emission decay of the polarized components by: ẵIVV tị GIVH tị ẵIVV tị ỵ 2GIVH ðtފ ð6Þ In this instrumental setup, G is a scaling factor which is independent of the emission wavelength and, in general, has values near It takes in account small instabilities of the laser and ⁄ or differences in accumulation times for the two polarized intensities It was determined by correlating the steady-state anisotropy value measured separately, to the anisotropy value resulting from integration of the IVV(t) and IVH(t) traces Decays Im(t) were fit to a sum of n exponential functions (n = 1, 2) by iterative convolution, using nonlinear global least-squares methods from the program globals unlimited (Urbana, IL, USA) [30] The emission FEBS Journal 275 (2008) 5393–5407 ª 2008 The Authors Journal compilation ª 2008 FEBS R E M Diederix et al Fluorescence studies of RepA anisotropy function, r(t) was determined by global analysis of the two polarized components of the fluorescence intensity, IVV(t) and IVH(t), as well as Im(t), using the same routines The analysis consisted in finding the r(t) numerical parameters that best fit the two polarized decay functions and iVH(t) = [i(t) ⁄ 3][1)r(t)], iVV(t) = [i(t) ⁄ 3][1+2r(t)] where i(t) = iVV(t)+2iVH(t) = 3im(t), to the experimental traces Lower case letters refer to the mathematical functions, whereas upper case letters refer to the convoluted experimental data and resulting fits In the case of AEDANS, individual analysis of 480 and 530 nm decays gave very similar lifetime values Therefore, anisotropy data from two emission wavelengths (480 and 530 nm) were fitted simultaneously, linking the corresponding component lifetimes to get a better defined set of fitting parameters The adequacy of the analyses was determined from the reduced weighted sum of squares of residuals, and visual inspection of the distribution of weighted residuals The general expression used for the emission anisotropy parameters in the fits is given by Eqn (7), in which /i are correlation times, and the pre-exponential factors bi are normalized In all the AEDANS anisotropy experiments the time zero anisotropy, r0, from the fit had an average value of 0.31 ± 0.015 In the case of Trp anisotropy analysis, r0 was kept fixed at a reasonable value, 0.28 [31], to suppress the contribution of otherwise unaccounted for scatter to useful parameters rtị ẳ r0 X bi exp½Àt=/i Š for comparative purposes The global rotational correlation times for the F plasmid RepE–operator complex were calculated in the same way, using its recently published crystal structure [9] Note that the correlation time actually measured depends on the orientation of the AEDANS absorption and emission transition dipoles with respect to the protein axes For a case where any of these are parallel to the long axis, the value takes the maximum limit For the other extreme, i.e transition dipoles perpendicular to the protein long axis, the correlation time takes a value slightly higher than the minimum value [32] For free RepA and the complex RepA–1IR, no (homologous) structures are available A prolate ellipsoid shape was assigned to both structures, with dimensions based on friction coefficients determined previously by sedimentation velocity experiments (f ⁄ f0 = 1.2 for free and 1IR-bound RepA), in which special care was taken regarding complex dissociation during the analytical ultracentrifugation [3] This value implies a ratio of long versus short axes of in the prolate ellipsoid, giving an observed average global rotational correlation time that is 1.6–3.4 times higher than expected if the molecule were spherical, depending on probe orientation [33,34] Just like for the cases where correlation times were calculated using hydropro, the limiting minimum and maximum rotational correlation times have been included in Table Partial specific volumes used for protein, DNA and hydration were 0.703, 0.55 and 0.28 mLặg)1, respectively 7ị Estimates of the global rotational correlation times for free dimeric RepA, and the complexes with 1IR, 1DR and 1DR-short were calculated in one of two ways: based on the 3D structure and based on a prolate ellipsoid shape with dimensions derived from sedimentation velocity experiments [3] In the case of iteron complexes, a crystal structure of the homologous protein F plasmid RepE in complex with its cognate iteron sequence is available [7] Using this, we built an homology model of the RepA–1DR structure [2,10] and simply elongated the DNA sequence assuming rigid DNA with no additional bending, to construct the complexes with 1DR and 1DR-short Note that the elongation is symmetric for 1DR-short, but asymmetric in the case of 1DR We did not account for the much stronger DNA bending observed in EMSA experiments (52°) than in the crystal structure (20°) [6] The structures thus obtained were used as input to the hydropro program, which calculates hydrodynamic parameters on the basis of atomic co-ordinates [17] Other input factors included the specific protein volume (0.702 mLỈg)1), solution density (1.019 gỈmL)1) and solvent viscosity (1.096 cP) The latter value was determined for the buffer at 23.5 °C using a capillary viscosimeter (Schott, Mainz, Germany) The lowest and highest of the five output global rotational correlation times for each geometry are included in Table Acknowledgements We thank Guillermo Bernabeu for excellent technical assistance and Dr Silvia Zorrilla for helpful discussions Financial support was from the Spanish Ministry for Education and Science (MEC grant nos.: BMC2003-00088 (to RG and MPL), BFU2006-00494 (to RG), and BFU2006-0395 ⁄ BMC (REMD and MPL)) REMD is supported by a ‘Juan de la Cierva’ fellowship (MEC grant no.: JCI-2005-1721-2) References Giraldo R (2003) Common domains in the initiators of DNA replication in Bacteria, Archaea and Eukarya: combined structural, functional and phylogenetic perspectives FEMS Microbiol Rev 26, 533–554 ´ Giraldo R & Fernandez-Tresguerres ME (2004) Twenty years of the pPS10 replicon: insights on the molecular mechanism for the activation of DNA replication in iteron-containing bacterial plasmids Plasmid 52, 69–83 ´ ´ Dı´ az-Lopez T, Lages-Gonzalo M, Serrano-Lopez A, Alfonso C, Rivas G, Dı´ az-Orejas R & Giraldo R (2003) Structural changes in RepA, a plasmid replication FEBS Journal 275 (2008) 5393–5407 ª 2008 The Authors Journal compilation ª 2008 FEBS 5405 Fluorescence studies of RepA 10 11 12 13 14 15 R E M Diederix et al inititator, upon binding to origin DNA J Biol Chem 278, 18606–18616 Giraldo R, Andreu JM & Dı´ az-Orejas R (1998) Protein domains and conformational changes in the activation of RepA, a DNA replication initiator EMBO J 17, 4511–4526 ´ Giraldo R, Fernandez-Tornero C, Evans PR, Dı´ azOrejas R & Romero A (2003) A conformational switch between transciptional repression and replication initiation in the RepA dimerization domain Nat Struct Biol 10, 565–571 ´ ´ Dı´ az-Lopez T, Davila-Fajardo C, Blaesing F, Lillo MP & Giraldo R (2006) Early events in the binding of the pPS10 replication protein RepA to single iteron and operator DNA sequences J Mol Biol 364, 909–920 Komori H, Matsunaga F, Higuchi Y, Ishiai M, Wada C & Miki K (1999) Crystal structure of a prokaryotic ˚ replication initiator bound to DNA at 2.6 A resolution EMBO J 18, 4597–4607 Swan MK, Bastia D & Davies C (2006) Crystal structure of p initiator protein–iteron complex of plasmid R6K: implication for initiation of plasmid DNA replication Proc Natl Acad Sci USA 103, 18481– 18486 Nakamura A, Wada C & Miki K (2007) Structural basis for regulation of bifunctional roles in replication initiator protein Proc Natl Acad Sci USA 104, 18484– 18489 Urh M, Wu J, Wu J, Forest K, Inman RB & Filutowicz M (1998) Assemblies of replication initiation protein on symmetric and asymmetric DNA sequences depend on multiple protein oligomerisation surfaces J Mol Biol 283, 619–631 Park K, Han E, Paulsson J & Chattoraj DK (2001) Origin pairing (‘handcuffing’) as a mode of negative control of P1 plasmid copy number EMBO J 20, 7323–7332 Paulsson J & Chattoraj DK (2006) Origin inactivation in bacterial DNA replication control Mol Microbiol 61, 9–15 Callis PR & Vivian JT (2003) Understanding the variable fluorescence quantum yield of tryptophan in proteins using QM–MM simulations Quenching by charge transfer to the peptide backbone Chem Phys Lett 369, 409–414 Dale RE, Eisinger J & Blumberg WE (1979) The orientational freedom of molecular probes The orientational factor in intramolecular energy transfer Biophys J 26, 161–194 Lillo MP, Beechem JM, Szpikowska BK, Sherman MA & Mas MT (1997) Design and characterization of a multisite fluorescence energy-transfer system for protein folding studies: a steady-state and time-resolved study of yeast phosphoglycerate kinase Biochemistry 36, 11261–11272 5406 16 Weber G & Shinitzky M (1970) Failure of energy transfer between identical aromatic molecules on excitation at the long wave edge of the absorption spectrum Proc Natl Acad Sci USA 65, 823–830 17 Garcia de la Torre J, Huertas ML & Carrasco B (2000) Calculation of hydrodynamic properties of globular proteins from their atomic-level structure Biophys J 78, 719–730 18 Fueki T, Sugiura S & Yamaguchi K (1996) Open strands to iterons promote the binding of the replication initiator protein (Rep) of pSC101 to the unit sequence of the iterons in vitro Biochim Biophys Acta 1305, 181– 188 19 Kunnimalaiyaan S, Kruger R, Ross W, Rakowski SA ă & Filutowicz M (2004) Binding modes of the initiator and inhibitor forms of the replication protein p to the c ori iteron of plasmid R6K J Biol Chem 279, 41058– 41066 20 Williams LD & Maher LJ III (2000) Electrostatic mechanisms of DNA deformation Annu Rev Biophys Biomol Struct 29, 497–521 21 Kruger R, Rakowski SA & Filutowicz M (2004) Particiă pating elements in the replication of iteron containing plasmids In Plasmid Biology (Funnell BE & Phillips GJ, eds), pp 25–45 ASM Press, Washington DC 22 Chattoraj DK (2000) Control of plasmid DNA replication by iterons: no longer paradoxical Mol Microbiol 37, 467–476 23 Clegg RM (1992) Fluorescence resonance energy transfer and nucleic acids Methods Enzymol 211, 353–388 24 Majumdar ZK, Hickerson R, Noller HF & Clegg RM (2005) Measurements of internal distance changes of the 30S ribosome using FRET with multiple donor–acceptor pairs: quantitative spectroscopic methods J Mol Biol 351, 1123–1145 25 Demchenko AP (1986) Ultraviolet Spectroscopy of Proteins Springer-Verlag, Berlin 26 Hudson EN & Weber G (1973) Synthesis and characterization of two fluorescent sulfhydryl reagents Biochemistry 12, 4154–4161 27 van der Meer BW (2002) Kappa-squared: from nuisance to new sense J Biotechnol 82, 181–196 28 Szabo AG & Rayner DM (1980) Fluorescence decay of tryptophan conformers in aqueous solution J Am Chem Soc 102, 554–563 29 Lillo MP, Canadas O, Dale RE & Acuna AU (2002) ˜ ˜ Location and properties of the taxol binding center in microtubules: a picosecond laser study with fluorescent taxoids Biochemistry 41, 12436–12449 30 Beechem JM, Gratton E, Ameloot M, Knutson JR & Brand L (1991) The global analysis of fluorescence intensity and anisotropy decay data: second generation theory and programs In Topics in Fluorescence Spectroscopy, Vol (Lakowicz JR, ed.), pp 241–305 Plenum Press, New York, NY FEBS Journal 275 (2008) 5393–5407 ª 2008 The Authors Journal compilation ª 2008 FEBS R E M Diederix et al 31 Ruggiero AJ, Todd DC & Fleming GR (1990) Subpicosecond fluorescence anisotropy studies of tryptophan in water J Am Chem Soc 112, 1003–1014 32 Lakowicz JR (1999) Principles of Fluorescence Spectroscopy, 2nd edn Kluwer Academic, New York, NY 33 Zorrilla S, Rivas G & Lillo MP (2004) Fluorescence anisotropy as a probe to study tracer proteins in crowded solutions J Mol Recog 17, 408–416 34 Waxman E, Laws WR, Laue TM, Nemerson Y & Ross JBA (1993) Human factor VIIa and its complex with soluble tissue factor: evaluation of asymmetry and conformational dynamics by ultracentrifugation and fluorescence anisotropy decay methods Biochemistry 32, 3005–3012 Fluorescence studies of RepA Fig S2 Anisotropy decays R(t) (kex = 375 nm, kem = 530 nm) of AEDANS C160–RepA free in solution (A) and bound to 1IR (B), 1DR (C) and 1DR-short (D) Table S1 Fluorescence lifetimes and decay amplitudes for W94 and AEDANS–C160, respectively, in free RepA and RepA bound to various cognate DNA sequences This supplementary material can be found in the online version of this article Please note: Wiley-Blackwell is not responsible for the content or functionality of any supplementary materials supplied by the authors Any queries (other than missing material) should be directed to the corresponding author for the article Supporting information The following supplementary material is available: Fig S1 Anisotropy decays R(t) (kex = 297 nm, kem = 345 nm) of wild-type RepA free in solution (A) and bound to 1IR (B), 1DR (C) FEBS Journal 275 (2008) 5393–5407 ª 2008 The Authors Journal compilation ª 2008 FEBS 5407 ... 295 nm) of free and bound unlabeled C160? ?RepA and wild-type RepA, with the protein concentration constant The ratio of the apparent FW94 of bound and free RepA was then used to estimate the fraction... DNA-binding domains, and rearrangement of the relative orientation of the two domains [7,9] The conformational change upon iteron binding may expose a recognition site for protein? ? ?protein interaction,.. .Fluorescence studies of RepA R E M Diederix et al Interestingly, in the latter case, the protein binds as a monomer [2–6] Free in solution, the protein is essentially dimeric,