Inactivation and destruction by KMnO4 of Escherichia coli RNA polymerase open transcription complex: recommendations for footprinting experiments

ANALYTICAL BIOCHEMISTRY Analytical Biochemistry 320 (2003) 239–251 www.elsevier.com/locate/yabio

Inactivation and destruction by KMnO4 of Escherichia coli RNA polymerase open transcription complex: recommendations for footprinting experiments
ski and Kazimierz L. Wierzchowski* Tomasz Łozin Institute of Biochemistry and Biophysics, Polish Academy of Sciences, Pawi
nskiego 5a, 02-106 Warszawa, Poland Received 17 March 2003

Abstract Potassium permanganate oxidation of pyrimidine residues in single-stranded DNA is commonly used in footprinting studies on formation of open transcription complex (RPo) by RNA polymerases (RNAP) at cognate promoters. Our own experience and literature search led us to conclude that KMnO4 doses often used in such studies might cause multiple-hit oxidation of promoter DNA and oxidative damage to RNAP in RPo and lead to false interpretation of footprints. We have therefore studied as a function of KMnO4 dose (i) transcription activity of RPo formed by Escherichia coli RNAP at a model cognate promoter Pa and (ii) RPoÕs structural integrity, by gel electrophoresis and footprinting assays. Kinetics of formation of this complex and melting of DNA in the transcription bubble region were thoroughly characterized by us previously. Here we show that (i) RPo becomes completely inactivated at oxidant doses much lower than those needed to cause a detectable footprint of the melted DNA region, (ii) footprinting patterns of the melted promoter region remain practically unaffected by RNAP oxidation within a range of low oxidant doses causing single-hit oxidation of DNA, and (iii) at higher oxidant doses, corresponding to multiple-hit DNA oxidation, the gross structure of RPo changes progressively until its complete collapse and dissociation into constituent components, so that only approximate interpretation of the footprinting data for the melted DNA region is possible. A protocol for accurate RPo footprinting with low single-hit KMnO4 doses and interpretation of the footprinting data in terms of kinetics of oxidation of pyrimidine residues in promoter DNA is recommended. Ó 2003 Elsevier Science (USA). All rights reserved. Keywords: Permanganate footprinting; Open transcription complex; Escherichia coli RNA polymerase; Promoter DNA; Kinetics of thymine oxidation; Thymine diglycol

Potassium permanganate is an extremely reactive and powerful agent able to oxidize most functional groups in organic molecules, including carbon–carbon double bonds [1,2]. In the field of nucleic acids research, of particular analytical interest is the oxidation of the 5,6 double bond in thymine and cytosine to corresponding diglycols, since reactivity of these bases has been shown to be distinctly faster in single- than in double-stranded DNA regions [3–5]. This finding has been widely exploited in permanganate footprinting studies on the

* Corresponding author. Tel.: +48-22-658-4729. E-mail address: [email protected] (K.L. Wierzchowski).

formation of RPo1 between RNA polymerases and DNA promoters (cf. [6–13] for representative work) involving separation of DNA strands in a so-called bubble region [14,15]. The commonly used footprinting conditions of RPo were based on earlier recommendations [7] concerning oxidant dosage, selected for semiquantitative evaluation of the content of open complexes formed in bacterial cells in vivo. Critical 1 Abbreviations used: ANS, aminonaphthalene sulfonate; BSA, bovine serum albumin; dsDNA, double-stranded DNA; DTT, dithiothreitol; EMSA, electrophoretic mobility shift assay; FDAI, fluorescence-detected abortive initiation; MH, multiple-hit; RNAP, RNA polymerase; RPo, open transcription complex; RPo,ox, oxidized open transcription complex; SH, single-hit; ssDNA, single-stranded DNA.

0003-2697/$ - see front matter Ó 2003 Elsevier Science (USA). All rights reserved. doi:10.1016/S0003-2697(03)00381-6

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evaluation of a large body of pertinent publications allowed us to conclude that KMnO4 doses (initial concentration of the oxidant
time of exposure) used in in vitro studies on RPo might often lead to MH oxidation of promoter DNA within its melted region and, consequently, to misinterpretation of the relative reactivity of pyrimidine bases therein. This is even more true since no attention has been paid to the reactivity of the RNAP component toward KMnO4 and the analytical consequences of oxidative damage to RPo structure and thermal stability. In these studies, footprinting experiments were usually carried out at a selected oxidant dose tacitly assumed to correspond to SH oxidation conditions. Only lately has it been shown how to evaluate quantitatively the reactivity of individual pyrimidines from footprints obtained under MH conditions [16], but even in this study protein oxidation was completely neglected. Recently, we have studied [17] the effect of Mg2þ ions on oxidation of thymines within the transcription bubble of RPo, formed at a model Pa promoter by Escherichia coli RNA polymerase, as a function of KMnO4 dose, and found that controlled use of SH oxidant doses allows reliable determination of the corresponding reaction rate constants. Here we present the results of further investigations on KMnO4 oxidation of this complex as a function of the oxidant dose and demonstrate that (i) RNAP inactivation occurs at doses well below those needed to obtain measurable SH footprint of the transcription bubble, (ii) progressive complex destruction due to RNAP oxidation takes place in the MH range of oxidant doses, and (iii) from footprints obtained under MH conditions only approximate kinetic data on oxidation of pyrimidine bases can be determined. On the basis of this and previous studies, we recommend a protocol for reliable KMnO4 footprinting of ssDNA regions in RPo and other DNA–protein complexes.

Materials and methods RNA polymerase RNA polymerase (EC 2.7.7.6) was prepared from E. coli C600 strain according to Burgess and Jendrisak [18]

except that Sephacryl S300 was used instead of Bio-Gel A5m, and the enzyme was stored in buffer S (50% glycerol, 100 mM NaCl, 10 mM Tris–HCl, pH 7.9, 0.1 mM DTT). Assessment of its activity according to Chamberlin et al. [19] showed that ca. 50% of the holoenzyme was active. Promoter E. coli promoter Pa, containing the consensus )35 and )10 hexamers separated by a 17-bp spacer (Fig. 1) was synthesized and cloned into pDS3 plasmid as described previously [20,21]. Reagents and chemicals c-ANS–UTP was prepared and purified according to Yarbrough et al. [22]. ANS was purchased from Fluka; UTP, ApA, and heparin were purchased from Sigma. All other chemicals were of reagent grade. FDAI assay of the activity of KMnO4 -oxidized RPo In this assay, c-ANS–UTP was used as the elongating NTP [23,24] and ApA as the initiating nucleotide, so that ApApUpU was the only abortive transcription product (cf. Fig. 1). The amount of fluorescent ANS– pyrophosphate liberated in the course of the reaction was measured spectrofluorimetrically at 500 nm (emission excited at 360 nm), using a laboratory-made double-monochromator ratio-recording and computercontrolled spectrofluorimeter, equipped with a thermoregulated cell compartment. A buffered solution of RNAP and a 226-bp-long DNA fragment of pDS3 plasmid carrying the investigated Pa promoter was aliquoted into 200-ll samples containing 1 pmol of DNA and 2 pmol of RNAP in buffer A (25 mM Tris–HCl, pH 7.9, 100 mM KCl, 0.2 mM EDTA, and, 10 mM MgCl2 when studying the effect of Mg2þ ions on oxidation of the complex). To form the open complex, samples were incubated for 10 min at 37 °C in a thermoregulated water bath. Subsequently 10 ll of a KMnO4 solution (freshly diluted from stock) was added to set its final concentration at ca (0.005–0.125 mM) and the reaction mixture was incubated at the same temperature for a time period t

Fig. 1. Sequence of the synthetic promoter Pa cloned into pDS3 plasmid. The melted DNA region of RPo is in bold face, the kinked arrow indicates transcription start site, and straight arrows indicate direction of extension of end-labeled primers Pr(t) and Pr(nt) by the Klenow enzyme (in brackets location relative to the start site).

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necessary to reach the desired oxidant dose c
t of up to 0.02 (M
s). The reaction was then quenched by addition of 10 ll of 20 mM DTT solution and supplemented with 20 ll of equimolar solution of ApA and ANS–UTP in buffer A (containing 10 mM MgCl2 ) to the final concentration of 0.1 mM. Immediately afterward, each sample was transferred into a fluorometric cuvette maintained at 37 °C in the thermoregulated cell compartment of the spectrofluorimeter and incubated for 3 min, and the progress of steady state abortive reaction was continuously monitored for 15 min. In control experiments, free RNAP holoenzyme was first oxidized at 0 °C and combined with promoter DNA at 37 °C, and its activity was measured according to the FDAI protocol. EMSA of KMnO4 -oxidized RPo The open complex was formed by incubation for 10 min at 37 °C of the mixture of 32 P-end-labeled DNA fragment carrying the promoter Pa (5 nM) and RNAP (10 nM) in buffer A (cf. above), with 10 mM MgCl2 when studying the effect of Mg2þ ions. An appropriate amount of KMnO4 stock solution was added to obtain the desired concentration of the oxidant and at selected time periods 10-ll aliquots were withdrawn and mixed with 3 ll of 20 mM DTT to quench the oxidation reaction. To estimate the stability of the oxidized complex in solution, the reaction was quenched in the total volume after application of a desired oxidant dose, incubated at 37 or 47 °C and sampled for EMSA at various times. To each 13-ll reaction sample 3 ll of gel loading buffer (25% sucrose, 0.02% bromophenol blue, 0.005% heparin) was added, the samples were loaded onto a native 4% polyacrylamide gel and resolved by electrophoresis in TB–Mg buffer (90 mM Tris–borate, pH 8.3, and 5 mM MgCl2 ). During electrophoresis current intensity was regulated to maintain the desired temperature inside the gel in the range of 35–42 °C. Quantitative KMnO4 footprinting of RPo Modification of nucleotides in DNA by KMnO4 oxidation as a function of the oxidant dose and their detection by primer extension were performed as described earlier [17]. A buffered solution of RNAP and supercoiled plasmid pDS3 carrying the promoter Pa was divided into two parts, one of which was made 10 mM in MgCl2 and the other one supplemented with water; each solution was then aliquoted into 50-ll samples containing 1 pmol of pDS3 and 2 pmol of RNAP in buffer A. To form the open complex each sample was placed at constant time intervals in a water bath at 37 °C for 15 min. Then 5 ll of an appropriately freshly diluted KMnO4 stock was added to obtain 0.05–1.6 mM final concentration of the oxidant, and the reaction mixture

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was incubated for 3 min at 37 °C. The reaction was quenched by addition of 150 ll of the stop solution (2 mM EDTA, 1% b-mercaptoethanol, and 5 nM marker DNA—the SalI–HindIII fragment of the same plasmid). The samples were deproteinized by phenol/ chloroform extraction, the aqueous phase was collected, and the oxidized DNA was purified using Sephadex G50 spin columns. The purified samples were then analyzed for the degree of oxidation of thymine residues in the nontemplate and template DNA strands within the transcription bubble by dividing them into halves and performing primer extension of each with an appropriate 32 P-endlabeled primer, Pr(nt) or Pr(t) (cf. Fig. 1). Reaction conditions were optimized for each strand separately to ensure the specificity of primersÕ hybridization. For the nontemplate strand, 35 ll of purified DNA was mixed with 4 ll 0.01 M NaOH containing 2 pmol of 32 P-endlabeled primer and denatured at 80 °C for 3 min. After cooling to 0 °C the mixture was neutralized with 8 ll of the Klenow reaction buffer (250 mM Tris–HCl, pH 7.2, 50 mM MgSO4 , 1 mM DTT, and 2.5 mM each dNTPs) and incubated at 45 °C for 15 min to form a Pr(nt)– DNA complex. Then 2 units of the Klenow fragment of DNA polymerase I in 2 ll of enzyme diluent (25% glycerol, 25 mM KH2 PO4 , pH 7.0, 1 mM DTT, and 50 mg/ml BSA) were added and the primer extension reaction was carried out at 45 °C for 10 min, followed by addition of 18 ll of stop solution (3 M ammonium acetate, 20 mM EDTA, and 2 lg tRNA). Duplex DNA products of the Klenow reaction were ethanol precipitated, rinsed with 70% ethanol, dried, dissolved in 10 ll of gel loading buffer (70% freshly deionized formamide, 7 M urea, 3 mM NaOH, 0.1 mM EDTA, and 0.02% bromphenyl blue and xylene cyanole), and denatured by heating at 95 °C for 2 min, 3-ll samples of ssDNA were resolved in duplicate on a 6% polyacrylamide sequencing gel in TBE buffer (0.089 M Tris–borate and 2 mM EDTA, pH 8.3). Dried gels were exposed to storage phosphor screens from Molecular Dynamics. For the template strand, all the reaction conditions were the same except that neutralization, hybridization, and primer extension steps were carried out at 60 °C, and cooling after denaturing was omitted. All footprinting experiments were performed in duplicate and each set of data was subject to two independent quantification procedures. Images of footprints were obtained with the use of a Molecular Dynamics Phosphorimager. Integrated intensities of bands (or groups of bands) and their intensity profiles along gel lanes were determined using the ImageQuant software, by the volume integration and area integration options, respectively. For area integration, the lowest intensity point in the graph was used as the horizontal baseline. In volume integration, local background was used for bands corresponding to

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marker DNA and promoter bubble DNA fragments, whereas for the whole lanes that of gel areas outside the lanes was used. These data were analyzed further with Origin (MicroCal) and Quattro Pro 4.0 (Borland) softwares. Estimation of an effective KMnO4 dose On account of the presence in footprinting reactions of oxidizable buffer A components such as Tris and EDTA, the redox capacity of the buffer solution with respect to KMnO4 , in the absence of RPo, was estimated by determination of an effective pseudo-first-order rate constant for KMnO4 reduction at 37 °C. This was calculated from single exponential decrease in time of the oxidant absorbance at 525 nm [2] for several of its initial concentrations, corresponding to the doses used in footprinting experiments. The resultant rate constant of ca. 0.1 M1 s1 was used to calculate the effective dose delivered to RPo under current footprinting conditions and found to equal 0.8 of the nominal dose applied, irrespective of the oxidant concentration. Accordingly, corrected doses were used in calculation of rate constants for inactivation of RPo and oxidation of Ts.

Results Inactivation of RPo Inactivation of RPo by KMnO4 oxidation as a function of the oxidant dose was examined at 37 °C by determining fraction F of transcription-competent complexes left after application of an effective oxidant dose x using the FDAI assay, described under Materials and methods. In view of the large enhancement by Mg2þ ions of the oxidizability of thymines within the transcription bubble of RPo [17], inactivation of the latter

was studied both in the absence of Mg2þ and in the presence of 10 mM MgCl2 . As shown in Fig. 2, the data points F ðxÞ could be fitted by a single-exponential function, F ðxÞ ¼ expðkin xÞ;

ð1Þ

where x ¼ c
t (M
s) is the effective oxidant dose applied at concentration c for a time period t, and kin is the pseudo-first-order rate constant for RPo inactivation. Values of the kin and kin;Mg rate constants obtained from the fits, 4.25(0.06)
102 M1 s1 and 2.50(0.07)
102 M1 s1 , (cf. legend to Fig. 2), indicated that inactivation of RPo in the presence of Mg2þ was about twofold slower than that in its absence. Comparison of these rate constants with those for oxidation of individual thymines in the transcription bubble region of the complex studied (cf. Table 1) showed that the process of RPo inactivation in the absence of Mg2þ was two to three orders of magnitude faster than the SH oxidation of the separated DNA strands in RPo; at 10 mM Mg2þ , these differences were an order of magnitude smaller. For comparison purposes inactivation of free RNAP was also studied using the same FDAI assay. At 37 °C the reaction proved to be extremely fast, even at the lowest oxidant doses used for RPo inactivation. Therefore, the free holoenzyme was treated with KMnO4 at 0 °C and its inactivation assayed at 37 °C (cf. insets to Figs. 2A and B). The effect of protection of the enzyme by Mg2þ was in this case distinctly smaller, as measured by the ratio of kin;Mg =kin ¼ 0:73 (cf. legend to Fig. 2). EMSA investigation of the oxidative destruction of RPo In view of the observed rapid oxidative inactivation of RPo, it was of analytical interest to examine by EMSA whether, apart from the destruction of its active center, the oxidized RPo,ox complex becomes thermally unstable and dissociates into components in the course of the reaction. Analysis of RPo oxidized at 37 °C with

Fig. 2. Kinetics of inactivation by KMnO4 of RPo formed by E. coli RNAP at the Pa promoter. (A) Activity of RPo oxidized in the absence of Mg2þ at 37 °C, measured by FDAI assay at the same temperature, as a function of the oxidant dose; the inset shows a similar plot for inactivation of the free holoenzyme at 0 °C, measured by FDAI assay at 37 °C. (B) Activity versus oxidant dose plots for RPo and free RNAP (inset) oxidized under similar experimental conditions in the presence of 10 mM MgCl2 . Solid lines in (A) and (B) correspond to the fitted pseudo-first-order functions with kin and kin;Mg inactivation rate constants: 4.25(0.06)
102 and 2.50(0.07)
102 M1 s1 for RPo, and 8.9(0.1)
102 and 6.5(0.4)
102 M1 s1 for RNAP, respectively. The data points are average values from n ¼ 4 independent experiments with SD errors indicated by vertical bars; in parentheses are standard errors from nonlinear weighted least squares analysis.

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Table 1 Pseudo-first-order rate constants for oxidation of thymines (ki ) by KMnO4 in the bubble region of the RPo open transcription complex and for the competing reactions (ri ) leading to the destruction of the open complex Thymine (DNA strand)

ki a

ria

ki b

T + 3 (nt)

2.2 (0.4) 2.28 (0.07) 1.15 (0.1) 0.94 (0.01) 2.6 (0.2) 2.46 (0.06) 6.1 (0.6) 5.25 (0.11) 2.2 (0.2) 1.54 (0.03) 8.1 (0.2) 7.07 (0.29) 4.8 (0.1) 3.8 (0.1) 4.3 (0.3) 3.80 (0.08) 1.5 (0.1) 1.18 (0.06) 1.7 (0.1) 0.98 (0.03) 0.5 (0.1) 0.29 (0.03)

19.4 (5.6)

5.2 (0.4) 5.90 (0.29) 2.7 (0.2) 2.41 (0.14) 4.9 (0.3) 3.95 (0.14) 13.5 (1.6) 9.35 (0.57) 3.7 (0.3) 2.49 (0.10) 11.5 (0.3) 11.2 (0.56) 7.4 (0.3) 6.0 (0.2) 6.4 (0.5) 5.04 (0.40) 2.6 (0.2) 1.95 (0.15) 8.0 (0.9) 4.17 (0.25) 2.1 (0.4) 1.04 (0.09)

T + 2 (nt) T ) 2 (nt) T ) 3 (nt) T ) 4 (nt) T ) 11 (t) T ) 9 (t) T ) 8 (t) T ) 5 (t) T ) 1 (t) T + 1 (t)

25.8 (6.3) 2.1 (1.9) 18.3 (4.5) 22.0 (2.9) 7.7 (0.6) 9.6 (0.5) 21.4 (3.0) 22.4 (2.4) 58.3 (4.4) 68.5 (19.6)

ri b 5.7 (2.2) 7.0 (2.4) 0.5 (1.5) 24.4 (5.2) 16.5 (3.4) 3.7 (0.8) 5.3 (0.7) 11.9 (2.9) 17.0 (3.3) 76.9 (11.0) 62.5 (15.5)

The rate constants (M1 s1 ) (standard errors in parentheses) were determined from global nonlinear weighted least squares fit of Eqs. (5)–(8) to the fi ðxÞ data from the whole (SH and MH) range of oxidant doses (Fig. 6). Italics indicate ki data from the SH range of oxidant doses from ref. [17], corrected for the redox capacity of buffer A. a [Mg2þ ] ¼ 0. b [Mg2þ ] ¼ 10 mM.

Fig. 3. EMSA analysis of KMnO4 oxidized RPo at 32 P-labeled Pa promoter DNA. (A and B) Samples treated with increasing oxidant dose (indicated) at 37 °C in the absence of Mg2þ and at 10 mM MgCl2 , respectively, resolved at 35 °C for 50 min; (C and D) fractions of stable RPo,ox left after treatment with increasing oxidant dose obtained by quantification of gels shown in (A and B), respectively. Solid lines in the plots represent fitted single-exponential function describing pseudo-first-order kinetics of RPo,ox dissociation (Eq. (1)), kdis ¼ 32ð2Þ M1 s1 and kdis;Mg ¼ 17ð3Þ M1 s1 . The data points are average values from n ¼ 3 independent experiments with SD errors indicated by vertical bars; in parentheses are standard errors from nonlinear weighted least squares analysis.

KMnO4 doses from 0.005 to 0.1 (M
s) (Figs. 3A and B) clearly shows that with the growing oxidant dose, (i) the amount of RPo decreases while the amount of free DNA increases (in the presence of Mg2þ this process is less pronounced) and (ii) mobility of RPo,ox becomes

somewhat reduced, while the DNA band broadens (in the presence of Mg2þ the DNA band first broadens and then reappears as a retarded narrow band against a diffuse background). Fractions of stable RPo,ox complexes, determined by quantification of gel images

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corresponding to reactions carried out in the absence and in the presence of Mg2þ (Figs. 3A and B) and plotted against the oxidant dose applied (Figs. 3C and D), could be fitted to a single-exponential function similar to Eq. (1) (cf. solid lines in Figs. 3C and D). Derived from the fits values of the apparent kdis ¼ 32ð2Þ M1 s1 and kdis;Mg ¼ 17ð3Þ M1 s1 for RPo,ox, formed in the absence of Mg2þ and at 10 mM MgCl2 , respectively, indicate that the protective effect of bound Mg2þ on complex dissociation is similar to that observed in the process of RPo inactivation (see above). Comparison of the dose–response plots for the FDAI and EMSA assays (cf. Figs. 2 and 3) and fitted kin and kdis parameters allowed us to conclude that inactivation of RPo by far precedes RPo,ox dissociation. For instance, exposure of RPo to an oxidant dose as low as 0.005 (M
s), shown to cause its inactivation to about 0.1 and 0.2 of the initial level in the absence and presence of Mg2þ , respectively (cf. Fig. 2), led to dissociation of 20–30% of the complexes (cf. Fig. 3). All these results demonstrate that as RPo becomes oxidized, its structure changes progressively, ultimately leading to dissociation of RPo,ox into components. Free RNAP holoenzyme inactivated by a similar KMnO4 dose at 0 °C lost its ability to bind promoter DNA under experimental conditions appropriate for RPo formation (EMSA experiments, data not shown). To evaluate the stability of the oxidized complexes in the experimental time scale, RPo was oxidized with increasing KMnO4 doses in the range of 0.005–0.1 (M
s) at 37 °C, the reaction was arrested, incubation was prolonged either at the same temperature or at 47 °C, and aliquots removed at various times were analyzed by EMSA at 35 °C on the same running gel. Results of this analysis are exemplified in Figs. 4A and B for the reaction products obtained at oxidant doses of 0.015 and 0.025 (M
s), respectively. Since the ratio of intensities of bands due to RPo,ox and DNA did not change over 30 min, we conclude that partially oxidized complexes are stable in solution both at 37 and at 47 °C. To check the thermal stability of RPo,ox under EMSA condi-

tions, the reaction products obtained after application of three different KMnO4 doses at 37 °C in the presence of 10 mM MgCl2 were first resolved by EMSA for 15 min at 37 °C and then electrophoresed for an additional 105 min either at 37 or at 42 °C (cf. Fig. 4C and D). While the gel patterns at 37 °C (Fig. 4C) proved similar to those obtained at 35 °C, results corresponding to 42 °C (Fig. 4D) indicated complete dissociation of the complexes oxidized with the KMnO4 dose as low as 0.005 (M
s). RPo,ox is thus extremely thermolabile above 37 °C under EMSA conditions. In such experiments gel temperature should therefore be strictly controlled and kept at or below 37 °C. In EMSA, the loading buffer contained heparin, a nonspecific DNA competitor, to challenge nonspecific complexes between RNAP and promoter DNA. Its presence could diminish somewhat the thermal stability of RPo,ox. An attempt was also made to check whether RPo,ox rearranges in the course of the oxidation reaction. For this purpose RPo was oxidized at 37 °C in (i) one step by treatment with KMnO4 at 20 mM concentration for 10 s (incident dose of 0.2 (M
s)) or in two steps with the same total dose, attained by treatment in the first step for either (ii) 60 s at 0.6 mM or (iii) 180 s at 0.2 mM oxidant concentration, followed by addition of the oxidant to 16.4 mM and continuation of the reaction for 10 s. Analysis of the reaction products by EMSA (not shown) indicated that both the extent of dissociation of RPo,ox into its components and the extent of DNA oxidation were the same in all the three (i)–(iii) experiments. This means that the reaction products accumulated in RPo,ox additively in the course of the reaction. Moreover, these experiments demonstrated that oxidant concentration and time of exposure are equivalent kinetic variables of the reaction. KMnO4 footprinting of the ssDNA region in RPo Our aim was to evaluate reaction rate constants for oxidation of thymine residues within each of the separated DNA strands in RPo from footprints obtained

Fig. 4. Stability of RPo,ox in solution and under EMSA conditions. (A and B) RPo,ox formed at 37 °C in the presence of 10 mM MgCl2 by treatment with KMnO4 doses of 0.015 and 0.025 (M
s), respectively, incubated at the same temperature (A) and at 47 °C (B) with aliquots withdrawn at indicated times analyzed by EMSA on the running gel. EMSA analysis at 37 °C (C) and at 42 °C (D) of oxidation products of RPo formed at 37 °C in the presence of 10 mM MgCl2 by treatment with indicated KMnO4 doses.

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Fig. 5. Examples of KMnO4 footprinting patterns of the melted DNA region in RPo. (A and B) Autoradiograms of 6% polyacrylamide sequencing gels (at the right side whole, at the left side enlarged fragments corresponding to the melted DNA region) showing resolved 32 P-end-labeled ssDNA products of Klenow primer extension reaction carried out on the template (A) and nontemplate (B) Pa promoter DNA strands; doses of KMnO4 (in (M
s)) are indicated at the top of the lanes 1–12, ‘‘)’’ and ‘‘+’’ signs indicate the absence and the presence of 10 mM MgCl2 in footprinting reactions; along the leftmost lines are indicated position of DNA bands corresponding to base residues of the Pa promoter in the melted region of the template and nontemplate promoter strands. (C and D) Selected (lanes 4) integrated intensity profiles of the melted Pa region (solid lines) of the template (C) and nontemplate (D) promoter strands, deconvoluted into Gaussian components P1–P10 (dotted lines) corresponding to individual oxidized thymines (marked by arrows). (E) Footprints of the melted nontemplate promoter region of RPo obtained at indicated KMnO4 doses and reaction times, at 37 °C and in the presence of 10 mM MgCl2 .

with a broad range of oxidant doses embracing SH and MH promoter DNA oxidation. The patterns of trial footprints obtained at a selected oxidant dose, attained either by exposure of RPo to various oxidant concentrations for the same reaction time or by varying the latter at a constant KMnO4 concentration, were similar (cf. Fig. 5E). This confirmed the equivalence of the two

kinetic parameters. Therefore, footprints as a function of the oxidant dose were obtained (exemplified in Fig. 5) by varying the concentration of KMnO4 at a constant exposure time. The concentration of the oxidant was always high enough to set the pseudo-first-order conditions for the oxidation reaction. The footprints were scanned and quantified as described under Materials

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and methods. Analysis of the scans was performed in three steps. First, the integrated intensity (IQ volume integration) of the group of bands corresponding to DNA fragments terminated at all the oxidized bases within the transcription bubble was measured and normalized to the integrated intensity of the whole lane. Then, the distribution of the integrated intensity within the bubble along the lane, an intensity profile (cf. solid contour line in Figs. 5C and D), was obtained using the IQ area integration function. Integrated intensities of particular bands (marked Pi in Figs. 5C and D) were evaluated by deconvolution of the intensity profiles, assuming Gaussian distribution of the intensity within the bands and a constant half-width for all the bands. Some of the bands were visibly doubled (cf. Figs. 5A and B) while those corresponding to Tn runs were unresolved n þ 1 multiplets. This was due to the termination of the primer extension reaction at a base preceding the thymine oxidized to corresponding 5,6-diglycol, when the latter was hydrolyzed to an urea derivative during alkaline denaturation [25,26]. Therefore, sums of integrated band areas corresponding to DNA fragments terminated at these two forms of oxidized thymines were calculated, assuming a constant ratio between the diglycol and the urea forms for all the thymines in a given experiment. This ratio was estimated from integrated areas of the doubled bands seen at the edges of sequences T(+3)G(+4) and A()12)T()11) in the nontemplate and template strands, respectively. In this manner fractions fi of all oxidized thymine bases at the selected oxidant doses were determined. In Fig. 6, fractions fi ðxÞ of reactive thymines in the template and nontemplate DNA strands are plotted as a function of the oxidant dose x. Owing to very low intensity of bands attributed to T ) 6, T ) 7, and T ) 10 and their abnormal behavior at high oxidant doses discussed later in the text, they were quantified as the whole group (not shown), and the corresponding data were not included in the kinetic analysis. The fi ðxÞ data were first analyzed according to pseudo-first-order reaction kinetics. The general pattern of both sets of plots indicates that approximately up to the oxidant dose of ca. 0.03 (M
s) the fractions fi grew single-exponentially, as observed previously [17], and started to deviate from this simple relationship when the reaction entered the MH regime. The plots clearly show the strong and greatly differentiated effect of Mg2þ ions on the reactivity of all thymines within the melted DNA region. Previous analysis of fi ðxÞ data obtained at SH oxidation of RPo at the Pa promoter [17] provided pseudo-first-order rate constants for oxidation of all Ts in the transcription bubble region, both in the absence and in the presence of 10 mM MgCl2 , and quantitative assessment of the effect of Mg2þ ions (cf. Table 1). For quantitative analysis of the fi ðxÞ data corresponding to the whole range of oxidant doses applied, from SH to

MH oxidation of bubble DNA, we attempted to apply the kinetic scheme proposed by Tsodikov et al. [16] for analysis of similar data for the open complex at the kPR promoter. According to this scheme, variation of fi fractions and the Pfraction of unoxidized DNA fragments, fu ¼ 1  fi , with the oxidant dose, x ¼ ct, compatible with pseudo-first-order reaction conditions, can be described by a set of equations, ! N X X fu ¼ 1  fi ; ¼ f exp  kj x þ ð1  f Þ; ð2Þ j¼1

f1 ¼ f ½1  expðk1 xÞ ; and " ! i1 X kj x  exp fi ¼ f exp  j¼1

ð3Þ 

i X

!# kj x

;

ð4Þ

j¼1

where f is the initial fraction of oxidizable promoter DNA, f1 is the fraction of the very first Ti encountered in the primer extension reaction by Klenow polymerase on either DNA strand, i.e., T + 3 and T ) 11 in our case, and k1 and kj are pseudo-first-order rate constants of oxidation. However, attempts to fit functions (2)–(4) to the experimental fi ðxÞ data, by initially setting the values of the kj rate constants at values determined from SH footprints (Table 1), proved unsuccessful. The calculated curves greatly deviated from fi vs x plots (cf. solid lines in plots in insets in Fig. 6). When kj were allowed to vary, their best-fit values differed greatly from those obtained from fi ðxÞ data corresponding to SH reaction conditions. The reason for the apparent inapplicability of the theoretical MH model is most obvious from the plots of the experimental fi ðxÞ data for T + 3 and T ) 11 and unoxidized DNA in comparison with those calculated with respective rate constants obtained from analysis of SH footprinting data (cf. respective panels and insets in Fig. 6). According to Eqs. (2) and (3), fractions f1 and fu at sufficiently high values of x should approach f and 1  f , respectively. A comparison of fi and fu values at x ¼ 0:23 (M
s) demonstrates that in neither case was this prediction fulfilled. The experimental values of f1 fractions for T + 3 and T ) 11 at x ¼ 0:23 differ greatly within the same footprinting experiment and are smaller than 1  fu . Dissociation of the oxidized complexes into components, shown by EMSA experiments, seems to be a major cause of the observed deviation of experimental data from those predicted by the theoretical MH model, which neglects the occurrence of this process. Results of the EMSA experiments demonstrated that accumulation of oxidative lesions both in the RNAP and in the promoter bubble region affects the structure of the open complex, which manifests itself in retardation and broadening of bands due to RPo,ox and DNA (cf. Fig. 3). Therefore, even partial collapse of the structure of RPo,ox before its dissocia-

T. Łozi
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247

Fig. 6. Kinetics of oxidation of thymines in the bubble region of the template and nontemplate DNA strands of the Pa promoter in RPo. Data points fi ðxÞ (average values: n ¼ 4, calculated standard errors in the range of 0.05–0.2) corresponding to DNA fractions of unoxidized DNA (fu ) and oxidized Ts (marked by position number relative to the transcription start point, cf. Fig. 1) in the template (left column) and nontemplate (right column) strands, in the absence (j) and in the presence (d) of 10 mM MgCl2 were obtained by quantification of footprints (exemplified in Fig. 5) as described under Materials and methods; the solid lines correspond to fitted functions (Eqs. (5)–(9)). In insets, the same fi ðxÞ data points (j) as in the main panel with simulated functions (solid lines) given by Eqs. (2)–(4) with SH ki values (17) from Table 1.

tion may lower the accessibility of some Ts to the oxidant. The 14-bp bubble region of the promoter, accumulating at most one to two oxidized Ts per strand, upon release of DNA from the complex is expected to

renature to the dsDNA form in which reactivity of pyrimidines toward KMnO4 is by two to three orders of magnitude lower than that in ssDNA [27]. Hence, further oxidation of the dissociated promoter DNA could

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not be observed in the range of oxidant doses used in this study.2 Since the fraction of dissociated oxidized DNA and that obtained after deproteinization of RPo,ox were simultaneously analyzed by the primer extension method, the observed extent of oxidation of Ts is lower than that expected from the theoretical model. Determination of the amount of DNA released from oxidized complexes under conditions of the footprinting reaction did not prove technically feasible. Therefore, an attempt was made to account for the overall effect by introducing into the MH kinetic scheme of Ts oxidation in RPo the following: (i) pseudo-firstorder rate constants ri for individual Ts, each corresponding to a sum of competitive processes leading to their apparent inaccessibility to the oxidant, viz. collapse of RPo structure and dissociation, or (ii) a rate constant r common for all Ts. Consequently, Eq. (3) was reformulated to express the total fraction of DNA oxidized at site i, ox

fi ¼ f ½ki =ðki þ ri Þ f1  exp½ðki þ ri Þx g;

ð5Þ

where f is the initial fraction of complexed promoter DNA. Probability Pi of the appearance of Ti in oxidized form in the promoter bubble DNA region can be defined as Pi ¼

ox

Thus, fraction f1 ¼ f1 of end-labeled DNA fragments, primer extended to T + 3 or T ) 11, is given by Eq. (5), where ki ¼ k1 and ri ¼ r1 or by a rearranged form of Eq. (6): f1 ¼ fP1 :

ð7Þ

Fractions fi ði P 2Þ of longer end-labeled DNA fragments corresponding to Ts located in either DNA strand upstream from the first one (T + 3 or T ) 11) at the primerÕs end are equal to the product of (fPi ) and respective probabilities ð1  Pj Þ of finding DNA unoxidized at sites occupied by preceding potentially oxidizable Tj ðj < iÞ residues: i1 Y fi ¼ fPi ð1  Pj Þ; i P 2: ð8Þ j¼1

Fraction fu of DNA unoxidized at all Ti sites is i Y ð1  Pj Þ þ ð1  f Þ:

ð9Þ

j¼1

Replacement in Eqs. (8) and (9) of Pi and Pj terms by appropriate functions in the form of Eq. (5) yields an2

Discussion

ð6Þ

fi =f : ox

fu ¼ f

alytical expression allowing fitting of all the experimental fi ðxÞ data to the model (i). Introducing in Eq. (5) a rate constant r common for all Ts allowed similar testing of the simpler model (ii). A global nonlinear least squares analysis of all the experimental fi ðxÞ data according to Eqs. (5)–(8) for each of the two models has shown that the model (i), assuming individual ri rate constant for each Ti , provides a better fit in terms of v2 values3 than the model (ii). This model was therefore used to evaluate the rate constants ki and ri for all the oxidizable Ts in both DNA strands. Solid lines drawn through the experimental data points in Fig. 6 correspond to the fitting functions. The similarity of ki values, within the indicated error range, obtained from the analysis of the fi ðxÞ data from SH [17] and SH–MH ranges of oxidant doses (cf. respective data in Table 1), validate the general applicability of the proposed model of competitive oxidation of the open complex. However, the accuracy and reliability of the kinetic data deduced from the SH–MH footprints with the help of this model are less satisfactory than those obtainable from SH footprints [17].

Structural and thermal melting studies on a model dsDNA containing a single thymine diglycol residue instead of thymine [28,29] have shown that the modified thymine adopts an extrahelical position and introduces a kink to the host duplex, which causes lowering of the thermal stability of DNA. Such perturbation of DNA structure may increase to some extent permanganate reactivity of neighboring thymines, such as extrusion of short hairpins with AT basepairs [27].

RNAP oxidation The high susceptibility of RNAP to oxidation by KMnO4 , both as free holoenzyme and bound to DNA in RPo, can be rationalized by the presence of amino acid residues expected to be particularly vulnerable to this reaction, e.g., side chains of cysteine and aromatic amino acids, in close proximity to the enzyme active center and r70 subunit domains responsible for promoter DNA and core RNAP recognition and binding [15,30–34] Cys-454, flanking the Mg2þ binding motif NADFDGD of the b0 subunit, next to which the major cleavage of the polypeptide chain in Fe2þ -induced Fenton reaction has been shown to occur [35], is located close to the active center of the open complex. One can thus hypothesize that MnO 4 diffused into the substrate site of RPo may primarily oxidize Cys-454 to cysteinic acid, thus severely perturbing the structure of the active site. This reaction can be expected to be competitive to the oxidation of some Ts in the adjacent template strand region, that is T(+1) and T()1). Indeed, these thymines are characterized by the largest values of ri rate constant (cf. Table 1) and appear in the footprints as the weakest DNA bands (cf. Fig. 5). Permanganate ions, isosteric with phosphate ions, were formerly used to oxidize 3

v2 values from fits of the fi ðxÞ data sets for Ts in the nontemplate and template (10 mM MgCl2 ) DNA strands to model (i) and (in parentheses) to model (ii): 3.4 (8.5) and 0.4 (8.3), respectively.

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selectively the thiol groups in the catalytic center of aspartate carbamylase [36] and of phosphoribose–pyrophosphate synthetase [37], without any measurable effect on the content of other amino acids in these proteins. Also oxidation of some of the four Cys residues forming the Zn2þ -binding domain at the N-terminal half of the b0 subunit, b0 1ZBD [38,39], can be expected to disturb severely the structure of both RNAP and RPo. This motif was recently shown to be involved in proper assembly of the E. coli holoenzyme [40] and specific recognition of region 3.1–4.2 of r70 [41], whereas in RPo, it may bind to the promoter DNA backbone in the spacer region between its extended )10 and )35 elements [15]. In addition, oxidative modification of aromatic amino acid side chains (Tyr, Trp, Phe) of the 2.3 domain of r70 , involved in specific recognition of the )10 promoter element through stacking interactions with base residues of the nontemplate strand [30,42], may lower RPo stability and increase accessibility of Ts in this region toward MnO 4 . A closer inspection of footprints in Fig. 5 indicates that residues T ) 10, T ) 7 and T ) 6 of the nontemplate strand, in contact with the aromatic amino acids of the 2.3 domain, appear more reactive in the MH range of KMnO4 doses (cf. lanes 5, 7, 9, and 11), particularly in the absence of MgCl2 . The increased reactivity of these Ts may be attributed to the loss of specific contacts between the )10 promoter region and the 2.3 domain bearing oxidized aromatic side chains. A concerted action of MnO 4 at the catalytic (Cys454) and promoter binding sites of RNAP, b0 ZBD, and 2.3 of r70 can be thus expected to strongly affect the structural organization and stability of RPo, eventually leading to its dissociation. On the other hand, the two OH groups and nonplanar 5,6-dihydro–5,6-dihydroxythymine moiety may also affect local pyrimidine–protein surface interactions and thus the reactivity of adjacent thymines before RPo,ox dissociation. In the absence of Mg2þ , for most Ts ri > ki , which indicates the occurrence of strongly competitive oxidation reactions of amino acid side chains in RNAP underlying the destruction of RPo. Only in the case of nontemplate T()2) and template T()11) are the ri and ki values comparable. This seems to be connected with their location near the RPo surface [15]. Addition of Mg2þ caused a several fold increase in ki values, as observed previously [17], with concomitant, but not proportional, decrease in ri rates. In effect, for most Ts the relationship between the two rate constants becomes reversed, i.e., ri < ki , whereas their sums become (ri þ ki ÞMg smaller than in the absence of Mg2þ . The latter observations are in full agreement with the documented general protective effect exerted by bound Mg2þ on RPo inactivation and RPo,ox dissociation. The greatly differentiated reactivity of Ts in the bubble promoter region is most probably determined by local

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sterical and electrostatic barriers for diffusion of MnO 4 to the reaction centers. Consequently, general enhancement of their reactivity by Mg2þ cations was interpreted [17] as due mainly to lowering of the electrostatic barrier by ions diffusely bound to ssDNA phosphates, whereas particularly strong enhancement of the reactivity of template strand T + 1 and T ) 1 was interpreted as due to further lowering of this barrier by the two cations chelated to the carboxylic groups of the active center b0 NADFDGD Mg2þ -binding motif [34,35]. The opposite, protective effect of Mg2þ observed in inactivation of RPo by permanganate should be therefore attributed to the interaction of these ions with the RNAP component of the complex. In the crystal structure of T. thermophilus RNAP holoenzyme [34], many Mg2þ ions form a coat on the protein surface, presumably facilitating wrapping of the DNA molecule around the enzyme upon formation of the transcription complex. It can be expected that some of these cations stabilize also the structure of RPo, preventing deeper penetration of MnO 4 into RNAP and thereby protecting most vulnerable amino acids from oxidation. Recommendations for footprinting of protein–DNA complexes In light of the presented results on oxidation of RPo by KMnO4 , it is evident that the generally neglected consequences of oxidative damage to RNAP might often lead to misinterpretation of footprints usually obtained at a single very high oxidant dose, similar to the recommended 1.2 (M
s) (10 mM KMnO4 and 2 min exposure time) [7]. This dose is several fold larger than the largest one used in this study and shown to cause MH oxidation of the promoter bubble ssDNA and severe oxidative damage to RNAP, leading to complete inactivation and dissociation of RPo,ox. We have demonstrated that interpretation of the footprints determined as a function of the oxidant dose up to 0.23 (M
s), according to the competitive model of RPo oxidation, allows one to obtain only approximate values of rate constants for thyminesÕ oxidation in the promoter bubble region. Moreover, this approach is tedious, time consuming, and costly. Thus footprinting experiments should be performed as a function of the oxidant SH dose (by varying either oxidant concentration or time of the reaction) and interpretation of the data should be based on the rate constants for pyrimidinesÕ oxidation derived from the pseudo first-order kinetics of the reaction. For kinetic analysis the doses applied should be corrected for using a fraction of the oxidant in reactions with organic components of binding buffers. The strongly pH dependent redox potential of the MnO 4/ Mn2þ pair at pH > 6 becomes lower than that of Cl / Clo , and oxidation of chlorides present abundantly in

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neutral or slightly alkaline binding buffers is negligible. We showed that oxidation of organic components of buffer A (Tris and EDTA) lowered the nominal oxidant dose applied by a factor of 0.8. The reduction capacity of Hepes buffer, occasionally applied in footprinting experiments [8,13], was found to be about 40-fold higher than that of Tris buffer A (data not shown), hence the use of Hepes should be avoided. Similarly, the addition to binding buffers of such compounds as DTT, mercaptoethanol, and BSA, used often to prevent oxidation of RNAP by oxygen, should also be avoided. Recommendations emerging from the results of this work are directly applicable to footprinting of transcription complexes of prokaryotic RNA polymerases. Moreover, they also provide a general methodology for consideration and designing of experimental conditions for KMnO4 footprinting of transcription complexes of eukaryotic RNA polymerases and other DNA–protein complexes.

Acknowledgments The authors express their gratitude to Dr. Krystyna Bolewska and Teresa Rak for preparation of RNA polymerase.

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