Tài liệu Báo cáo khoa học: Kinetics and thermodynamics of nick sealing by T4 DNA ligase pptx

Kinetics and thermodynamics of nick sealing by T4 DNA ligase Alexey V. Cherepanov* and Simon de Vries Kluyver Department of Biotechnology, Delft University of Technology, the Netherlands T4 DNA ligase is an Mg 2+ -dependent and ATP-dependent enzyme that seals DNA nicks in three steps: it covalently binds AMP, transadenylates the nick phosphate, and cata- lyses formation of the phosphodiester bond releasing AMP. In this kinetic study, we further detail the reaction mechan- ism, showing that the overall ligation reaction is a super- imposition of two parallel processes: a ễprocessiveế ligation, in which the enzyme transadenylates and seals the nick without dissociating from dsDNA, and a ễnonprocessiveế ligation, in which the enzyme takes part in the abortive adenylation cycle (covalent binding of AMP, transadenylation of the nick, and dissociation). At low concentrations of ATP (< 10 l M ) and when the DNA nick is sealed with mismatching base pairs (e.g. ve adjacent), this super- imposition resolves into two kinetic phases, a burst ligation (% 0.2 min )1 ) and a subsequent slow ligation (% 2 ã 10 )3 min )1 ). The relative rate and extent of each phase depend on the concentrations of ATP and Mg 2+ .The activation energies of self-adenylation (16.2 kcalặmol )1 ), transadenylation of the nick (0.9 kcalặmol )1 ), and nick- sealing (16.318.8 kcalặmol )1 ) were determined for several DNA substrates. The low activation energy of transadeny- lation implies that the transfer of AMP to the terminal DNA phosphate is a spontaneous reaction, and that the T4 DNA ligaseAMP complex is a high-energy intermediate. To summarize current ndings in the DNA ligation eld, we delineate a kinetic mechanism of T4 DNA ligase catalysis. Keywords: DNA ligase; end-joining; kinetics; mechanism of action; mismatching nick. T4 DNA ligase is an enzyme that catalyses formation of the phosphodiester bond between the adjacent 5Â-PO 4 and 3Â-OH groups of two dsDNA fragments [1]. It is able to join two dsDNAs (blunt-end or sticky-end ligation), or seal a break between two ssDNA fragments annealed on the complementary DNA strand (nick-ligation). It can join phosphodiester linkages on triple-stranded nucleic acids [2], seal single-stranded 15-nucleotide gaps [3], and act as a lyase, removing apurininc/apyrimidinic (AP) sites in DNA [4]. The enzyme requires a bivalent metal cation such as Mg 2+ or Mn 2+ , and joins DNA using ATP as a coenzyme. One phosphodiester bond in DNA is formed per ATP molecule hydrolysed to AMP and pyrophosphate. The mechanism of catalysis of T4 DNA ligase comprises three steps and involves two covalent reaction intermedi- ates: E ỵ ATP $ EAMP ỵ PP i 1ị E AMP ỵ ndsDNA $ EAMPndsDNA ! E ỵ AMPndsDNA 2ị E ỵ AMP ndsDNA $ E AMPndsDNA ! E ỵ dsDNA ỵ AMP 3ị where ndsDNA is nicked dsDNA, AMPndsDNA is ndsDNA adenylated at the 5Â-phosphate of the nick, and a one-sided arrow indicates that the ễreverseế reaction is at least three orders of magnitude slower than the ễforwardế reaction. On the basis of the electrophoretic mobility shift assay experiments, it has been suggested that adenylated ligase forms transient Tcomplexes, EAMPndsDNA, in search of a phosphorylated 5Â-end of (n)dsDNA. When the nick phosphate is found, it is adenylated, and a stable ễScomplexế is formed, EAMPndsDNA [5]. The enzyme in this complex has been suggested to ễstallế on DNA until the nick is sealed and dsDNA is released. Formation of the rst phosphodiester bond during joining of the blunt ends has been proposed to happen accordingly; the main difference is the 2 : 1 dsDNA to enzyme stoichiometry in step 3. It has been suggested [5] that during blunt end joining, the ternary complex ligaseDNA is formed via two second-order associative reactions: E ỵ AMPdsDNA ! EAMPsDNA (Scomplex) Scomplex ỵ dsDNA ! EAMPdDNAdsDNA Despite a good understanding of the overall reaction mechanism, relatively few articles have been dedicated to the kinetic studies of catalysis performed by this enzyme. The optimal concentration of Mg 2+ in the nick- joining reaction (810 m M ), apparent K m for the nicked Correspondence to S. de Vries, Kluyver Department of Biotechno- logy, Delft University of Technology, Julianalaan 67, 2628 BC Delft, the Netherlands. Fax: + 31 15 2782355, Tel.: + 31 15 2785139, E-mail: s.devries@tnw.tudelft.nl Abbreviations: ndsDNA, nicked dsDNA. Enzymes: DNA ligase (EC 6.5.1.1). *Present address: Metalloprotein & Protein Engineering Group, Leiden Institute of Chemistry, Gorlaeus Laboratories, Leiden University, Einsteinweg 55, PO Box 9502, 2300 RA Leiden, the Netherlands. (Received 28 May 2003, revised 8 September 2003, accepted 9 September 2003) Eur. J. Biochem. 270, 43154325 (2003) ể FEBS 2003 doi:10.1046/j.1432-1033.2003.03824.x dsDNA (1.5 n M ), and ATP (14–20 l M ) as well as the apparent inhibition constant K i for dATP (10–35 l M ) had been determined during the initial characterization of the enzyme [6,7]. It was shown that T4 DNA ligase binds ATP covalently forming a lysine (e-amino)-linked adenosine monophosphoramidate [8]. Harvey et al. [9] have demonstrated that sealing of the pre-adenylated nick in dsDNA is inhibited when T4 DNA ligase is preincubated with ATP and Mg 2+ . Later it was shown that the enzyme obeys Ping-Pong kinetics, and joins dsDNA with multiple nicks in the nonprocessive mode. The true K m for ATP (100 l M ) was determined in the joining reaction with polydAÁd(pT) 10 as substrate (true K m ¼ 0.6 l M ) [10]. It was shown that T4 DNA ligase seals nicks containing base pair mismatches [11–15], and the effect of the ionic strength on the apparent K m of the mismatching dsDNA nick has been evaluated (e.g. 200 n M at 0.2 M NaCl vs. 50 n M without salt) [12,16,17]. The pH-dependent equilibrium constant for step 1 (0.0213 at pH ¼ 7.0 and 25 °C, pK a ¼ 8.4) and the standard free energy for cleavage of ATP to AMP ()10.9 kcalÆmol )1 ) have been determined [18]. It was shown that T4 DNA ligase is capable of synthesizing the dinucleoside polyphosphates, such as Ap 3 A, Ap 4 A, Ap 4 G, and Ap 4 dA, using ADP (d)ATP, GTP, and P 3 as substrates [19,20]. This secondary enzyme activity may stem from the fact that ligase has two closely located nucleotide-binding sites [21]. Recently, a dynamic mechanism of nick recognition by DNA ligase has been proposed [22]. The key feature of the mechanism is the B-to-A DNA helix transition of the enzyme-bound dsDNA motif, which results in DNA contraction, bending, and unwinding. For non-nicked dsDNA, this transition is reversible, leading to dissoci- ation of the enzyme. For ndsDNA, this transition was proposed to (a) trigger an opened–closed conforma- tional change in the enzyme, and (b) force the motif to accommodate the strained A/B-form hybrid conforma- tion, the transition state in the nick-sealing reaction. In our previous work, we assessed the ability of T4 DNA ligase to seal ndsDNA containing one to five adjacent mismatching base pairs [14,23], aiming to use this enzyme in the novel protocol of saturated scanning mutagenesis. In all cases, kinetic traces displayed pronounced biphasic beha- vior, which was most spectacular with the nick containing five base pair mismatches. Apparently, this effect has not been previously reported. To understand its origin, we decided to study the mechanism of T4 DNA ligase in more detail. We have previously reported a pre-steady-state kinetic analysis of the first step of DNA ligase catalysis (covalent binding of AMP [24]), showing that the enzyme employs a two-metal-ion mechanism for this nucleotidyl transfer reaction, using the dimagnesium ATP form (ATPÁMg 2 ) as a true substrate. The monocoordinated form, ATPÁMg, and/or free ATP bind DNA ligase noncovalently, with K d < 150 n M [21]. Nucleotidyl transfer is reversible, and the monomagnesium pyrophosphate form MgÁP 2 O 7 participates in the T4 DNA ligase-promoted synthesis of ATP [24]. This work concentrates on the steady-state kinetic analysis of the overall ligation reaction and an initial thermodynamic characterization of the catalysis. We aimed to achieve the following goals: (a) to deepen the general understanding of the kinetic mechanism of the end-joining reaction, the biphasic behavior in particular, using an ndsDNA substrate with 5¢-mismatching base pairs; (b) to extract kinetic and thermodynamic parameters of the elementary steps of ligase catalysis; and (c) to summarize current findings on the mechanism of action of DNA ligase in a single reaction scheme. Experimental procedures Enzymes and oligonucleotides Three commercial batches of T4 DNA ligase were used, and were purchased from Amersham Biosciences (Uppsala, Sweden), Roche Molecular Biochemicals (Basel, Switzer- land), and MBI Fermentas (Vilnius, Lithuania). All enzyme batches showed similar activity and substrate specificity. The proteins were essentially pure as judged by SDS/PAGE. Protein concentration in the purchased enzyme stocks was determined using the BCA protein determination kit (Pierce Biotechnology, Rockford, IL, USA). Synthetic oligonucleo- tides were purchased with Eurogentec (Seraing, Belgium), except for the one labeled with the Cy5 fluorescent marker, which was obtained from Amersham Biosciences. Model system For studies on T4 DNA ligase-promoted repair of nicks in dsDNA, we used 72/24/(6–24)-mer synthetic DNA sub- strates. Nonphosphorylated 72-mer B had the sequence 5¢-GTCCAAACAGCTATCTGCATCCGTCGACCTGC TCGGTTCCTTGGCTACACTGGCCGTCGTTTTACA ACGTCG-3¢.The24-mer5¢-DNA oligonucleotide C (5¢- here refers to the fact that the 5¢-oligomer is located upstream of the nick) had the sequence 5¢-CGACGTT GTAAAACGACGGCCAGT-3¢, and contained on the 5¢-end the fluorescent label Cy5 (Dye 667, No. 27-1801-02; Amersham Biosciences), allowing easy quantification of the products of the joining reaction. 3¢-DNA oligonucleotides (located downstream of the joining site) of different lengths (6–24-mers; Figs 1, 2, 3, 4 and 8) were 5¢-phosphorylated; oligonucleotides M5C19, M1C6, and M4C7, in addition, contained base pair mismatches next to the joining site. Ligation of ndsDNA Theligationreactionwasperformedin30lL66m M Tris/ HCl/1 m M dithiothreitol/0.05 mgÆmL )1 BSA, pH ¼ 7.6. The buffer pH (measured at +20 °C) was adjusted to 7.19 (7.34) when the ligation was performed at +4 °C(+10°C), counting dpH/°C ¼ )0.026 for the Tris/HCl buffer pair. To follow the formation of the adenylated DNA intermediate, [ 32 P]ATP[aP] was used. Concentrations of ATP, MgCl 2 , oligonucleotides, and the incubation temperature were varied according to the comments in the text. ndsDNA was prepared by mixing the necessary amount of 72-mer oligonucleotide B, a stoichiometric amount of 5¢-Cy5- labeled 24-mer C, and the required donor oligonucleotide, followed by 5 min incubation at 65 °C, 5 min incubation at 37 °C, and 10 min incubation at room temperature. T4 DNA ligation buffer was added after pre-annealing of the 4316 A. V. Cherepanov and S. de Vries (Eur. J. Biochem. 270) Ó FEBS 2003 oligonucleotide and before addition of the enzyme; the reaction mixture was preincubated at the assay temperature for 5 min. The reaction was initiated by addition of the enzyme. Over time, aliquots of 0.5 lL were withdrawn and mixedwith10lL 100% formamide/10 m M NaOH/10 m M EDTA/5 mgÆmL )1 Blue Dextran, pH ¼ 9.5. Stability of DNA ligase at ambient temperature It is known that preparations of T4 DNA (RNA) ligase gradually lose their activity when incubated at temperatures above 0 °C [6,25]. In this work, ligase was assayed at +4 °C for periods up to 25–40 h. To avoid irreversible inactivation of T4 DNA ligase during prolonged incubations, BSA was added to the reaction mixture to a concentration of 0.05 mgÆmL )1 , as reported previously [21]. Separation of ligation products and data analysis All DNA separations were performed on an ALF Express DNA sequencer (Amersham Biosciences) using 6–15% acrylamide gels (Tris/borate/EDTA/7 M urea). Usually, runs were performed at 55 °C, with 80 mA current and lasted for 1–3 h. For each data point, the fluorescence of two chromatographically separated peaks of starting mater- ial and ligation product was obtained. Separation of [ 32 P]DNA was visualized using a Molecular Dynamics Phosphorimager SI (Amersham Biosciences) and quantified using IMAGEQUANT software. Data obtained from both Cy5-labeled and 32 P-labeled DNA was imported into IGOR PRO version 4.0 (WaveMetrics Inc., Salt Lake City, UT, USA); further data analysis such as integration of peaks and fitting was performed using the built-in functions of this software package. Rate values for the burst ligation were determined as V init ¼ F ¢[t] t ¼ 0 , where F [t] is the exponential fitting function. Steady-state rates were determined by interpolating the steady-state region of the product forma- tion curve with a linear regression function. Numbers of turnovers were calculated by dividing the initial concentra- tion of ndsDNA in the reaction mixture by the concentra- tion of T4 DNA ligase. Pre-steady-state kinetic analysis Transient-state kinetic experiments were performed on the Bio Sequential Stopped-Flow Reaction Analyzer SX-18MV (Applied Photophysics, Leatherhead, Surrey, UK) using the ozone-free 150 W xenon-arc light source. The SX -18 MV software package for a single-wavelength operation mode was used for the optical measurements. Tryptophan emission was excited at 280 nm and measured as the light passing through a < 320 nm cut-off filter. Kinetic traces of protein fluorescence emission were obtained by averaging three to ten shots. Error estimates for the data values in graphs and tables represent 95% confidence intervals calculated using Student’s distribution function. In the stopped-flow instrument, we studied the transient- state kinetics of the binding of Mg 2+ to the EÁATP complex at different pH values. For this experiment, a ligase solution (7.5 l M , 0.41 mgÆmL )1 ) was prepared by dilution of a10mgÆmL )1 enzyme stock into 0.075 mgÆmL )1 BSA solution in deionized water containing 1.5 m M dithioerythr- itol and % 230 l M ATP(ATPwasaddedfroma50-m M stock solution pre-equilibrated to pH ¼ 7). This weakly buffered solution at pH % 7.5 was stored on ice until further use. A 150 m M Tris/HCl buffer of the desired pH was prepared separately, as well as the 15 m M solution of MgCl 2 in deionized water. At 5 min before the mixing shots, 150 m M Tris/HCl buffer was diluted threefold into both enzyme and Mg 2+ stocks. Then 200-lL aliquots of the resulting solutions were withdrawn, mixed with each other, and the pH of the mixture measured. In parallel, the solutions were pre-equilibrated to ambient temperature in the drive syringes and rapidly mixed in the stopped flow instrument, triggering the reaction. Results and discussion Time course of the joining reaction To study the T4 DNA ligase-promoted end-joining reaction, we used synthetic DNA oligonucleotides as described in Experimental procedures. Two ndsDNA substrates were assembled: a substrate containing a 5-bp mismatching fragment at the nick (BÁMÁ5C19), and a complementary nick substrate, BÁC24. T4 DNA ligase effectively utilizes both of these substrates (Figs 1, 3 and 4). Fig. 1. Biphasic kinetics of joining of 3¢-oligonucleotides (M5C19, C24, M1C6, and M4C7) to 72/24-mer BÁC. [dsDNA] was 1 l M ;[ATP]was 5.6 l M for M5C19, C24, and 1 m M for M1C6 and M4C7. Ligation of M5C19 and C24 was performed at +10 °C as described in Experi- mental procedures. M1C6 and M4C7 were joined at +4 °Caspre- viously described [14]. For joining of C24, the concentration of T4 DNA ligase in the assay mixture was 8 n M , 0.1 l M for M5C19, and 0.4 l M for M1C6 and M4C7. [dsDNA] (1 l M ) corresponds to the 125 turnovers of T4 DNA ligase in the case of C24 joining, 10 turnovers in the case of M5C19, and 2.5 turnovers for M1C6 and M4C7. Ó FEBS 2003 Kinetics of nick sealing by T4 DNA ligase (Eur. J. Biochem. 270) 4317 The mismatching nick (BÁMÁ5C19) is sealed in two kinetic phases (Fig. 1A). In the case of C24, these phases are less pronounced, and are observed only at low concentrations of ATP (Fig. 1B). Similar biphasic behavior is observed with nicks containing one to four mismatching base pairs (e.g. Fig. 1C,D). The origin of the two phases becomes clear when the formation trace of the adenylated ndsDNA intermediate is plotted together with the trace of the ligation product (Fig. 2). Virtually all ndsDNA is conver- ted into AMP-dsDNA before 20% of the ligation product is formed, and during this period the initial burst phase of ligation takes place. The slow ligation phase starts when all available DNA substrate and T4 DNA ligase are converted into their respective AMP-bound intermediates. The following mechanistic interpretation of the biphasic kinetics is suggested. During the burst phase, the enzyme performs ligation ÔprocessivelyÕ, i.e. by transadenylating and sealing the nick without dissociating from the DNA complex or being re-adenylated. This process is not ÔidealÕ: a fraction of ligase molecules dissociates after the transfer of AMP to the nick phosphate, rebinds AMP, and performs another transadenylation step, which in parallel to the ÔprocessiveÕ ligation leads to the accumulation of AMP–ndsDNA. The slow steady-state ligation phase starts when the concentration of the AMP–ndsDNA intermediate reaches its maximum. The overall end-joining rate decreases because the adenylated DNA is ligated either by the adenylated enzyme with a notably lower rate, or, and what is more likely, by a small fraction of the AMP-free enzyme (in agreement with previous data [9,14]). It is also clear that the formation of the phosphodiester bond rate-limits sealing of the mismatch- ing nicks, and not the adenylation of the 5¢-nick phosphate. For example, the rate of burst ligation of the oligonucleotide M1C6 (19 h )1 ) is, within experimental error, identical with the sealing rate of pre-adenylated AMP–M1C6 (18 h )1 ) [14]; adenylation of the oligonucle- otide M5C19 (1 min )1 ) is almost fivefold faster than the burst ligation (0.2 min )1 ). A complex of T4 DNA ligase with AMP–ndsDNA is more stable in the case of a complementary nick ([5]), and the enzyme hardly dissociates from ndsDNA between the steps of transadenylation and nick-sealing. As a result, the steady-state concentration of AMP–ndsDNA during liga- tion would be lower, and a difference in joining rates between the burst ligation and slow ligation phases at the same [ATP] is less pronounced. For example, in the case of BÁCÁ24, the rate of the burst phase is only about sixfold higher than the rate of the slow phase, in contrast with > 100-fold difference in the case of the mismatching nick (Fig. 1). The amplitude of the burst phase reaches % 50% of the total extent of ligation when [ATP] is taken of the same Fig. 2. Formation of the ligation product (d) and the AMP–DNA intermediate (s) during the joining of M5C19–72/24-mer BÆC. Ligation was performed at +10 °C. T4 DNA ligase (0.4 l M )sealedndsDNA (1 l M ) in the presence of 1 m M ATP and 5 m M MgCl 2 .Dottedtraces were obtained by fitting exponential functions to the experimental data points. [TP] 0 represents the number of turnovers required for the joining of all ndsDNA in the reaction mixture. Fig. 3. Joining of C24 to 72/24-mer BÆA at different concentrations of ATP. (A) Product formation curves were computed by fitting single/ double exponential functions to the experimental data points. [ATP] 0 for each curve is shown in the inset. The linear/steady-state phase of the joining reaction is magnified in the inset. Rate values were determined by fitting linear regression to the kinetic traces (fitted traces are shown in the inset). (B) (d) Turnover for the ligation of C24 and (s) M5C19 (burst ligation) at different [ATP]. The 0–50 l M region of ATP concentrations is magnified in the inset. (C) Line- weaver–Burk plot of the (d) data shown in (B). Ligation was per- formed under the conditions described in Fig. 1. The concentration of Mg 2+ was 5.1 m M . 4318 A. V. Cherepanov and S. de Vries (Eur. J. Biochem. 270) Ó FEBS 2003 order of magnitude as [ndsDNA] in the reaction mixture, e.g. [ATP] % 5 l M (Figs 2 and 4). In general, biphasic kinetics are observed because the ligase in complex with AMP–ndsDNA does not re-adeny- late itself. Otherwise, the enzyme would already be saturated with AMP at the beginning of the reaction, irrespective of the presence of ndsDNA, and would promote only slow ligation. In this sense, AMP–ndsDNA seems to ÔshieldÕ ligase from ATP. Biphasic kinetics support the previous proposals that: (a) the adenylyl moiety in the ligaseÁAMP– ndsDNA complex occupies the ATP-binding pocket of the protein [5], preventing a second nucleotide molecule from entering the active site; and/or (b) ligase-bound ndsDNA covers the ATP-binding site, hindering the access of ATP from solution [26,27]. Effect of ATP concentration The rate of the end-joining reaction catalysed by T4 DNA ligase depends on the concentration of ATP in the reaction mixture. For example, the sealing rate of the complementary nick in BÁCÁ24 increases more than 10-fold with increasing [ATP] from 5 to 400 l M , yielding K app m (ATP) ¼ 1.1 · 10 )4 M (Fig. 3), similar to the previously reported value obtained with poly(dA)Áoligo(dT) 10 [10]. There are two possible explanations for the decrease in the end-joining rate at low [ATP]: either the binding of the nucleotide to the ligase becomes rate limiting, or the equilibrium shifts towards the nonadenylated enzyme. From the pre-steady- state kinetic analysis of ATP binding to T4 DNA ligase in the absence of ndsDNA [24], both of these possibilities seem unlikely. At 5 m M Mg 2+ , the rate constant for binding of ATP is k on % 9 · 10 5 M )1 Æs )1 , giving a binding rate of 4.5 s )1 at 5 l M ATP, which is more than 100-fold faster than the observed rate of ligation at this [ATP] (% 1min )1 ). Therefore, binding of ATP could by no means limit the enzyme turnover, unless inhibition by DNA is considered. Furthermore, the apparent K d for ATP in the noncovalent EÁATP complex is below 150 n M [21], implying that at 5 l M ATP the concentration of free enzyme in the absence of ndsDNA is negligible; the ligase is essentially ATP bound and/or AMP bound. Pre-steady-state kinetic experiments in which binding of (n)dsDNA to T4 DNA ligase was studied suggest that this is a rapid process with k obs % 10 8 M )1 Æs )1 (A. Cherepanov, D. Pyshny & V. Chikaev, unpublished results). Thus, the binding of DNA at low [ATP] would be notably faster than binding of ATP (10 2 s )1 for dsDNA vs. 4.5 s )1 for ATP at 1 l M ndsDNA and 5 l M ATP). The situation is reversed at high ATP concentrations, when binding of ATP is faster than binding of DNA (10 2 s )1 for DNA vs. 4.5 · 10 3 s )1 for ATP at 1 l M ndsDNA and 5 m M ATP). Modeling studies indicate that ndsDNA in complex with ligase hinders the access of solution ATP to the nucleotide- binding site [26,27]. Therefore, it is likely that the decrease in the nick-sealing rate at low [ATP] occurs because DNA prevents binding of ATP to the ligase. This agrees with the fact that the joining rate of the mismatching oligonucleotide M5C19 does not decrease with [ATP] in the range 5 l M to 3m M (Fig. 4). In contrast, an approximately twofold increase in the joining rate is observed, compared with more than 10-fold decrease in the latter in the case of C24. In the case of M5C19, formation of the phosphodiester bond (< 0.2 min )1 ) limits the rate of the enzyme turnover, being slower than binding of a DNA substrate (10 2 s )1 ), ATP (> 4.5 s )1 ), and joining of the complementary nick under the same conditions (> 1 min )1 , Table 1). The increase in the joining rate of M5C19 at low [ATP] may indicate a shift of equilibrium towards the catalytically competent nonadenylated form of the enzyme. At [ATP] above 3 m M , inhibition of joining is observed: at 10 m M ATP the nick-sealing rate is roughly 20-fold lower than at 1 m M ATP (Fig. 3). T4 DNA ligase is known to synthesize dinucleoside polyphosphates, such as Ap 4 A[19]. In this reaction the enzyme binds a second ATP molecule in the DNA-binding site with K d 0.1–0.25 m M [21]; the Ap 4 A synthesis is inhibited by ndsDNA [20]. In our case, at low [ndsDNA] and high [ATP], the opposite situation may arise, when ATP inhibits binding of ndsDNA and subsequent ligation by occupying the dsDNA-binding site, leading to the synthesis of Ap 4 A (similar to that reported for GTP [20]). Fig. 4. Joining of M5C19 to 72/24-mer BÆC at different concentrations of ATP. (A) Product formation curves at T ¼ 10 °C. Concentration of ATP is shown in the graph. (h)130l M ATP; (j)362l M ;(s) 1.16 m M ;(d)3.15m M . The burst-ligation phase of joining is magni- fied in the inset. Reaction turnovers of the burst ligation (C) and of the slow ligation (D) are weakly [ATP]-dependent, increasing at [ATP] < 100 l M % 2-fold. The amplitude of the burst-ligation phase (B) increases more than 10-fold from % 0.3 turnovers at 3.15 m M ATP to 4.5 turnovers at 5.5 l M ATP, and the increase starts at [ATP] < 100 l M . Ligation was performed under conditions equival- ent to those described in Fig. 1. The concentration of Mg 2+ was 5 m M . [dsDNA] (1 l M ) corresponds to 10 enzyme turnovers (T4 DNA ligase was 0.1 l M ). Ó FEBS 2003 Kinetics of nick sealing by T4 DNA ligase (Eur. J. Biochem. 270) 4319 In the case of joining of the mismatching oligonucleotide M5C19, the amplitude of the burst phase increases more than 10-fold from % 0.3 turnovers at 3 m M ATP to 4.5 turnovers at 5 l M ATP (Fig. 4), while the joining rate remains roughly the same. The amplitude of the slow phase (extent of ligation) is independent of the concentration of ATP, approaching % 90% of the total concentration of the ndsDNA substrate in the reaction mixture. Effect of Mg 2+ concentration Mg 2+ is essential for DNA joining catalysed by T4 DNA ligase. It has previously been shown that the optimal concentration of Mg 2+ in the ligation mixture is % 10 m M [6]. As shown in Fig. 5A, two kinetic phases of the end- joining of M5C19 have different requirements for [Mg 2+ ]. The rate of burst ligation increases with [Mg 2+ ], and the optimum is above 5 m M Mg 2+ . In contrast, the rate of the slow phase of joining reaches its maximum at % 1m M Mg 2+ , above which joining is inhibited. It is interesting that the dependence of joining rates on [Mg 2+ ] follows the changes in the equilibrium concentrations of the mono- magnesium and dimagnesium forms of ATP (Fig. 5B). The maximal rate of the slow phase of the joining of the mismatching oligonucleotide M5C19 corresponds to the maximal concentration of ATPÁMg. On the other hand, the rate of the burst phase of joining of M5C19, and the joining rate of the complementary oligonucleotide C24 resembles more the increase in the concentration of ATPÁMg 2 . Temperature-dependence and pH-dependence of self-adenylation of T4 DNA ligase We have previously shown that self-adenylation of T4 DNA ligase in the absence of ndsDNA proceeds according to Scheme 1 [24]. Fig. 5. [Mg 2+ ]-dependence of the joining of M5C19 (A) or C24 (inset in A) to 72/24-mer BÆC and the equilibrium concentrations of different ATP forms (B). (A) Turnovers of the burst ligation for M5C19 (s), of the slow ligation for M5C19 (d); inset in (A) ligation turnover of C24. (B) Equilibrium concentrations of ATP forms were calculated using the K d values in Table 2. Table 2. K d values used to calculate equilibrium concentrations of ATP forms. Concentration of the nicked dsDNA, 1 l M (0.12 m M DNA phosphorus); ATP, 1 m M ; T4 DNA ligase, 0.1 l M in the case of M5C19 (8 n M in case of C24). Reaction K d ( M ) Reference H + + ATP 4– « HATP 3– 2.7 · 10 )7 [38] Mg 2+ + HATP 3– « MgHATP 1– 6.6 · 10 )3 Mg 2+ + ATP 4– « MgATP 2– 8.9 · 10 )6 Mg 2+ + MgATP 2– « Mg 2 ATP° 1.7 · 10 )2 0.6Mg + dsDNA-PO 4 « Mg 0.6 dsDNA-PO 4 5 · 10 )6 [39,40] Table 1. Kinetic and thermodynamic parameters of T4 DNA ligase catalysis (at 20 °C, 5 m M Mg 2+ and pH = 7.6, [ATP] = 1m M and [nds- DNA] = 1 l M ). Thermodynamic parameters were calculated by fitting Eqn (6) to the experimental data points, taking the transmission coefficient j ¼ 1. Reaction Substrate Rate, t )1 E A (kcalÆmol )1 ) DS (calÆdeg )1 Æmol )1 ) Self-adenylation Mg 2 ATP 12.5 ± 0.2 s )1 16.7 ± 0.3 1 ± 0.6 Transadenylation M5C19 1 ± 0.1 min )1 0.9 ± 0.1 ) 65.9 ± 0.2 End-joining C6 a 83.7 ± 1.9 min )1 16.3 ± 1.3 ) 4.3 ± 2.5 C24 34.3 ± 0.3 min )1 16.4 ± 0.4 ) 6.1 ± 1.3 M5C19 b 4.7 ± 0.2 · 10 )3 min )1 18.8 ± 0.7 ) 15.7 ± 2.8 a Mean values. b Value relates to the slow ÔnonprocessiveÕ ligation. Scheme 1. Kinetic scheme of self-adenylation of T4 DNA ligase. 4320 A. V. Cherepanov and S. de Vries (Eur. J. Biochem. 270) Ó FEBS 2003 T4 DNA ligase binds ATPÁMg noncovalently, but not ATPÁMg 2 . Subsequently, EÁATPÁMg binds a second Mg 2+ ion, forming a catalytic intermediate, EÁATPÁMg 2 . ATPÁMg 2 is the true substrate in the adenylation reaction, while the monomagnesium pyrophosphate form, MgÁP 2 O 7 isthetruesubstrateforthereversereaction,thesynthesisof ATP [24]. In this work, as part of an initial thermodynamic characterization of T4 DNA ligase, the activation energy of the adenylation reaction (cleavage of ATP) was deter- mined in the absence of dsDNA using the stopped-flow instrument. To minimize the contribution of the reverse reaction to the observed reaction rate, we excluded pyro- phosphate from the reaction, mixing the EÁATP complex with Mg 2+ . In the absence of excess pyrophosphate, k app À2 contributes negligibly to k obs 2 , and k obs 2 % k app 2 , and 2 E A (obs) % 2 E A (app) (for the k abbreviations see Scheme 1; for the values of k¢ see [24]). In our experiments we used Tris/HCl buffer, which is known for its pronounced temperature/pH-dependence ()0.026 pH units per °C). To determine the activation energy of self-adenylation, the reaction temperature was varied between 10 and 30 °C, and pH therefore drifted by % 0.5 unit. We avoided using different buffering systems because the kinetic parameters depend on the choice of buffer. For example, the use of Tris/maleimide instead of Tris decreases the adenylation rate % 1.5-fold (not shown). pH-dependence It is known that the adenylation of T4 DNA ligase strongly depends on pH [18], because of the protonation of the catalytic Lys159. To take into account the temperature- induced pH drift of Tris/HCl buffer, we determined k obs 2 at a fixed temperature (20 °C) and different pH (6–9.5). In this set of experiments, we used Tris/HCl buffer in part out of its useful pH range, and special care was taken to measure pH directly in order to account for the weakly buffering components such as ATP, ligase and Mg 2+ (see Experi- mental procedures). The stopped-flow traces recorded at different pH and fitted values of k obs 2 are shown in Fig. 6. Eqn (4) was fitted to the experimental data: k obs 2 ðpH x Þ%k app 2 pH x ðÞ¼k app 2 K a K a þ 10 ÀpH ð4Þ (pH x ) is the pH-dependent rate constant of the forma- tion of the enzyme–AMP adduct, k app 2 is the pH-inde- pendent rate constant, and K a is the protonation constant of the 6-ammonium group of the catalytic lysine residue. Interestingly enough, the determined pK a ¼ 9.8 ± 0.3 for the protonation of the Lys159 is more than 1.2 pH units higher than obtained in the equilibrium binding studies [18] and 1 pH unit lower than the pK a for the e-amino group of lysine in solution [28]. This discrepancy between the results in [18] and our data could possibly be explained by the difference in the experimental conditions: buffer system (Tris/HCl in our case vs. Ches, Taps and Hepes for [18]); concentration of Mg 2+ (5 m M vs. 1 m M ); reaction tem- perature (20 °Cvs.25°C). In the next set of experiments, we determined the k app 2 values at different temperatures (10–30 °C) and pH ¼ 7.6 at 20 °C. Corresponding kinetic traces are shown in Fig. 7. To correct the k app 2 values obtained for the temperature- induced pH drift, we employed the results of pH-depend- ence studies shown in Fig. 6. The relation used for this purpose was as follows: Fig. 6. pH-dependence of the self-adenylation of T4 DNA ligase. Left: kinetic traces obtained at different pH (values are shown next to each trace). Right: the corresponding values of the observed rate constant k obs 2 . The solid trace was computed by weighted fitting using Eqn (4), yielding values for the apparent pK a for the catalytic lysine residue and pH-independent k app 2 showninthegraph.Thereactionwasstartedby rapid mixing of Mg 2+ solution with EÁATP complex pre-equilibrated at the desired pH, resulting in the following final concentrations of components: 2.7 ± 0.1 l M ligase, 67.35 ± 0.07 l M ATP, 5±0.05m M Mg 2+ and pH values of 6.9, 7.05, 7.32, 7.48, 7.76, 7.99, 8.28, 8.6, 8.79 and 9.2. Fig. 7. Kinetics of the self-adenylation of T4 DNA ligase. Fluorescence emission traces were recorded at different temperatures (values are shown in the graph). The reaction was started by rapid mixing of Mg 2+ solution with EÁATP complex pre-equilibrated in buffer A for 5 min at temperatures of 11.3, 13, 14.8, 16.5, 18.4, 20.2, 22.1, 24, 26, and 27.6 °C, resulting in final concentrations of components: 2.6 ± 0.1 l M ligase, 71.42 ± 0.06 l M ATP, 5 ± 0.05 m M Mg 2+ . Ó FEBS 2003 Kinetics of nick sealing by T4 DNA ligase (Eur. J. Biochem. 270) 4321 k app 2 ðT x Þ k app 2 20  CðÞ ¼ K a þ 10 À7:6 K a þ 10 À7:6þ0:026ðT x À20Þ ð5Þ k app 2 (T x ) is the apparent rate constant at certain tem- perature T x , k app 2 (20 °C) is the value at 20 °C, 7.6 is the pH of buffer A at 20 °C, and 0.026 is the pH drift of Tris/HCl buffer per °C. The Arrhenius plot of k app 2 , corrected for the temperature- induced pH changes is shown in Fig. 8. The following equation was used to fit the experimental data points: ln k app 2 ÀÁ ¼ ln j ekT h  À DS # 2 R À 2 E A RT ð6Þ The plot is essentially linear with the slope of 16.7 ± 0.3 kcalÆmol )1 (Table 1). Combining this value with the reaction constants determined in [24], the Mg 2+ -depend- ent and pH-independent rate constant of adenylyl transfer at 37 °C can be estimated as 10 3 )10 4 s )1 . This indicates that at physiological pH, temperature and [Mg 2+ ], T4 DNA ligase binds ATP under strongly suboptimal conditions, resulting in two orders of mag- nitude lower reaction rates. Partial rates and activation energies of T4 DNA ligase catalysis Joining of nicked dsDNA by T4 DNA ligase involves three catalytic steps: formation of the enzyme–adenylate, forma- tion of the ndsDNA–adenylate, and sealing of the nick. These processes were studied at several temperatures, to estimate the activation energies of the individual reaction steps under the assumption that binding of the substrate(s) and dissociation of the product(s) do not limit the rate of catalysis. Formation of the E–AMP intermediate was studied using the stopped-flow technique. Synthesis of AMP–ndsDNA and sealing of the nick were monitored using [ 32 P]ATP and/or Cy5-labeled DNA. All three proces- ses yield essentially linear Arrhenius plots in the temperature range +4 to +30 °C (Fig. 8). Nonlinearity for the joining of the mismatching oligonucleotide M5C19 at high tem- peratures may be the result of the melting of the duplex AMP–M5C19ÁBÁC. Formation of the E–AMP intermediate is the fastest measured process at all temperatures studied with a rate constant more than 10-fold higher than sealing of the complementary ndsDNA. Both adenylation of T4 DNA ligase and sealing of the complementary nick have reason- ably high activation energies, which are identical within the experimental error (Table 1). In terms of the transition-state theory, the marked differences between the observed rates of these reactions (i.e. adenylation of the ligase and nick sealing) arise because of differences in the activation entropies (Table 1). The apparent activation energy for transadenylation of ndsDNA is % 15 kcalÆmol )1 lower than for adenylation of the ligase and/or sealing of the nick (Table 1; determined for the mismatching nick M5C19ÁBÁC). It implies that the transfer of AMP to the terminal DNA phosphate is a thermodynamically sponta- neous reaction, and that the T4 DNA ligase–AMP complex is a high-energy intermediate, as suggested in [18]. Kinetics of T4 DNA ligase catalysis In a simple description, the nick-joining activity of T4 DNA ligase is a three-step enzymatic reaction which involves ndsDNA, ATP and an inorganic cofactor Mg 2+ .The reaction proceeds according to a Ping-Pong mechanism via formation of two intermediate products: E–AMP and AMP–ndsDNA [1,10]. In this work we observed the following phenomena: (a) biphasic kinetics of the nick-sealing, especially pronounced at low [ATP] (Figs 1, 2 and 4); (b) increase in the amplitude of the burst-ligation phase at low [ATP] (Fig. 4); (c) different [Mg 2+ ]-dependence of each kinetic phase (Fig. 5); (d) decrease in the end-joining rate at high and/or low [ATP] (Fig. 3). To take these observations into account, we produced Scheme 2. Biphasic kinetics. According to Scheme 2, the initial burst- ligation phase results from the ÔprocessiveÕ ligation (route 1 -…-6–7). In parallel to the ÔprocessiveÕ ligation, a fraction of ligase molecules enters a nonproductive adenylation cycle (route 1 -…-6–1), aborting the catalysis between the steps of transadenylation and nick-sealing. Abortive adenylation leads to the build-up of the AMP–ndsDNA pool and to the Fig. 8. Arrhenius plot of the individual steps of T4 DNA ligase catalysis: self-adenylation of the enzyme, transadenylation of the nick, and the end- joining. Trace 1, self-adenylation of the ligase. The values of the observed rate constant k obs 2 were corrected for the temperature-induced pH drift of the Tris/HCl buffer using Eqn (5). Traces 2, joining of C6 to 72mer/24mer BÁC. C6 to (BÁC)ratiosare1:1;3:1;10:1;30:1,and 100 : 1. Trace 3, joining of C24 to 72mer/24mer BÁC. C24 to (BÁC) ratio is 30 : 1. Trace 4, adenylation of M5C19 in complex with 72mer/ 24mer BÁC. M5C19 to (BÁC) ratio is 2 : 1. Trace 5, joining of M5C19 to 72mer/24mer BÁC. M5C19 to (BÁ C) ratio is 2 : 1. The dotted traces were obtained by fitting Eqn (6) to the experimental data points. In the case of trace 3 (joining of C24), and trace 5 (joining of M5C19), several data points obtained at high temperatures were omitted to account for melting of the DNA duplex. 4322 A. V. Cherepanov and S. de Vries (Eur. J. Biochem. 270) Ó FEBS 2003 slowing of the ligation rate, because the adenylated enzyme (4) does not seal preadenylated DNA. The slow ligation phase starts when the concentration of AMP–ndsDNA reaches its maximum (steady-state conditions, Fig. 2, 20–100 min). TherelativeratiooftheÔprocessiveÕ nick-sealing and the abortive adenylation depends on the quality of the ndsDNA substrate. In the case of a complementary nick, ligase forms high-affinity complexes with the adenylated ndsDNAs [5], and the kinetic contribution of the adenylation cycle to the overall ligation is minor. EÁAMP–ndsDNA complexes are less stable in the case of a 5¢-mismatching nick [12,14], and the contribution of the adenylation cycle pathway is more significant. For example, the complex of ligase with AMP- M5C19ÁBÁC is unstable, transadenylation of M5C19 is rapid (% 1min )1 ), and the burst ligation is slow (0.2 min )1 ) (Table 1). As a result, the slow ligation phase starts only when all available ndsDNA substrate is converted: it is either adenylated (via 5–6–1), or joined (via 5–6–7), and, at the same time, the dominant enzyme fraction (4)isAMP bound (Fig. 2, 20–100 min). The adenylation of ligase is reversible: there is always a fraction of the free enzyme (1) in the reaction mixture formed via the routes 4–3–2–1, or 4–11–1). According to Scheme 2, the slow ligation of M5C19 is performed by this enzyme fraction via the route 1–6–7. Increase in amplitude of the burst phase at low [ATP]. At high [ATP] (1–5 m M ), free ligase (1) binds ATP (1–2) faster than it binds DNA (1–6) [(1–5) · 10 3 s )1 for ATP vs. 10 2 s )1 for 1 l M ndsDNA]. The repetitive abortive cycling (1–2 -…-6–1) leads to accumulation of the AMP–ndsDNA intermediate, and its removal via the ÔnonprocessiveÕ route 1–6–7 is kinetically insignificant. ÔNonprocessiveÕ ligation 1–6–7 becomes significant only during the slow ligation phase, when all available ndsDNA substrate is adenylated, and the complexes 2–5 are no longer productive. At low [ATP] (< 40 l M ), ligase binds ATP slower than DNA (< 36 s )1 for ATP vs. 10 2 s )1 for 1 l M ndsDNA). During burst ligation, the free ligase (1) is thus engaged in both ÔprocessiveÕ ligation (1–2 -…-6–7)andtheÔnonproces- siveÕ scavenging of the preadenylated nicks via the route 1–6–7. ÔNonprocessiveÕ nick sealing (1–6–7) slows down the accumulation of AMP–ndsDNA, and the start of the slow ligation is delayed. Delay of the slow ligation results in an increase in the amplitude of the burst-ligation phase (for M5C19 more than 10-fold; Fig. 4). Similar results were recently reported [15] when a large number of the dsDNA substrates with one or two mismatching base pairs on both sides of the nick opposite tandem canonical bases were tested for ligation by T4 DNA ligase. There, the highest ligation efficiency was observed at [ATP] of 10–100 l M . [Mg 2+ ]-dependence. The fact that the two ligation phases have different [Mg 2+ ] optima stems, in terms of Scheme 2, from the [Mg 2+ ]-regulated redistribution between the two enzyme forms: the adenylated ligase (4), and the free enzyme (1). E–AMPÁMg (4) is engaged in the ÔprocessiveÕ ligation (4 -…-7). An increase in [Mg 2+ ] stimulates self-adenylation 3 fi 4, and inhibits the reverse reaction 4 fi 3 [24], causing the increase in 4, and, accordingly, the increase in the burst-ligation rate. On the other hand, slow ÔnonprocessiveÕ ligation is performed by the free ligase (1) (route 1–6–7). The reverse reaction 4 fi 3 is the most efficient at % 1–3 m M [Mg 2+ ] [24], causing the increase in 1. As a result, the rate of the slow phase reaches its maximum at this [Mg 2+ ]. When mismatching nicks containing tandem canonical bases at the site of ligation are sealed, the same narrow optimum of [Mg 2+ ]between1and3m M has been reported [15]. Decrease in end-joining rate at high and/or low [ATP]. According to Scheme 2, the inhibition of ligation at high [ATP] occurs because T4 DNA ligase binds the nucleotide at the dsDNA-binding site [20] with K d between 0.1 and 0.25 m M [21] (routes 2–9, 3–10, and 4–11).Thedecreasein the rate of ligation at low [ATP] occurs because ndsDNA forms a complex with the ligase (route 1–8). From the modeling studies [26,27] one may conclude that, if the ligase in complex (8) binds ATP at all, it will do so at a reduced rate. The structural considerations are that ndsDNA in complex with T4 DNA ligase prevents access of ATP to the nucleotide-binding pocket of the enzyme, preventing ATP from either leaving or binding to the active site. This is reflected in Scheme 2: ligase in complex with (n)dsDNA does not bind ATP at all. Scheme 2. Nick-joining by T4 DNA ligase. The rate constants k 1 –k 3 correspond to the three steps of covalent catalysis. Ó FEBS 2003 Kinetics of nick sealing by T4 DNA ligase (Eur. J. Biochem. 270) 4323 Scheme 2 only quantitatively addresses the AMP- dependent reversal or inhibition of ligation. Another important assumption is that the ssDNA fragments to be joined do not dissociate from the opposite DNA strand or the EÁdsDNA complex. The latter is certainly not the case when short (4–12-mer) oligonucleotides are joined near their T m . We further assumed that ndsDNA is present in the Mg 2+ -coordinated form, neglecting the exchange between the dsDNA-bound Mg 2+ and K + or Na + at elevated concentrations of the latter ions (> 100 m M ). High con- centrations of K + and Na + inhibit DNA ligases [10,12, 29–31], perhaps, among other reasons, because ndsDNA changes to the K + -(Na + )-coordinated form. In summary, this kinetic scheme of T4 DNA ligase catalysis includes a two-metal-ion mechanism of ligase adenylation, binding of the second nucleotide molecule at the DNA-binding site, and synthesis of dinucleoside tetra- phosphates, and treats the reaction in terms of ÔprocessiveÕ burst ligation and ÔnonprocessiveÕ nick sealing. Physiological relevance of joining of mismatching nicks by T4 DNA ligase In vitro pre-steady-state kinetic studies performed in this work suggest that T4 DNA ligase could rapidly adenylate mismatching nicks in the infected cells, either at the late stages of degradation of the cellular DNA or during the subsequent replication of the coliphage T4 DNA: the ligase gene is transcribed at steady levels throughout the eclipse, reaching its maximum 3 min after infection [32]. The sealing of these pre-adenylated nicks, however, would be slow because of relatively high intracellular [ATP] (1–3 m M )[33]. Under these circumstances, T4 DNA ligase would act on the mismatching nicks more like an mRNA capping enzyme, a member of the same superfamily of nucleotidyl- transferases [34,35]. In the cell, capping of the 5¢-end of mRNA ensures protection from degradation by specific exonucleases [36,37]. In the case of capping the mismatching nick, however, similar reasoning does not seem logical. One should consider that, in the cell at relatively high ionic strength, both adenylation and joining of the mismatching nicks could be suppressed to a large extent, as in the case of in vitro ligations at % 0.2 M NaCl [12,16,17]. Without supportive experimental data in vivo, we will refrain from assigning any physiological meaning to the ÔlowÕ substrate specificity of T4 DNA ligase reported here and in earlier contributions [14,15,23]. Instead, we demonstrate that this ÔunconventionalÕ activity of T4 enzyme is an invaluable tool for elucidating the general kinetic mechanism of DNA ligase catalysis. Conclusion T4 DNA ligase-promoted end joining of nicked DNA is a superimposition of two processes. During the burst phase, the main enzyme fraction performs ligation ÔprocessivelyÕ, i.e. by transadenylating ndsDNA and sealing the nick without dissociation from the complex. In parallel, a fraction of the ligase molecules dissociates after transadenylation, and rebinds ATP. 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HP2O3 À , ADP, and CDP J Am Chem Soc 94, 9198–9204 7 39 Shack, J & Bynum, B.S (1959) Determination of the interaction of deoxyribonucleate and magnesium ions by means of a metal ion indicator Nature (London) 184, 635–636 40 Cavalieri, L.F (1951) Studies on the structure of nucleic acids V On the mechanism of metal enzyme interactions J Am Chem Soc 74, 1242–1247 . of Mg 2+ was 5 m M . [dsDNA] (1 l M ) corresponds to 10 enzyme turnovers (T4 DNA ligase was 0.1 l M ). Ó FEBS 2003 Kinetics of nick sealing by T4 DNA ligase. orders of mag- nitude lower reaction rates. Partial rates and activation energies of T4 DNA ligase catalysis Joining of nicked dsDNA by T4 DNA ligase involves