Dissection of the ATP-driven reaction cycle of the bacteriophage T4 DNA replication processivity clamp loading system1

doi:10.1006/jmbi.2001.4687 available online at http://www.idealibrary.com on

J. Mol. Biol. (2001) 309, 869±891

Dissection of the ATP-driven Reaction Cycle of the Bacteriophage T4 DNA Replication Processivity Clamp Loading System Paola Pietroni, Mark C. Young, Gary J. Latham and Peter H. von Hippel* Institute of Molecular Biology and Department of Chemistry University of Oregon, Eugene OR, 97403-1229, USA

Processive DNA replication requires the loading of a multisubunit ringshaped protein complex, known as a sliding or processivity clamp, onto the primer-template (p/t) DNA. This clamp then binds to the replication polymerase to form a processive polymerase holoenzyme. The processivity of the holoenzyme derives from the topological properties of the clamp, which encircles the DNA without actually binding to it. Multisubunit complexes known as clamp-loaders utilize ATP to drive the placement of this ring around the DNA. To further understand the role of ATP binding and hydrolysis in driving clamp-loading in the DNA replication system of bacteriophage T4, we report the results of a series of presteady-state and steady-state kinetic ATPase experiments involving the various components of the reconstituted system. The results obtained are consistent with a mechanism in which a slow step, which involves the binary ATP-bound clamp-clamp loader complex, activates this complex and permits p/t DNA to bind and stimulate ATP hydrolysis. ATP hydrolysis itself, as well as the subsequent (after clamp-loading) dissociation of the clamp-loader and the slippage of the loaded clamp from the p/t DNA construct, are shown to be fast steps. A second slow step occurs after ATP hydrolysis. This step involves the dissociated clamp loader complex and may re¯ect ADP release. Only one molecule of ATP is hydrolyzed per clamp-loading event. Rate constants for each step, and an overall reaction mechanism for the T4 clamp-loading system, are derived from these data and from other results in the literature. The principles that emerge ®t into a general framework that can apply to many biological processes involving ATP-driven reaction cycles. # 2001 Academic Press

*Corresponding author

Keywords: DNA replication; polymerase accessory proteins; ATPase; presteady-state kinetics; ATPase reaction cycles

Introduction Present addresses: M. C. Young, Of®ce of Information Systems, University of Puget Sound, 1500 North Warner St., Tacoma, WA 98416, USA; G. J. Latham, Research Division, Ambion, Inc., 2130 Woodward St., Austin, TX 78744, USA. Abbreviations used: gp44/62, a complex of the protein products of phage T4 genes 44 and 62, with a stoichiometry of four gene 44 protein subunits to one gene 62 protein subunit; gp45, the trimer of the protein product of T4 gene 45; p/t, primer/template; ATPgS, adenosine 50 -O-(3-thiophosphate); TLC, thin-layer chromatography. E-mail address of the corresponding author: [email protected] 0022-2836/01/040869±23 $35.00/0

Organisms ranging from bacteriophages through prokaryotes to eukaryotes share a common mechanism to achieve processive replication of their genomic DNA. The catalytic core of the machinery involved in the replication process is the polymerase, which is topologically linked to its DNA template by attachment to a ring-shaped multisubunit protein complex that completely encircles the DNA.1 ± 3 As a class, these ring-shaped proteins are known as sliding clamps4,5 because they ``clamp'' the relevant polymerases to the template DNA. However, these clamps do not bind to the DNA directly. Rather they are linked to it only topologically, thus leaving the tethered polymerases free to # 2001 Academic Press

870 ``slide'' along the DNA while catalyzing processive primer extension DNA synthesis.6 These sliding clamps are assembled around DNA by the action of multisubunit clamp-loader complexes, which utilize ATP to drive the loading function. These clamp loaders act catalytically; after placing a clamp on the DNA, they dissociate and again become available for a subsequent clamp-loading process.7 ± 9 The sliding clamp of the bacteriophage T4 system is a homotrimer of gp45 subunits3,10 and the clamp-loader is a tightly associated complex containing one gp62 and four gp44 subunits.10 Recently, the gp44/62 complex has been reconstituted in vitro from puri®ed subunits.11 The intrinsic ATPase activity of the gp44/62 complex itself is very low; it is stimulated many-fold, and in a synergistic manner, by both gp45 and p/t DNA.12,13 The gp44 tetramer carries the ATP-binding sites of the complex (one per subunit) and is activated directly by DNA cofactor binding, whereas gp62 is required to trigger the stimulation of the gp44/62 ATPase activity by the gp45 sliding clamp.10,14 Several studies had suggested that ATP binding is suf®cient to assemble the gp44/62-gp45 protein complex onto primer-template (p/t) DNA, but that interaction of the polymerase with the sliding clamp to achieve processive holoenzyme formation absolutely requires ATP hydrolysis.9,15 ± 17 However, two ¯uorescence studies, one conducted in our laboratory18 and the other in the Benkovic laboratory,19 suggested that ATP hydrolysis is required for the formation of a stable gp44/62gp45 complex in which gp45 had undergone a conformational change, presumably related to ring opening. The ¯uorescence change associated with this process was not observed when the nonhydrolyzable analogue ATPgS was used in place of ATP. Addition of p/t DNA reversed the ATPinduced ¯uorescence change in the gp44/62-gp45 complex, suggesting that dissociation of the clamploading complex accompanies the loading of the sliding clamp onto the p/t DNA. Thus, both of these ¯uorescence studies suggested that ATP hydrolysis might be required to ``activate'' the gp44/62-gp45 complex and therefore that p/t DNA might bind to the activated loading complex complex after ATP hydrolysis. In a subsequent study, we used a photoactivatable crosslinker to look for conformational changes within the accessory proteins complex during clamp binding and loading.20 These crosslinking studies revealed that ATP and ATPgS induce very similar conformational changes in the gp44/62gp45 complex, while the conformational change induced by ADP differs. These ®nding suggested that, in the absence of p/t DNA, ATP hydrolysis by the gp44/62-gp45 complex might be slow and release of the ADP hydrolysis product might be fast. On the other hand, since p/t DNA binding seemed to reverse the ATP-induced conformational change, it appeared that the addition of p/t DNA

Kinetics of Loading the T4 Processivity Clamp

might accelerate ATP hydrolysis, resulting in a relatively short lifetime for the ATP-bound state. Thus, our crosslinking results suggested a different loading pathway than that inferred from the ¯uorescence studies, with p/t DNA entering the gp44/62-gp45 complex before, rather than after, ATP hydrolysis. In order to establish de®nitively whether ATP hydrolysis occurs before or after p/t DNA binding, and to identify the rate-limiting step (or steps) during the ATP hydrolysis cycle in the presence and absence of p/t DNA, we have performed a series of presteady-state and steady-state kinetic experiments. The results of these studies are reported here. Previous kinetic studies led to the following proposed reaction cycle for the clamp-loading process in the T4 DNA replication system.21,22 Sedimentation studies had shown that the gp45 trimer ring displays an unexpectedly high degree of structural asymmetry in hydrodynamic experiments, suggesting that the gp45 ring might be partially open when free in solution.10,23 Kinetic studies then suggested that a slow conformational change occurs in the presence of ATP after the binding of the clamp to the gp44/62 complex, and that this conformational change results in the further opening of the ring. This conformational change appeared to be linked to the hydrolysis of two molecules of ATP. It was proposed that the next step was to bind the clamp loader complex carrying the open ring to the DNA, with this binding inducing ring closure and the subsequent hydrolysis of two more ATP molecules. This second ATP hydrolysis event was considered to be rate limiting in the loading process. The overall proposed clamp-loading cycle concluded with the release of gp44/62 from the overall complex, leaving the ring assembled around the DNA and available for interaction with the replication polymerase.21,22 The clamp-loading process in Escherichia coli has been found to be quite different from that suggested above for the T4 system. In the E. coli system, ATP binding induces a conformational change in the clamp loader that allows interaction with the clamp. The E. coli clamp seems to be fully closed in solution, and a slow conformational change that seems to result in ring opening is observed on binding the clamp to the ATP-bound clamp-loader.24 ± 26 This conformational change occurs before DNA binding and ATP hydrolysis, and represents the rate-limiting step in the loading process.27,28 The ATP-bound open clamp-clamp loader complex of E. coli then binds rapidly to p/t DNA,26,28,29 and this binding is followed by the hydrolysis of one or two molecules of ATP.30,31 Finally, the E. coli clamp loader dissociates from the ternary structure, leaving the clamp encircling the DNA.5,26,28,29 The rate-limiting step in this process had been proposed to be a slow step between the sequential hydrolysis of two ATP molecules.30 However, a more recent study has suggested that the two ATP molecules are hydrolyzed rapidly

Kinetics of Loading the T4 Processivity Clamp

and simultaneously, and that the rate-limiting step in the ATP hydrolysis cycle occurs in solution after the clamp loader has dissociated from the loaded clamp, probably re¯ecting slow ADP release.31 The presteady-state and steady-state kinetic experiments with the T4 clamp-loading system that are described here are more congruent with the mechanisms described recently for the E. coli system than with the reaction cycle previously proposed for T421,22 that is outlined above. Our experiments show clearly that a slow conformational change in the binary clamp-clamp loader complex occurs before hydrolysis and limits the rate of the clamp-loading process, with p/t DNA then also binding to this ``activated'' complex prior to ATP hydrolysis. The subsequent hydrolysis of ATP and dissociation of the clamp loader from the clamp and the p/t DNA now turn out to be relatively fast, whereas a second slow step, possibly related to ADP release, occurs after clamp-loading is complete and the clamp loader has dissociated.

Results ATP hydrolysis, or a step before hydrolysis, is at least partially rate-limiting during the loading of the gp45 clamp onto p/t DNA

871 burst phases that are seen in the presence of p/t DNA will be discussed below), indicating that ATP hydrolysis, or a step before hydrolysis, is relatively slow and at least partially rate-limiting for this system. In the absence of p/t DNA, the gp44/62-gp45 protein complex (Figure 1, open circles) hydrolyzes ATP at a rate of 0.1 mM sÿ1 per mM gp44/62 complex while, in the presence of p/t DNA, ATP is hydrolyzed at a rate of 5.5 mM sÿ1 per mM gp44/62 complex. Thus the step that limits the ATP hydrolysis reaction in the absence of p/t DNA is accelerated  55-fold when p/t DNA is added. However, despite this substantial rate stimulation, the overall kinetic pro®le remains linear, indicating that the rate-limiting step in the absence of p/t DNA is not accelerated to the point that a step after hydrolysis becomes rate-limiting (because then biphasic kinetics would be observed). Since we know that the hydrolysis of ATP in the presence of p/t DNA results in clamp-loading onto the DNA,9,15 ± 17 we must conclude from these results that, in addition to release of the products of ATP hydrolysis (ADP and Pi), the steps corresponding to the dissociation of gp44/62 from gp45 and from the p/t DNA, and the sliding of the loaded gp45 clamp off the ends of the short linear p/t DNA, must all be relatively fast processes that

Rapid quench experiments In a presteady-state kinetics investigation of ATPase mechanisms, the reaction is quenched at very early times (milliseconds), allowing detection of the very ®rst ATP hydrolysis cycle. If the steady-state rate is limited by a step that follows the chemical hydrolysis step (for example, product release), the ATPase activity measured under presteady-state conditions and plotted as a function of time will display biphasic kinetics that are characterized by an initial burst of product formation corresponding to the ®rst ATP hydrolysis reaction, followed by a slower steady-state phase in which a slower step not involved in the ®rst kinetic phase (e.g. product release) limits the rate of the subsequent hydrolysis cycles. Conversely, when steps after hydrolysis are fast relative to the chemical hydrolysis step or to a step before hydrolysis, then all the ATP hydrolysis cycles, including the very ®rst, occur at identical rates, resulting in an overall linear time-course for the reaction that corresponds to the steady-state rate throughout. Presteady-state ATPase experiments were performed to identify the rate-limiting step for the reaction in which ATP is hydrolyzed by the gp44/ 62-gp45 complex in the absence and in the presence of p/t DNA. Figure 1 shows the presteadystate time-course of ATP hydrolysis catalyzed by 2 mM gp44/62 in the presence of 3 mM gp45 and in either the absence (open circles) or the presence (®lled circles) of 2.4 mM p/t DNA. It is apparent that both time-courses display approximately linear kinetics (the very small lag and subsequent

Figure 1. The ATP hydrolysis step, or a step before hydrolysis, is at least partially rate-limiting during the stimulation of the gp44/62 ATPase activity by gp45 and p/t DNA. The presteady-state time-courses of ATP hydrolysis by gp44/62 and gp45 in the presence (®lled circles) and absence (open circles) of p/t DNA were determined as described in Experimental Procedures. Final component concentrations were 2 mM gp44/62, 3 mM gp45, 2.4 mM 30/50mer p/t DNA (when present), and 500 mM ATP. An overall linear pro®le was observed in both time-courses, with a rate of 5.5 mM sÿ1 per mM gp44/62 in the presence of both gp45 and p/t DNA, and a rate of 0.1 mM sÿ1 per mM gp44/62 when only gp45 was present.

872 do not limit the overall rate of the ATPase reaction cycle.{ Therefore, assuming that steps involved in the assembly of the gp45-gp44/62 complex are also relatively fast, the overall rate of the clamploading process must be limited either by the chemical hydrolysis step itself or by a conformational change in the loading complex prior to hydrolysis that could involve: (i) the ternary reaction complex (gp45-gp44/62-p/t DNA); (ii) a binary complex (gp45-gp44/62 or gp44/62-p/t DNA) that must be ``activated'' prior to assembly of the complete loading complex; or (iii) the gp44/ 62 clamp-loading complex alone. In any case, the ternary complex must be formed before ATP hydrolysis occurs in the overall reaction cycle. This follows because a burst phase is not observed when gp44/62 is mixed with gp45 in the absence of DNA, nor is a burst phase observed when gp44/62 is mixed with p/t DNA in the absence of gp45 (data not shown). Therefore, we conclude that both reaction cofactors, i.e. gp45 and p/t DNA, must combine with the gp44/62 clamploading complex before the ATP hydrolysis reaction takes place. Pulse-chase experiments The lack of a burst phase in the rapid-quench experiments prevented us from determining the number of ATP molecules hydrolyzed by the gp44/62 complex during the loading of gp45 onto DNA. Had the ®rst hydrolysis cycle been faster than the subsequent cycles, then the amplitude of the burst phase would have corresponded to the number of ATP molecules cleaved in the ®rst hydrolysis cycle and, presumably, in all that follow. Pulse-chase experiments were performed to address this issue in another way. In a pulse-chase experiment the enzyme is ®rst mixed with radiolabeled substrate and then, after varying times, is mixed with excess unlabeled ATP instead of being inactivated with acid as in a rapid-quench experiment. After incubation for an additional period of time suf®cient to convert all of the labeled enzymebound substrate to product (six to ten half-lives of the steady-state turnover rate), the reaction was stopped by quenching with acid. During the chase period, the enzyme hydrolyzes only the fraction of the radiolabeled ATP that was tightly bound to the enzyme at the initiation of the chase. More loosely bound radiolabeled substrate { We note that the short linear p/t DNA constructs used in this study have no blocking groups at the ends to prevent ``sliding off'' of the closed circular gp45 clamps after loading, and, as shown here, this sliding off process is very rapid. Kaboord and co-workers32 have shown that such sliding off can be blocked effectively by incorporating biotin-streptavidin groups at the ends of the p/t DNA construct. However, in this study we have used the p/t DNA ``catalytically'' in our dissection of the overall clamp-loading cycle, and thus have not used blocked DNA constructs.

Kinetics of Loading the T4 Processivity Clamp

Figure 2. Only one ATP molecule is hydrolyzed per loading complex during the ATPase reaction cycle in the presence of both gp45 and p/t DNA. Pulse-chase experiments, carried out as described in Experimental Procedures, were run at two different ®nal concentrations of clamp-loading complex and p/t DNA. The ®rst set contained 2 mM gp44/62, 3 mM gp45, 2.4 mM p/t DNA, and 500 mM ATP (®lled circles). The second set contained 1 mM gp44/62, 1.5 mM gp45, 1.2 mM p/t DNA, and 500 mM ATP (open circles). The burst amplitude, which re¯ects the number of ATP molecules bound and eventually hydrolyzed during the chase in the ®rst ATP turnover reaction, is proportional to the enzyme concentration and corresponds to 1.9 mM ATP for the 2 mM gp44/62 reaction, and to 1 mM ATP for the 1 mM gp44/ 62 reaction. The measured steady-state rate was 10.5 mMÿ1 sÿ1 and 5 mMÿ1 sÿ1 for the 2 mM and 1 mM gp44/62 reactions, respectively.

may be released before hydrolysis during the chase period, and would then be replaced by unlabeled ATP. Thus, the amount of radiolabeled product that is measured at the end of the chase corresponds to the fraction of labeled substrate that was hydrolyzed before the unlabeled chase ATP was added, together with the fraction that was bound suf®ciently tightly to the enzyme to be hydrolyzed during the chase period rather than lost by exchange with unlabeled ATP. This means that hydrolyzed radiolabeled ATP can be detected in a pulse-chase experiment when the enzyme in the initial reaction has been mixed with radiolabeled substrate for a time suf®cient only to bind ATP, but not to hydrolyze it, since time is available for hydrolysis of the bound ATP during the chase period. Therefore, the rate of development of the initial burst phase in this type of experiment measures the rate of ATP binding, whereas in a rapid-quench experiment it measures the rate of ATP hydrolysis. Furthermore, the amplitude of the normalized burst phase in such a pulse-chase experiment corresponds to the total

Kinetics of Loading the T4 Processivity Clamp

number of ATP molecules that are bound (and eventually hydrolyzed) by the enzyme in a single ATPase reaction cycle. On the other hand, the linear phase of the reaction in a pulse-chase experiment re¯ects the steady-state rate of ATP hydrolysis, just as it does in a rapid-quench experiment. Since the rate of binding of ATP is always faster than the steady-state hydrolysis rate in the presence of a large excess of substrate, a burst phase is always observed in such a pulse-chase experiment if the rate at which the substrate dissociates from the enzyme is slower than the rate of hydrolysis. As seen in Figure 2, an initial fast burst of ATP hydrolysis is observed when gp44/62 is mixed with radiolabeled ATP in the presence of excess gp45 and DNA for varying times, and then chased with unlabeled ATP. Under the conditions used in this experiment (an initial ATP concentration of 500 mM), the rate of binding of ATP is so fast that even at the earliest observable time-point (8.5 ms) the enzyme is saturated with ATP and only the linear steady-state reaction is observed. The steady-state rate measured in these pulse-chase experiments is similar to that observed in the rapid-quench experiment (Figure 1, ®lled circles), and corresponds to 10.5 mM sÿ1 when 2 mM gp44/ 62 complex was used (Figure 2, ®lled circles), and to 5 mM sÿ1 for 1 mM gp44/62 (Figure 2, open circles). The burst amplitude also depends on the concentration of gp44/62 used, and is 1.9 mM for the 2 mM gp44/62 reaction (®lled circles), and 1 mM for the 1 mM gp44/62 reaction (open circles). This experiment ®nally permits us to conclude that the amplitude of the burst phase in these pulse-chase reactions corresponds to the binding and subsequent hydrolysis of only one molecule of ATP per reaction cycle catalyzed by the gp44/62 clamp loader complex.{ In order to determine the rate constant for ATP binding to the gp44/62 complex, the concentration of ATP that was added to the reaction was lowered and the experiment was repeated at 100 mM and 60 mM ATP. The results are presented in Figure 3 and show that the time-course of the exponential phase corresponding to ATP binding can be resolved and measured at these reduced ATP concentrations. A ®rst-order binding rate of { Of course, the amplitude of the burst phase of radiolabeled ATP in such experiments will be decreased if the rate of dissociation of the bound ATP from the enzyme is fast relative to the rate of hydrolysis, and if the concentration of substrate is not suf®cient to saturate the enzyme. Both of these issues are investigated below, and are shown not to be a problem in this experiment. Nevertheless, we point out here that both of these potential artifacts, if present, would reduce, rather than increase, the observed amplitude of the burst phase, and a reduced burst amplitude caused by these artifacts would lead to an under-estimate of the number of ATPs bound and hydrolyzed by the gp44/62 complex per clamp-loading reaction cycle.

873

Figure 3. The bimolecular rate constant for the binding of ATP to gp44/62 (in the presence of gp45 and p/t DNA) is 7  105 Mÿ1 sÿ1. Pulse-chase experiments with gp44/62 in the presence of both gp45 and DNA at ATP concentrations of 100 mM (®lled circles) and 60 mM (open circles) were conducted as described in Experimental Procedures. The ®nal concentrations of protein and p/t DNA after mixing were 2 mM gp44/62, 3 mM gp45, and 2.4 mM p/t DNA. The burst rate re¯ects the rate of ATP binding, which is 73(10) sÿ1 in the 100 mM ATP reaction and at 45(15) sÿ1 in the 60 mM ATP reaction, yielding a bimolecular rate constant for ATP binding of 7  105 Mÿ1 sÿ1. The burst amplitude, which measures the number of ATP molecules that are bound and eventually hydrolyzed during the chase period in the ®rst ATP turnover cycle, is 1.3 mM at 100 mM ATP and 1 mM at 60 mM ATP, indicating that about half of the enzyme present (2 mM gp44/62 was used) is saturated with substrate at the concentrations of ATP used. Accordingly, the steady-state rates measured in these experiments are about one-half (6.7 mM sÿ1 at 100 mM ATP and 4.6 mM sÿ1 at 60 mM ATP) of the maximal rate (10.5 mM sÿ1 with 2 mM gp44/62) measured with saturating (500 mM) ATP (Figure 2).

73(10) sÿ1 was obtained with 100 mM ATP (Figure 3, ®lled circles) and a rate of 45(15) sÿ1 was measured for 60 mM ATP (open circles). These rates correspond to a calculated bimolecular rate constant for ATP binding of 7  105 Mÿ1 sÿ1. In these experiments, the concentration of gp44/62 was 2 mM, and the extrapolated amplitude of the burst phase was 1 to 1.3 mM (see Figure 3), indicating that approximately one-half of the enzyme was saturated with substrate under these conditions. Accordingly, the steady-state rates observed at these concentrations of ATP (6.7 mM sÿ1 and 4.6 mM sÿ1 at 100 mM and 60 mM ATP, respectively) are about half of the rate seen under saturating concentration of ATP (11 mM sÿ1; Figures 1 and 2, ®lled circles). This ®nding is consistent with the Km value of 100 mM that was measured previously for the steady-state ATPase reaction cata-

874 lyzed by the gp45-gp44/62-p/t DNA complex,13,33,34 and is consistent with our conclusion that only one ATP molecule is hydrolyzed per clamploading reaction cycle by the T4 system. At least two steps, one involving gp45 in a binary complex and the other the gp44/62 complex alone, are partially rate-limiting during the loading of the gp45 clamp onto p/t DNA Steady-state ATPase kinetics The presteady-state kinetic experiments described above did not address the question of whether the entire ternary clamp-loading complex (gp45-p44/62-p/t DNA) is involved in the ratelimiting step during the loading process, or whether the rate-limiting step occurs at an earlier stage of clamp-loader assembly. Thus, a slow conformational change in the gp44/62 complex might be required to activate the loading complex before gp45 and DNA bind and stimulate ATP hydrolysis. Alternatively, and assuming that the rates of binding of gp44/62 to the gp45 clamp and the p/t DNA are fast, a slow conformational change after assembly of two components might be required before p/t DNA or gp45 can join the complex and activate ATP hydrolysis. In another scenario, the assembly of the ternary complex might be fast and the ATPase rate would then be limited by a reaction involving the ternary complex, with the slow step corresponding to either a conformational change involving the entire complex or to a slow step within the chemical hydrolysis reaction itself. Depending on which of the above scenarios applies, the steady-state ATPase activity will be proportional to the concentration of gp44/62 alone, to the concentration of a binary complex, or to the concentration of the ternary complex, respectively. If the rate-limiting step occurs at the level of the ternary complex, titration of either one of the binary complexes, composed of gp44/62 and either gp45 or p/t DNA, with the third component (either DNA or gp45, respectively) should display the same extent of stimulation of the ATPase activity when the limiting component is added, since the concentration of the limiting component, (either gp45 or DNA) would also correspond to the concentration of ternary complex in solution (assuming that all three components bind tightly). Conversely, if a binary complex is involved in the rate-limiting step, then the ternary complex would have a relatively short lifetime and the component that is not part of the rate-limiting step would act catalytically on the ATPase rate. In this case, a substoichiometric concentration of the catalytic component would stimulate the ATPase activity to its maximum level, while the maximal stimulation of the ATPase activity would be reached only at a stoichiometric concentration of the limiting component if the component added becomes part of the rate-limiting binary complex. Finally, both

Kinetics of Loading the T4 Processivity Clamp

cofactors could act catalytically if a conformational change in the gp44/62 complex itself is ratelimiting. The experiments shown in Figure 4 were performed to sort out these possibilities. In these experiments, a solution containing gp44/62, together with an excess of either p/t DNA or gp45, was titrated with either gp45 (®lled circles) or DNA (open circles), respectively, and the steadystate ATPase rate was measured at each titration point and is plotted as a function of the concentration of the titrating component. It is apparent that the stimulation of the ATPase activity follows a very different course for the two experiments, with the titration with gp45 of a solution containing gp44/62 and excess p/t DNA resulting in a weaker stimulation of the ATPase activity than that observed in a comparable experiment with p/t DNA as the titrant. In the latter case, the ATPase rate increases very rapidly, with maximal

Figure 4. Two steps are partially rate-limiting during the ATPase cycle catalyzed by gp44/62 in the presence of both gp45 and p/t DNA, with one involving gp45 in a binary complex and the other the gp44/62 complex alone. Steady-state ATPase titrations were performed as described in Experimental Procedures. gp44/62 (1 mM) in the presence of either excess (2 mM) gp45 (open circles), or excess (1.2 mM) p/t DNA (®lled circles), were titrated with the third component (either p/t DNA or gp45, respectively). p/t DNA stimulates the binary gp44/62-gp45 complex catalytically (open circles). Therefore, p/t DNA is not part of the rate-limiting step. In contrast, gp45 stimulates the ATPase of gp44/62 ``less catalytically'' in the presence of excess p/t DNA (®lled circles). The latter kinetic pro®le displays a stoichiometry of about one-half gp45 molecule per gp44/62 complex, suggesting that besides a slow step involving gp45 in a binary complex (see the text), a second slow step, involving gp44/62 alone, occurs with a similar rate. The continuous curves were derived by ®tting the data to the equations developed in the Appendix.

Kinetics of Loading the T4 Processivity Clamp

stimulation attained at a concentration of p/t DNA that is well below the stoichiometric level. As Figure 4 shows (open circles), the plateau rate for this experiment is effectively reached at a p/t DNA concentration that is at least ten times lower than the 1 mM concentration of gp44/62-gp45 complex present in the reaction. Therefore, p/t DNA acts catalytically in these reactions and cannot be part of the rate-limiting step. In contrast, the approach to the plateau rate occurs over a much wider concentration range for the comparable experiment in which gp45 was used to titrate a mixture of gp44/62 and p/t DNA (®lled circles), suggesting that a slow step involving gp45 must be at least partially rate-limiting under these conditions. We note, however, that even here the titration with gp45 does not proceed entirely stoichiometrically. Fitting the gp45 titration data as a binding reaction con®rms this conclusion and shows that the kinetic pro®le obtained corresponds to a binding stoichiometry of 0.5 gp45 clamp per gp44/62 complex, showing that gp45 also acts somewhat catalytically, although to a much lesser extent than p/t DNA. To rule out the possibility that part of the gp44/62 protein preparation was not fully active in these experiments (which would, of course, alter both the stoichiometry at which maximal stimulation is attained and the calculated number of ATP molecules consumed per reaction cycle), we performed the same titration as that shown in Figure 4 but in the absence of p/t DNA (Figure 5). Previous experiments18,19 had shown that the titration of ¯uorescently labelled gp45 with gp44/62 (in the absence of p/t DNA and in the presence of ATP) demonstrates a 1:1 (stoichiometric) binding of protein complexes. In support of this conclusion, in Figure 5 we titrate the ATPase activity of gp44/62 with gp45 in the absence of p/t DNA. The ATPase rates obtained are, of course, much lower than those measured with the ternary complexes of Figure 4, but the data in this experiment also show a reaction stoichiometry that is very close to one gp45 per gp44/62 complex. Therefore, the catalytic effect of gp45 on the ATPase activity of gp44/62 observed in the presence of p/t DNA, although small, is nevertheless real and not an artifact due to inactive protein in the gp44/62 preparation. Consequently, we must conclude that, in addition to the slow and rate-determining step that involves gp45, a second slow step that involves only gp44/62 must occur with a comparable rate, and must also be partially rate-limiting within the overall clamp-loading reaction cycle. From the steady-state results, we cannot tell whether the observed slow steps occur before or after ATP hydrolysis. However, we know from our { The release of reaction product Pi has been shown to be fast.9,34

875

Figure 5. The stimulation of the ATPase activity of gp44/62 by gp45 is stoichiometric in the absence of p/t DNA. The steady-state ATPase titration was performed as described in Experimental Procedures. gp44/62 (1 mM) (in the absence of p/t DNA) was titrated with gp45. The gp44/62 ATPase rate increases linearly until a stoichiometric concentration of gp45 has been added.

previous presteady-state results (Figure 1, ®lled circles) that at least one slow step must occur before hydrolysis. Moreover, as pointed out above, a careful inspection of this presteady-state rapid-quench experiment (Figure 1, ®lled circles) reveals the presence of both a small lag phase and a small burst phase. This shows that, in addition to the slow step before hydrolysis, a second slow step, with a similar rate, must occur after hydrolysis. The evidence to support this is consistent with a number of possibilities for these partially rate-limiting steps. Thus, the slow step that involves the gp45 clamp could be: (i) a conformational change in the binary gp44/62-gp45 complex before ATP hydrolysis is stimulated by p/t DNA; (ii) a step after hydrolysis (such as the gp44/62-gp45 dissociation reaction); (iii) the sliding of the gp45 ring off of the p/t DNA after the loading event (in this scenario, the ``more catalytic'', relative to gp45, activity of p/t DNA could be explained by the possibility of loading more than one gp45 onto the 30/50mer p/t DNA); or (iv) a combination of effects at several of these steps. Similarly, the slow step that involves gp44/62 alone could be either a slow conformational change in gp44/62 that is required before the ternary complex can be assembled and ATP hydrolysis achieved, or a conformational change in the gp44/62 complex after hydrolysis that occurs after the sliding clamp has been loaded onto the p/t DNA and gp44/62 dissociates from both DNA and gp45 (a conformational change of the latter type could also be related to the slow release of the reaction product, ADP).{

876

Kinetics of Loading the T4 Processivity Clamp

A slow conformational change in the gp44/62-gp45 complex limits the stimulation of ATP hydrolysis by p/t DNA Three-syringe rapid-quench experiments The steady-state ATPase titrations described above have shown that at least two partially ratelimiting steps occur in the loading of the gp45 clamp onto p/t DNA. One of these steps involves a binary complex (either gp45-gp44/62 or gp45-p/t DNA), and the other involves gp44/62 alone. In addition, a closer inspection of a previous rapidquench experiment (Figure 1, ®lled circles) has suggested that one of these steps must occur before, and the other after ATP hydrolysis. This follows because a small lag and burst phase can be recognized in the kinetic pro®le when the three components (gp44/62, gp45 and DNA) are mixed together (Figure 1), which is consistent with the presence of two rate-limiting steps of comparable rate that occur before and after hydrolysis. To determine which complex is involved in the slow step that occurs before hydrolysis, three-syringe rapid-quench experiments were designed in which the gp44/62 complex was preincubated with ATP and either gp45 or p/t DNA for a time suf®cient to reach the steady-state ATPase rate (®ve seconds). After this interval, the third component (either p/t DNA or gp45, respectively) was added from a third syringe and the reaction was continued for varying times before stopping the reaction with an acid quench from a fourth syringe (the experimental design is shown in Figure 6). If the slow step before hydrolysis involves only gp44/62, then this step would be ``overcome'' during the ®ve second preincubation period (regardless of whether the gp44/62 component had been pre-mixed with gp45 or with p/t DNA), and addition of the third component would result in a fast burst of ATP hydrolysis followed by a slower steady-state ATP hydrolysis rate determined by the two partially rate-limiting steps that must occur before each subsequent ATP hydrolysis cycle. Conversely, if the slow step before hydrolysis involves the gp44/62-gp45 complex, then a fast burst of ATP hydrolysis would be seen only if gp44/62 had been preincubated with gp45, and then p/t DNA is added, and not if gp44/62 had been preincubated with p/t DNA before the addition of gp45. (In the latter case, the kinetic pro®le would be very similar to that seen in the ``normal'' two-syringe rapid-quench experiment (Figure 1) in which all the components are mixed with ATP at the same time.) { We have observed, in the reaction involving the 1.5 seconds preincubation time, that less than one-half of an equivalent of ATP (relative to the amount of gp44/62 present) was hydrolyzed (data not shown). This con®rms that ATP hydrolysis is not required for the subsequent stimulation of the steady-state ATPase rate by p/t DNA.

Figure 6. A slow step involving the binary gp45gp44/62 complex limits the stimulation of ATP hydrolysis by p/t DNA. Presteady-state, three-syringe experiments were performed as described in Experimental Procedures. gp44/62 (2 mM) was premixed, in the presence of ATP, with either 3 mM gp45 (®lled circles) or 2.4 mM p/t DNA (open circles), before the addition of the third component, either 2.4 mM p/t DNA or 3 mM gp45, respectively. (The concentrations listed are ®nal concentrations after adding the third component.) A burst of ATP hydrolysis is observed only when gp44/62 is pre-incubated with gp45 (®lled circles) and not when gp44/62 is pre-incubated with p/t DNA (open circles). Therefore, a slow conformational change in the gp45gp44/62 binary complex must limit the rate of the hydrolysis step. The burst amplitude (1.9 mM) corresponds to only one ATP molecule hydrolyzed per gp44/62 complex in a single turnover. The measured steady-state rate is 5.5 mM sÿ1 per mM gp44/62.

Figure 6 shows the outcome of such experiments. The results have been corrected for the amount of ATP that is hydrolyzed by gp44/62 (stimulated either by the gp45 or the p/t DNA cofactor) during the preincubation time. This amount was determined by adding acid quench reagent instead of the third component from the third syringe, and was subtracted from the ADP concentration measured at the time-points shown. As seen in Figure 6, an initial fast burst of ATP hydrolysis was observed in this reaction only when gp44/62 is premixed with gp45, and then rapidly mixed with p/t DNA (Figure 6, ®lled circles). The burst phase is followed by a steady-state rate of ATP hydrolysis of 5.5 mM sÿ1 per mM gp44/ 62, which is similar to that obtained in previous rapid-quench and pulse-chase experiments (Figure 1, ®lled circles and Figure 2). Changing the preincubation time from ®ve seconds to 1.5 or ten seconds, does not change the burst size.{ Conversely, a fast burst phase of ATP hydrolysis is not

877

Kinetics of Loading the T4 Processivity Clamp

observed when the order of addition of the two cofactors is reversed, so that gp45 is added to gp44/62 that has been pre-incubated with ATP and p/t DNA (Figure 6, open circles). Rather, the kinetic pro®le obtained is similar to that observed previously when all three components are mixed with ATP at the same time (Figure 1, ®lled circles). Therefore, these three-syringe experiments prove that the slow step before hydrolysis can be overcome only if gp44/62 is preincubated with gp45 (and not with p/t DNA), indicating that the slow step before hydrolysis involves the binary gp44/ 62-gp45 complex. The burst amplitude obtained re¯ects the number of ATP molecules hydrolyzed in a single reaction cycle when p/t DNA is added to an ``activated'' gp44/62-gp45 complex. This value, as expected, was found to be dependent on the concentration of gp44/62 used in the experiment (in the presence of excess gp45 and p/t DNA), and was 1.9 mM when 2 mM gp44/62 was used (Figure 6, ®lled circles) and 1.1 mM when 1 mM enzyme was used (data not shown). Therefore, in both experiments, the burst amplitude is very close to one ATP molecule hydrolyzed per gp44/62 complex present, con®rming our conclusions from the pulse-chase experiments described above (Figure 2). In light of these results, we ask again about the slow step that involves gp44/62 alone. If this also had occurred before hydrolysis (and the slow step after hydrolysis again involved gp45, as could occur if either dissociation of the gp44/62-gp45 complex or gp45 sliding off the p/t DNA were

also slow steps), then we would have expected to see a more pronounced burst phase in the experiment in which gp44/62 is pre-incubated with ATP and p/t DNA, followed by gp45 addition (Figure 6, open circles), compared to the experiment in which all three components are mixed with ATP at the same time (Figure 1, ®lled circles). Since the kinetic pro®les of both experiments appear very similar, we conclude that the slow step involving gp44/62 alone must occur after ATP hydrolysis.

Discussion A model for the T4 processivity clamp-loading reaction cycle The results presented here and elsewhere now permit us to design and test a detailed reaction cycle for the loading of the T4 gp45 sliding clamp onto the template DNA at primer-template junctions. The overall mechanism is outlined in Figure 7. The following results underlie the proposed reaction cycle. We have shown here that a relatively slow (and rate-limiting) step that involves only gp44/62 and gp45 occurs before ATP hydrolysis in the clamp-loading reaction (Figures 4 and 6). A variety of studies have suggested that this step corresponds to a conformational change in the gp45-gp44/62 complex. After this step has occurred, p/t DNA can bind to this ``activated'' binary complex and stimulate ATP hydrolysis (Figures 4 and 6). Previous studies have shown

Figure 7. Proposed clamp-loading mechanism for the T4 system. The gp45 ring is shown as partially open in solution.23 A slow conformational change, which does not require DNA, must occur in the binary gp45-gp44/62 complex before hydrolysis can be stimulated by p/t DNA. This conformational change is related to the completion of the ring-opening process.22,26,29 Previous crosslinking and footprinting studies16,17 have shown that hydrolysis leads to dissociation of the clamp loader from the ternary complex, leaving the gp45 ring assembled around the p/t DNA. Further closure of the open gp45 subunit interface has been observed upon interaction with the polymerase gp43 via binding of the C terminus of gp43 at the subunit interface.22 We have found that only one ATP molecule is hydrolyzed per clamp-loading cycle, and that this hydrolysis reaction and the various associated protein and DNA dissociation processes are relatively fast. A second slow step, which is probably related to ADP release, occurs after gp44/62 release. The clamp-loading reaction cycle presented here has many features in common with that recently proposed for the E. coli DNA replication system.31

878

Kinetics of Loading the T4 Processivity Clamp

that ATP hydrolysis is required both for the loading of the gp45 clamp onto p/t DNA and for the release of the gp44/62 loading complex into solution.9,15 ± 17 Combining these results has led us to depict the ATP hydrolysis step in Figure 7 as preceding gp45 loading and gp44/62 dissociation from p/t DNA. Finally, our results have shown that a second partially rate-limiting step occurs after ATP hydrolysis. This step involves only the gp44/62 complex (Figures 4 and 6) and we suggest that this step is related to ADP dissociation from the recycled clamp loader. Fitting of parameters and kinetic simulation of the proposed reaction-cycle model In order to test the validity of this proposed minimal reaction cycle, we have performed a simultaneous ®tting (to the equations shown in the Appendix) of the data obtained from the titrations of the steady-state ATPase activity of gp44/62 with cofactors (Figure 4). The equations were formulated on the basis of the model presented in Figure 7, plus two kinetic assumptions. These assumptions are: (i) that the dissociation rates of the p/t DNA and the gp45 clamp from the gp44/ 62 loading complex are both fast relative to the ATP hydrolysis step; and (ii) that the reaction involving the sliding of the closed gp45 clamp off the unblocked p/t DNA used in this study is fast relative to the rate of the pre-hydrolysis conformational change in the binary (gp45-gp44/62) loading complex. The simultaneous ®ts obtained are shown as the continuous curves in Figure 4 and provide best-®t values for the ®ve rate constants that describe the steps of the mechanism of Figure 7. These rate constants are: (i) k45bind, the bimolecular rate constant for gp45 binding to gp44/62; (ii) kb.c., the ®rst-order rate constant for the prehydrolysis conformational change in the binary gp44/62-gp45 complex; (iii) kDNAbind, the bimolecular rate constant for p/t DNA binding to the activated gp44/62-gp45 complex; (iv)kt.c., the ®rst-order rate constant for the hydrolysis step within the ternary complex; and (v)k44c., the ®rstÿ1

{ Note that this rate constant (17(3) s ), which is the lower limit value for the rate constants for the steps in which gp45 is involved in either a binary or the ternary complex, is approximately equal to the rate constant (18(3) sÿ1) for the pre-hydrolysis conformational change that was calculated assuming that the protein dissociation and the sliding off of gp45 from p/t DNA reactions are both fast. This follows because the rate constant for the hydrolysis of ATP by the ternary complex (270(70) sÿ1) is much larger than that measured for the pre-hydrolysis conformational change (18(3) sÿ1) and, therefore, the lower limit for the rate constant for the steps involving gp45 (which is a combination of the two rates constants referred to above) turns out to be very close to that measured for the latter (slower) step.

order rate constant for the ®nal partially ratelimiting step that involves gp44/62 alone, which we have tentatively attributed to ADP release. (Here, the subscript abbreviation b.c. represents binary complex, t.c. represents ternary complex, and 44c. represents gp44/62 complex.) All the rate constants obtained using these ®tting routines are ``apparent'', meaning that they represent minimal values for the rate constants of the respective forward reactions. The apparent rate constant values obtained by ®tting the model are: kb.c. ˆ 18(3) sÿ1; kt.c. ˆ k44c. ˆ 11.6(1) sÿ1; k45bind ˆ 31 270(70) sÿ1; ÿ1 ÿ1 (5) mM s ; and kDNAbind ˆ 190(17) mMÿ1 sÿ1. We show in Figure 8(b) that the simulated data points (open symbols) that have been ®tted to the above apparent rate constants and the minimal reaction scheme (shown in Figure 7 and summarized in Figure 8(a)) reproduce very well the experimental steady-state ATPase reaction rate data that we obtained above (®lled symbols in Figure 8(b)), thus also justifying the approximations involved in using a constant concentration of free non-titrating cofactor in deriving the equations shown in the Appendix. In addition, as indicated above, in creating the ®tting equations in the Appendix we assumed that the dissociation rates of gp45 and p/t DNA from gp44/62 are fast relative to the rate of ATP hydrolysis, and that the sliding of gp45 off the p/t DNA is fast relative to the pre-hydrolysis conformational change in the binary loading complex. If these assumptions are incorrect, the derived rate constants serve, at least, to set lower limits for the rate constants of the steps involved. Rate constant kt.c. is associated with the steps in which p/t DNA is involved as a cofactor. Therefore the ®tted value of this rate constant (270(70) sÿ1) represents a lower limit for both the rate of ATP hydrolysis and the rate of p/t DNA dissociation from the gp44/62 clamp-loading complex. The rate constant that describes the steps in which gp45 is involved (kE
45) is given by: 1 kE
45

ˆ

1 kb:c:

‡

1 kt:c:

As a consequence, the calculated value of this parameter (17(3) sÿ1) represents the lower limit for the rate constant of the pre-hydrolysis conformational change, as well as for the rate constants for gp45 dissociation from gp44/62 and for the sliding off of gp45 from p/t DNA.{ Finally, the rate constant that describes the steps that involve gp44/62 alone (k44cˆ11.6(1) sÿ1) represents an upper limit for the rate constant of the post-hydrolysis conformational change in the gp44/62 complex, which we have attributed to ADP release. We have used the rate constants obtained by ®tting the steady-state ATPase titrations to simulate independently the results of the rapid-quench experiment (Figure 1, closed circles). The simu-

879

Kinetics of Loading the T4 Processivity Clamp

Figure 8. Simulations of the steady-state ATPase titrations and the rapid-quench experiments based on the model presented in Figure 7. (a) The overall kinetic scheme, and the relative rate constants utilized in the simulations shown in (b) and (c). (b) Data from the ATPase titrations (Figure 4) were simulated to show that the approximations made in deriving the equations in the Appendix do not appreciably affect the ®tting results. In fact, simulations performed using the ®tting parameters (open symbols) closely resemble the experimental data (®lled symbols). (c) Simulation (continuous curve) of the rapid-quench experiment shown in Figure 1 (filled circles), using the rate constants derived from the steady-state ATPase data (see the text).

lation is shown in Figure 8(c). The simulated curve ®ts very closely to the experimental data points. In particular, the small lag and burst phases, and the subsequent steady-state rate are ®t extremely well by the simulated curve. This suggests strongly that the relative rate assumptions (listed above) that we made in creating the equations for ®tting the steady-state ATPase titrations are very likely to be correct. In fact, if the protein and p/t DNA dissociation rates were not fast relative to the hydrolysis step, then the burst phase would have been more pronounced than that seen experimentally. Similarly, if the rate of sliding of gp45 off the p/t DNA was not fast relative to the pre-hydrolysis conformational change, the burst phase would have been more pronounced, and the observed lag

phase would have been smaller, because in that scenario the pre-hydrolysis conformational change that is responsible for the appearance of the lag phase would have been faster. The kinetics of the T4 clamp-loading reaction cycle On the basis of the above considerations, we can comment further on the clamp-loading reaction scheme shown in Figure 7. First, we note that the presteady-state kinetic experiments of Figure 1 have shown that the ternary complex must be assembled before ATP hydrolysis can take place. The fact that a burst of ATP hydrolysis is not observed when gp45 and gp44/62 are mixed with

880 ATP in the absence of p/t DNA means that the rate-limiting step is either the chemical hydrolysis step itself, or a step that precedes hydrolysis. Stimulation of the ATPase activity by p/t DNA must re¯ect the acceleration of this step, and therefore p/t DNA must enter the gp44/62-gp45 complex before ATP hydrolysis takes place. This is con®rmed by the fact that a burst phase is not observed in the ATPase rate when gp44/62 and p/t DNA are mixed in the absence of gp45 (data not shown). In addition, the kinetic pro®le of the rapid-quench experiment remains linear even when both cofactors (gp45 and p/t DNA) are present together, showing again that either the chemical hydrolysis step or a step that precedes it must be at least partially rate-limiting. In addition, the steady-state titrations (Figure 4) have shown that p/t DNA is not part of the rate-limiting step, since it acts catalytically in stimulating the ATPase activity of gp44/62 in the presence of excess gp45. This means that the action of p/t DNA in the reaction cycle must be fast, and therefore not rate-determing. In support of this conclusion, a catalytic effect of p/t DNA on the gp44/62-gp45 complex was observed in our ¯uorescence study.18 On the other hand, the titration with gp45 has revealed the existence of at least two slow steps with similar rates, one that involves gp45 and another that involves the gp44/62 complex alone. This follows because gp45 does not act completely stoichiometrically in the presence of p/t DNA, unlike its behavior in the absence of DNA (Figure 5). Thus, a stoichiometric pro®le would be observed if the titrating cofactor becomes part of the slowest step in the reaction. However, in contrast, we ®nd (in the presence of p/t DNA) that approximately one-half of an equivalent of gp45 (relative to the amount of gp44/62 present) suf®ces to stimulate the ATPase activity of the clamp loader to its maximum level. This means that one molecule of gp45 is able to stimulate approximately two molecules of the gp44/62 loading complex before the ®rst gp44/62 complex has completed its loading cycle, requiring that there must be a slow step involving the gp44/62 complex alone that occurs at a rate similar to that of the reaction in which the gp45 clamp plays a part. The three-syringe experiments then served to demonstrate that a slow step involving gp44/62 and gp45 does exist, and that this step occurs before ATP hydrolysis. In fact, preincubation of gp44/62 with gp45 in the presence of ATP before the addition of p/t DNA results in a fast burst phase of ATP hydrolysis (Figure 6, ®lled circles). This means that the step that limits the hydrolysis reaction must be a conformational change in the binary gp45-gp44/62 complex. Once this conformational change has occurred, DNA binding and hydrolysis follow rapidly, in agreement with the catalytic action of DNA observed in the steadystate kinetic experiments (Figure 4, open circles)

Kinetics of Loading the T4 Processivity Clamp

and in our previous ¯uorescence studies.18 The slow step that is observed in the steady-state ATPase reaction and involves only the gp44/62 complex must occur after hydrolysis, since preincubation of gp44/62 with DNA and ATP before gp45 addition does not change the kinetic pro®le relative to that obtained in an experiment in which all three components were mixed together simultaneously with ATP (compare Figure 6 (open circles) and Figure 1 (filled circles)). The results presented here are consistent with a model in which gp45 binds to gp44/62 and, in the presence of ATP, undergoes a slow conformational change. We relate the pre-hydrolysis conformational change in the binary gp45-gp44/62 complex to ring opening, by analogy with the E. coli system (see below). In addition, although the gp45 ring has been shown to be partially open in solution,23 it has been observed to open further upon gp44/62 binding in the presence of ATP.22 Once this conformational change has occurred, p/t DNA binds rapidly to the activated gp44/62-gp45 complex and stimulates hydrolysis, which also occurs rapidly. Following this ternary complex formation, gp44/62 must dissociate from both p/t DNA and gp45, leaving the gp45 ring assembled around the DNA. We cannot tell from our present data whether hydrolysis precedes or follows the protein and DNA dissociation steps. However, earlier work16,17 has shown that hydrolysis leads to dissociation of the ternary complex. Therefore, we have depicted the hydrolysis step (Figure 7) as preceding the protein and DNA dissociation processes. After the clamp-loading event is complete, a second slow step occurs when the gp44/62 is free in solution. We relate this slow step, which involves gp44/62 alone, to the process of ADP release. This conclusion is based both on analogy with the E. coli system (see below), and on the fact that product inhibition studies using aluminum tetra¯uoride have shown that ADP release from gp44/62 is at least partially rate-limiting in the T4 system.9 The steady-state titration experiments (see Appendix) were ®tted to two equations that are de®ned by our model, together with the two relative rate assumptions described above. A ``global'' ®tting process then provided the rate constants associated with the steps involved, with gp45 binding to gp44/62 with a bimolecular rate constant of 31(5) mMÿ1 sÿ1, which is consistent with the value (31(16) mMÿ1 sÿ1) of this parameter measured by others.21 The pre-hydrolysis conformational change in the gp45-gp44/62 complex occurs at a rate of 18(3) sÿ1. DNA binds to the activated gp44/62-gp45 complex with a bimolecular rate constant of 190(17) mMÿ1 sÿ1, and hydrolysis of one ATP molecule follows at a rate of 270(70) sÿ1. Finally, the slow step (presumably ADP release), which involves the gp44/62 complex once it has dissociated from the p/t DNA and from the gp45 clamp, occurs at a rate of 11.6(1) sÿ1.

Kinetics of Loading the T4 Processivity Clamp

Simulation of the rapid-quench experiment using the above rate constants ®ts closely to the experimental data (Figure 8(c)). In particular, the small lag and burst phases observed experimentally (Figure 1) are in agreement with the existence of the two slow steps that precede and follow hydrolysis with rate constants of 18(3) sÿ1 and 11.6(1) sÿ1, respectively. This signi®es that the assumptions that were made in ®tting the steady-state data are most likely correct. For example, if the rates of gp45-gp44/62 dissociation and the gp45 sliding off of the p/t DNA were not signi®cantly faster than the above rates, then the pre-hydrolysis conformational change would occur at a faster rate, and the post-hydrolysis slow step would occur at a slower rate, leading experimentally (in Figure 1) to both a smaller lag phase and a more pronounced burst phase of ATP hydrolysis relative to the simulated data of Figure 8(c). Our kinetic data have demonstrated that ATP hydrolysis does not occur until after a ternary complex has been formed. This says that hydrolysis is not needed to activate the gp44/62-gp45 complex prior to p/t DNA binding, contrary to what we and others had suggested previously on the basis of ¯uorescence studies.18,19 In those earlier studies it was shown that ATP, but not ATPgS, was able to induce a conformational change in the gp45gp44/62 complex, and it was this observation that led to the conclusion that ATP hydrolysis was required to produce the conformational change. The present data have shown that the hydrolysis step is most likely rate-limiting in the absence of p/t DNA, and that the DNA component acts to accelerate this step to the point that a conformational change in the gp44/62-gp45 complex preceding hydrolysis becomes partially rate-limiting. Thus, in the absence of p/t DNA, the predominant gp45-gp44/62 species present in solution must be the one that has experienced the pre-hydrolysis conformational change, but has not yet hydrolyzed ATP. The result of all this is that we now believe that the ATP-induced change observed in the spectra of the ¯uorescently labeled gp45 mutants as a consequence of the action of the gp44/62 complex re¯ects a conformational change in the gp44/62gp45 complex that occurs on ATP binding, and thus precedes ATP hydrolysis. This is con®rmed by the observation in our previous crosslinking experiments that a conformational change is indeed induced by ATP binding, since this conformational change could be induced by the binding of ATPgS.20 It may be that this change was not observed in the ¯uorescence studies because ATP and ATPgS bind to the complex somewhat differently. We note, in this context, that a subsequent ¯uorescence study21 showed that the ATP-induced ¯uorescence change in the gp45-gp44/62 complex occurs with a rate constant of 14(1.4) sÿ1, which is similar to the rate constant of 18(3) sÿ1 that we

881 measured for the pre-hydrolysis conformational change reported here. Presteady-state kinetic measurements of the gp44/62 ATPase activity have been performed previously in our laboratory34 and in the Benkovic laboratory.33 We have found that the fast burst phase for ATP hydrolysis catalyzed by gp44/62 in the presence of both gp45 and p/t DNA that we reported previously as corresponding to the hydrolysis of four ATP molecules per reaction cycle34 was actually an artifact produced by the quenching agent that was used in the experiment. Berdis & Benkovic33 reported that a burst of ATP hydrolysis is observed when the gp44/62 loads gp45 onto a p/t DNA that is ``blocked'' at both ends, thus preventing gp45 from sliding off over the ends. This result is not inconsistent with our present results and model, since gp45 dissociation from the p/t DNA becomes the slowest step in the reaction cycle when a blocked p/t DNA is used, and thus the observation of a presteady-state burst of ATP hydrolysis would be expected under these conditions. However, it was reported in the Berdis & Benkovic study33 that 1(0.5) mM ATP was hydrolyzed by 250 nM gp44/62 complex per loading event, which would correspond to the hydrolysis of four ATP molecules per gp44/62 complex in each clamp-loading event, in contrast to the one ATP molecule hydrolyzed per reaction cycle that we have demonstrated in the present study. It may be that the low concentration of proteins and DNA used in the Berdis & Benkovic study33 (250 nM gp44/62 compared to the 2 mM concentration used in the present study), together with the high concentration of ATP that they used (1 mM) might have made this measurement more prone to error and have led to the result obtained. Finally, we note that Sexton et al.21 have reported a burst phase in the ATPase kinetics of the gp44/62 complex in the presence of gp45 and in the absence of DNA. We cannot reconcile this observation with our ®nding of no burst phase under these conditions. The T4 clamp-loading reaction cycle has remarkable similarities to that observed with the E. coli system To this point, the clamp-loading mechanism that is a central component of the DNA replication process has been characterized most extensively for the E. coli system. Here, the clamp loader is the g-complex, which is composed of ®ve different subunits. One of these subunits (d), which has no ATPase activity, is able to bring about the opening of the dimeric sliding clamp (b) of E. coli by itself, meaning that in this instance a simple protein-protein interaction can drive clamp opening. However, when the d subunit is integrated into the reconstituted g-com-

882 plex, the b ring cannot be opened unless ATP (or ATPgS) is added to the solution. To explain this, it has been proposed that the d subunit is buried within the g-complex, and that ATP binding (not hydrolysis) is suf®cient to trigger a conformational change in the g-complex that exposes the d subunit for interaction with the b ring.26,29 This pre-hydrolysis conformational change in the clamp-clamp loader complex occurs with a ®rst-order rate constant of 12 to 14 sÿ1, and is the rate-limiting step of the E. coli clamp-loading cycle.27,28 The ATP-bound open clamp-clamp loader complex of E. coli binds to p/t DNA with high af®nity,28,29 and with a second-order rate constant of 2  108 Mÿ1 sÿ1.31 Hingorani et al.30 have used measured burst phase amplitudes to propose that two molecules of ATP are hydrolyzed sequentially as a consequence of p/t DNA binding, and that the rate-limiting step of the reaction cycle occurs between the two hydrolysis steps. In a subsequent report31 (involving some of the same authors) it was shown that the hydrolysis of the two molecules of ATP that occurs as a consequence of p/t DNA binding is simultaneous and is characterized by a lower-limit reaction rate constant of 34 sÿ1. The complex then dissociates with a ®rst-order rate constant of 22 sÿ1, leaving the b clamp assembled around the p/t DNA.26,28 In this scheme, the ratelimiting step of the ATP hydrolysis cycle occurs in solution after the clamp loader has dissociated from the p/t DNA, and probably re¯ects a slow rate of ADP release.31 Our present model for the T4 clamp-loading system resembles, in many ways, this latter E. coli clamp-loading mechanism, although there are important differences (see below). Thus, we observe a slow step prior to hydrolysis that involves only the clamp-clamp loader complex, and which occurs with a ®rst-order rate constant of 18 sÿ1. This value is close to the rate constant of 12-14 sÿ1 measured in the E. coli system for the comparable reaction.27,28 The binding of p/t DNA to this activated complex in the T4 system is fast and occurs with a bimolecular rate constant of 2  108 Mÿ1 sÿ1, which is identical with the rate constant measured for this process with the E. coli system.31 The ATP hydrolysis, clamp loader release, and p/t DNA dissociation steps are fast in both systems, and the slowest step in the ATP hydrolysis cycle of the E. coli system is one that involves the clamp loader complex alone. This step occurs in solution after the loading event, and (as in T4) probably re¯ects ADP release from the dissociated clamp loader complex.31 It has been proposed for the E. coli system that this slow rate of release of ADP from the clamp loader complex may have the bene®cial effect of transiently inactivating the clamp loader, thus preventing it from competing with the polymerase for the loaded b-clamp.35 The same argument could be applied to the apparent

Kinetics of Loading the T4 Processivity Clamp

slow rate of ADP release from the gp44/62 clamp loader in the T4 reaction cycle. Only one ATP molecule is hydrolyzed by the T4 gp44/62 complex in the clamploading process The pulse-chase experiments, in which the clamp loader complex was ®rst mixed with [a-32P]ATP and then chased with excess non-radiolabeled ATP before quenching, showed that only one ATP molecule per gp44/62 complex is hydrolyzed during the chase period in a single turnover of the reaction cycle (Figure 2). A pulse-chase experiment can detect only ATP molecules that are bound suf®ciently tightly to the enzyme to be hydrolyzed without exchange with the ``cold'' ATP added in the chase phase. Thus, if the rate of dissociation of the ATP substrate from the active site of the enzyme is faster than the hydrolysis rate, then bound [a-32P]ATP will dissociate during the chase and be diluted out by the excess nonradiolabeled ATP. Therefore, in principle, it is possible that more than one ATP molecule might be hydrolyzed, but clearly only one is suf®ciently tightly bound to the enzyme to yield radioactive ADP in an experiment of this kind. However, the three-syringe experiments argue against the possibility that other, more weakly bound, ATP molecules are hydrolyzed during the clamp-loading reaction cycle, since we have shown that a fast burst of hydrolysis is observed when the [a-32P]ATP-bound gp44/62-gp45 activated complex is mixed rapidly with p/t DNA in the absence of any cold ATP (Figure 6, ®lled circles), and that the burst amplitude here also corresponds to only one ATP molecule hydrolyzed per gp44/62 complex. In this experiment, any additional loosely bound [a-32P]ATP would have had the opportunity to rebind to the active sites of the gp44 subunits, and then to be hydrolyzed. In addition ATP hydrolysis has been shown to be irreversible,9,34 and therefore the measured burst phase of one ATP molecule is not the result of an equilibrium between forward and backward reactions at different sites. These experiments force us to conclude that only one ATP molecule is indeed hydrolyzed per gp44/62 complex during the loading of a gp45 sliding clamp onto p/t DNA. The situation differs in E. coli, where at least two ATP molecules are bound to the g subunits of the g-clamp loader complex29,36 and two ATP molecules are hydrolyzed per clamp-loading reaction cycle. In contrast, the gp44/62 clamp loader of T4 contains four gp44 subunits, each of which carries an ATP-binding site.10 Detailed binding studies (P.P. & P.H.v.H., unpublished results) of the T4 system have shown that more than one ATP molecule binds to these four subunits at various stages of the clamp-loading process. Yet, in the present study (Figures 2 and 6), only one ATP molecule appears to be hydro-

Kinetics of Loading the T4 Processivity Clamp

lyzed in the T4 clamp-loading cycle{ What role do the other ATPs (and ATP-binding sites) play in the T4 clamp-loading process, and why are they not hydrolyzed? This question will be dealt with in detail elsewhere. A general role for nucleotide triphosphate (NTP) binding and hydrolysis in biological systems These studies of the clamp-loading reaction cycles of DNA replication can be put into a more general context. Extensive studies of different biological systems in which a speci®c molecular function is driven by the repetition of an NTPdependent binding and hydrolysis cycle have led to the appreciation that a common mechanism may be used to ``couple'' NTP binding and hydrolysis to such cyclic reaction processes (for a review, see Vale & Milligan).37 Such systems include the motor proteins involved in intracellular transport,37 ± 40 the G proteins involved in intraand intercellular signalling,39,41 and the chaperone protein unfolding and refolding complexes (reviewed by Fink and by Saibil).42,43 In reaction cycles of this sort, the macromolecular component(s) involved must interact with their ``target(s)``, perform their function, dissociate and return to the starting conformation to permit another reaction cycle to begin. Free energy input is needed to reset the system; i.e. to return the system to the initial higher free energy level from which the next cycle can start. This general perspective has recently been applied to the reaction cycle of nucleic acid helicases.44 { A reviewer has suggested that the possibility remains open that a second ATP molecule could, in principle, be hydrolyzed in each reaction cycle if the rate of hydrolysis of this additional ATP were slow, since this ATP would then ``disappear'' into the steadystate rate. Additional simulations showed that the hydrolysis of one (but not more) additional ATP molecule of this type might be possible within the error of our experiments, and so we tested for this possibility by means of an additional pulse-chase experiment. The design of this experiment was similar to that shown in Figure 6, with the difference that excess unlabeled ATP was added with the fourth syringe instead of the acid quench solution, and the reaction was permitted to run for an additional period before quenching (approximately ten turnovers) that should be suf®cient to convert any tightly bound ATP to product. The expected burst size for this experiment would be two ATP molecules if a second ATP molecule were (as proposed by the reviewer) hydrolyzed slowly by the gp44/62 complex. However, only one ATP molecule was observed in the burst in this experiment as well, showing that if a second ATP molecule is indeed hydrolyzed in the reaction it must both be hydrolyzed slowly and bound very weakly. This issue is being investigated further (P.P. & P.H.v.H., unpublished results), but at this point we continue to conclude that clamp loading in the T4 system is driven by the hydrolysis of only one ATP molecule per reaction cycle.

883 It has become clear that the NTP hydrolysis process itself need not be correlated directly with the conformational change in the protein component(s) that is (are) responsible for the speci®c function. In fact, NTP-bound states are often associated with important conformational changes that occur during the reaction cycle. In many of the systems studied, it has been found that the free energy of NTP binding is suf®cient to induce a major conformational change in the protein complex involved, and that NTP hydrolysis then represents a mechanism for ``relaxing'' this conformation back to its starting state. Therefore, the major structural changes that occur upon NTP binding and upon NTP hydrolysis product release are often opposite in direction, with one of these two opposite conformational changes being utilized to drive the speci®c reaction cycle, while the other is used to reset the system. The NTP hydrolysis step can either precede or follow the central change that corresponds to the biological function that is performed by the system. For example, in the well-known myosin-actin motor protein system, ATP hydrolysis precedes the sliding of the actin ®lament. In fact, the ``power stroke'' in this system results from the conformational change that occurs upon binding of the ADP-bound myosin to the actin ®lament and is related to product release. ATP binding is needed to induce an opposite conformational change that dissociates the complex, with ATP hydrolysis occuring subsequently in solution so that the cycle can restart. In contrast, in the kinesin-tubulin motor system, ATP hydrolysis follows the forward movement of the kinesin on the tubulin ®ber. In this case, the equivalent power stroke occurs upon ATP binding, with ATP hydrolysis resulting in dissociation of the complex. Similarly, the G proteins are ``switched on'' by a conformational change that occurs upon ATP binding. Thus G proteins in their ATP-bound state bind to the target protein and activate the signal. ATP hydrolysis follows and induces dissociation, switching off the signal. Chaperonin protein-unfolding systems work in a manner similar to that of actin-myosin, in that ATP binding leads to dissociation of hydrophobic residues within the chaperonin cavity wall from the target protein, and hydrolysis leads to rebinding. Thus, in general, NTP binding and hydrolysis can be viewed as representing a ligand-driven switch between two conformations of a macromolecular machine, with one being responsible for the binding of a protein component (or components) to the target to ful®l the speci®c function of the reaction cycle, while the other is utilized to dissociate the complex and reset the system. (For further discussion and application of these concepts, see von Hippel & Delagoutte.)44 The clamp-loading machinery of DNA replication seems to function in a similar way. Initially, it was thought that ATP hydrolysis was required to overcome the energetically unfavorable process of opening the closed ring-like sliding clamp, and it

884 was surprising to ®nd (initially with the E. coli system) that ATP binding was suf®cient to induce the conformational change required to drive ring opening.26,29 In addition, in keeping with the generalities presented above, it was found that the g-complex of the E. coli system undergoes reversible conformational changes during the ATP hydrolysis cycle that are associated, respectively, with an ATP-bound and an ADP-bound state of the system.35 One of these binding states (here the ATP-bound form), switches on the clamp loader, driving it to the p/t DNA target. Binding to the DNA target then triggers ATP hydrolysis, which relaxes the clamp-loading complex back to its lowaf®nity (ADP-bound) DNA-binding state.

Kinetics of Loading the T4 Processivity Clamp

Experimental Procedures Materials DNA oligonucleotides were purchased from Genosys (The Woodlands, TX) and gel-puri®ed. ATP was purchased from Amersham Pharmacia (Piscataway, NY) and [a-32P]ATP was from DuPont New England Nuclear (Boston, MA). All other biochemicals and chemicals were obtained from Sigma Chemical Co. (St. Louis, MO), Aldrich Chemical Co. (Milwaukee, WI), or BoehringerMannheim (Indianapolis, IN). Primer/template DNA The p/t DNA used in these experiments was:

.

The ®ndings that we have presented here suggest that the T4 clamp-loading system of bacteriophage T4 works in a similar way. The gp44/62 clamp loader associates with its cofactor targets (the gp45 sliding clamp and the p/t DNA) before ATP hydrolysis takes place. In addition, a pre-hydrolysis slow step involving the gp44/62gp45 complex has been observed, suggesting that this binary complex undergoes a conformational change upon ATP binding, which may switch on or activate the protein complex for p/t DNA binding. By analogy with the E. coli system, this conformational change may be associated with ring opening. Although it has been suggested that the T4 ring may be partially open even when free in solution,23 it has been observed that a further opening of the ring occurs on binding of the clamp to the gp44/62 clamp loader complex, and this further opening seems to be required for ef®cient loading of the sliding clamp.22 Earlier studies have shown that ATP hydrolysis leads to dissociation of the loading complex.16,17 Therefore, here also, in common with the varying systems described above, ATP hydrolysis may serve to relax the ATPinduced conformational change back to the starting state of the reaction cycle, leading both to dissociation of the clamp loader and to reclosing of the ring around the p/t DNA. It is clear that the ATP-driven clamp-loading systems of the DNA replication complexes of a variety of organisms utilize similar regulatory principles, and future work will show how these regulatory mechanisms are functionally incorporated into the reaction cycles of the DNA replication complexes as they interact with the components and events of transcription, recombination, and repair.

and was assembled by annealing equimolar amounts of the two single-stranded component DNA oligonucleotides. Oligonucleotide concentrations were determined from measurement of UV absorbance at 260 nm, using calculated extinction coef®cients of 460.3 mMÿ1 cmÿ1 for the 50-mer and 304.3 mMÿ1 cmÿ1 for the 30-mer. Annealing was performed by raising the temperature of the oligonucleotide mixture to 90  C, and then slow cooling to room temperature over two hours. The completeness of the hybridization process involved in forming the p/t DNA construct was con®rmed by native gel electrophoresis. Proteins The T4 clamp and clamp-loader proteins (gp45 and gp44/62) were puri®ed as described.34 Protein concentrations were determined by measurement of UV absorbance at 280 nm, using the following calculated molar extinction coef®cients: gp44/62, e280 nm ˆ 1.2  105 Mÿ1 cmÿ1; and gp45, e280 nm ˆ 5.7  104 Mÿ1 cmÿ1.45 Presteady-state kinetic experiments Presteady-state kinetic measurements were made using an Update Instruments (Madison, WI) Quench Flow apparatus modi®ed to the design of Johnson;46 see Young et al.34 for a description of the apparatus. A 17 ml solution containing gp44/62 in a buffer composed of 25 mM Hepes (pH 7.5), 160 mM potassium acetate, 6 mM magnesium acetate, and 0.5 mM DTT, with or without gp45, was rapidly mixed with an equal volume of solution in the same buffer containing ATP, as well as a trace amount of [a-32P]ATP and p/t DNA, when present. The ®nal reaction concentrations (after mixing) were 1 or 2 mM gp44/62, 1.5 or 3 mM gp45 (when present), and 1.2 or 2.4 mM p/t DNA (when present). The concentration of ATP in the reaction solution ranged from 60 to 500 mM. [a-32P]ATP was used in these studies instead of [g-32P]ATP, since the (a-32P)-containing preparations contained a lower background (<0.5 %) of contaminating radiolabeled components than did the (g-32P)containing material. Reactions were done at room temperature (23  C).

885

Kinetics of Loading the T4 Processivity Clamp

The reaction was allowed to proceed as indicated by the time-points before mixing with a quench solution of 0.7 M formic acid dispensed from a third syringe. Although formic acid is not supposed to hydrolyze ATP, we noted that even spotting the quenched sample onto the TLC plate soon after collection showed that some slow hydrolysis does occur with time, and that this hydrolysis could become signi®cant by the time all the samples for a given run had been collected and the TLC plates had been run (one or two hours). Therefore samples (200 ml each), were collected into tubes containing 200 ml of chloroform, vortex mixed, and adjusted to pH 7 by adding 55 ml of 3.2 M Tris base. The products of the reaction were analyzed by spotting 1 ml of solution onto PEI-F cellulose TLC plates, which were developed in 0.35 M potassium phosphate buffer (pH 3). Radioactive ADP and ATP were quanti®ed using a Molecular Dynamics Storm model 860 PhosphoImager (Sunnyvale, CA). In pulse-chase experiments, a solution containing excess (15 mM) non-radiolabeled ATP and 15 mM magnesium acetate in reaction buffer was dispensed from the third syringe instead of the formic acid quench solution. After a chase time equivalent to at least ten turnovers of the ATP reaction (about ten seconds), the chase reactions were quenched by adding 15 ml of 7 M formic acid and 200 ml of chloroform, vortex mixed, adjusted to pH 7, and analyzed as described above. In the three-syringe experiments (see Results), the solutions of gp44/62 and gp45 or gp44/62 and p/t DNA were ®rst mixed with ATP. The reactions were then allowed to reach steady-state by incubating for 1.5 to ten seconds, and then mixed with the third component (either p/t DNA or gp45, respectively), which was dispensed from the third syringe. After varying additional incubation times, these reactions were quenched with 0.7 M formic acid from a fourth syringe, treated with chloroform (300 ml added to the 300 ml samples), neutralized, and analyzed as described above. Data were corrected for the amount of ATP hydrolyzed during the preincubation period. This amount was determined by adding acid instead of the third component from the third syringe, and the result was subtracted from the data points. Steady-state kinetic measurements Steady-state ATPase rates were measured spectrophotometrically in a Cary 3E spectrophotometer, using a coupled enzyme system as described.47 The assay monitors ATPase activity by following the decrease in absorbance at 340 nm as NADH is converted to NAD during the regeneration of the ADP product of the ATP hydrolysis reaction. The regeneration system contains 3 mM phospho(enol)pyruvate, 0.5-1.5 mM NADH, 0.025 unit mlÿ1 of pyruvate kinase, and 0.025 unit mlÿ1 of lactate dehydrogenase. Under these conditions, regeneration of ADP to ATP occurs at least 20 times faster than the measured rate of ATP hydrolysis, and therefore the monitoring system is never rate-limiting. The reactions used to measure the steady-state activation of gp44/62 ATPase by cofactors contained 1 mM ATP, 1 mM gp44/62, and 2 mM gp45 or 1.2 mM p/t DNA (whichever was present as the non-titrating cofactor), together with the indicated amounts of the titrating cofactor. The reaction buffer was the same as that used in the presteady-state experiments. Reaction volumes were 60 ml contained in a 0.3 cm path-length spectropho-

tometer cuvette. The assay temperature was maintained at 23  C using a temperature-controller. Data analysis The presteady-state experiments in which a burst phase was observed were ®tted to a single exponential followed by a linear phase, using non-linear regression analysis (KaleidaGraph; Synergy Software): y ˆ A…1 ÿ eÿlt † ‡ kt where A is the burst amplitude, l is the burst rate, k is the linear (steady-state) rate and t is the reaction time. The steady-state ATPase titrations were ®tted to the equations described in the Appendix. The simultaneous ®ttings involved were performed by using the ProFit program (QuantumSoft Software), and the kinetic simulations were performed using the KINSIM program.48

Acknowledgements This research was supported, in part, by USPHS Research Grants GM-15792 and GM-29158 (to P.H.v.H.), and by American Cancer Society Postdoctoral Fellowship PF-4303 (to G.J.L.). P.H.v.H. is an American Cancer Society Research Professor of Chemistry. We are indebted to Dr Manju Hingorani for helpful advice on the rapid-quench and pulse-chase experiments, and to Stephen Weitzel for help with the protein preparations.

References 1. Kong, X. P., Onrust, R., O'Donnell, M. & Kuriyan, J. (1992). Three-dimensional structure of the beta subunit of E. coli DNA polymerase III holoenzyme: a sliding DNA clamp. Cell, 69, 425-437. 2. Krishna, T. S., Kong, X. P., Gary, S., Burgers, P. M. & Kuriyan, J. (1994). Crystal structure of the eukaryotic DNA polymerase processivity factor PCNA. Cell, 79, 1233-1243. 3. Moare®, I., Jeruzalmi, D., Turner, J., O'Donnell, M. & Kuriyan, J. (2000). Crystal structure of the DNA polymerase processivity factor of T4 bacteriophage. J. Mol. Biol. 296, 1215-1223. 4. Huang, C. C., Hearst, J. E. & Alberts, B. M. (1981). Two types of replication proteins increase the rate at which T4 DNA polymerase traverses the helical regions in a single-stranded DNA template. J. Biol. Chem. 256, 4087-4094. 5. Stukenberg, P. T., Studwell-Vaughan, P. S. & O'Donnell, M. (1991). Mechanism of the sliding beta-clamp of DNA polymerase III holoenzyme. J. Biol. Chem. 266, 11328-11334. 6. Hingorani, M. M. & O'Donnell, M. (2000). Sliding clamps: a (tail)ored ®t. Curr. Biol. 10, R25-R29. 7. Maki, S. & Kornberg, A. (1988). DNA polymerase III holoenzyme of Escherichia coli. III. Distinctive processive polymerases reconstituted from puri®ed subunits. J. Biol. Chem. 263, 6561-6569. 8. Kaboord, B. F. & Benkovic, S. J. (1995). Accessory proteins function as matchmakers in the assembly of the T4 DNA polymerase holoenzyme. Curr. Biol. 5, 149-157. 9. Berdis, A. J. & Benkovic, S. J. (1997). Mechanism of bacteriophage T4 DNA holoenzyme assembly: the

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Kinetics of Loading the T4 Processivity Clamp 24. Naktinis, V., Onrust, R., Fang, L. & O'Donnell, M. (1995). Assembly of a chromosomal replication machine: two DNA polymerases, a clamp loader, and sliding clamps in one holoenzyme particle. II. Intermediate complex between the clamp loader and its clamp. J. Biol. Chem. 270, 13358-13365. 25. Hingorani, M. M. & O'Donnell, M. (1998). Toroidal proteins: running rings around DNA. Curr. Biol. 8, R83-R86. 26. Turner, J., Hingorani, M. M., Kelman, Z. & O'Donnell, M. (1999). The internal workings of a DNA polymerase clamp-loading machine. EMBO J. 18, 771-783. 27. Bloom, L. B., Turner, J., Kelman, Z., Beechem, J. M., O'Donnell, M. & Goodman, M. F. (1996). Dynamics of loading the beta sliding clamp of DNA polymerase III onto DNA. J. Biol. Chem. 271, 30699-30708. 28. Bertram, J. G., Bloom, L. B., Turner, J., O'Donnell, M., Beechem, J. M. & Goodman, M. F. (1998). Presteady state analysis of the assembly of wild type and mutant circular clamps of Escherichia coli DNA polymerase III onto DNA. J. Biol. Chem. 273, 2456424574. 29. Hingorani, M. M. & O'Donnell, M. (1998). ATP binding to the Escherichia coli clamp loader powers opening of the ring-shaped clamp of DNA polymerase III holoenzyme. J. Biol. Chem. 273, 24550-24563. 30. Hingorani, M. M., Bloom, L. B., Goodman, M. F. & O'Donnell, M. (1999). Division of labor ± sequential ATP hydrolysis drives assembly of a DNA polymerase sliding clamp around DNA. EMBO J. 18, 51315144. 31. Bertram, J. G., Bloom, L. B., Hingorani, M. M., Beechem, J. M., O'Donnell, M. & Goodman, M. F. (2000). Molecular mechanism and energetics of clamp assembly in Escherichia coli: the role of ATP hydrolysis when gamma complex loads beta on DNA. J. Biol. Chem, 275, 28413-28420. 32. Kaboord, B. F. & Benkovic, S. J. (1993). Rapid assembly of the bacteriophage T4 core replication complex on a linear primer/template construct. Proc. Natl Acad. Sci. USA, 90, 10881-10885. 33. Berdis, A. J. & Benkovic, S. J. (1996). Role of adenosine 50 -triphosphate hydrolysis in the assembly of the bacteriophage T4 DNA replication holoenzyme complex. Biochemistry, 35, 9253-9265. 34. Young, M. C., Weitzel, S. E. & von Hippel, P. H. (1996). The kinetic mechanism of formation of the bacteriophage T4 DNA polymerase sliding clamp. J. Mol. Biol. 264, 440-452. 35. Ason, B., Bertram, J. G., Hingorani, M. M., Beechem, J. M., O'Donnell, M., Goodman, M. F. & Bloom, L. B. (2000). A model for Escherichia coli DNA polymerase III holoenzyme assembly at primer/template ends. DNA triggers a change in binding speci®city of the gamma complex clamp loader. J. Biol. Chem. 275, 3006-3015. 36. Tsuchihashi, Z. & Kornberg, A. (1989). ATP interactions of the tau and gamma subunits of DNA polymerase III holoenzyme of Escherichia coli. J. Biol. Chem. 264, 17790-17795. 37. Vale, R. D. & Milligan, R. A. (2000). The way things move: looking under the hood of molecular motor proteins. Science, 288, 88-95. 38. Johnson, K. A. (1985). Pathway of the microtubuledynein ATPase and the structure of dynein: a comparison with actomyosin. Annu. Rev. Biophys. Biophys. Chem. 14, 161-188.

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39. Vale, R. D. (1996). Switches, latches, and ampli®ers: common themes of G proteins and molecular motors. J. Cell Biol. 135, 291-302. 40. Mandelkow, E. & Johnson, K. A. (1998). The structural and mechanochemical cycle of kinesin. Trends Biochem. Sci. 23, 429-433. 41. Sprang, S. R. (1997). G protein mechanisms: insights from structural analysis. Annu. Rev. Biochem. 66, 639-678. 42. Fink, A. L. (1999). Chaperone-mediated protein folding. Physiol. Rev. 79, 425-449. 43. Saibil, H. (2000). Molecular chaperones: containers and surfaces for folding, stabilising or unfolding proteins. Curr. Opin. Struct. Biol. 10, 251-258. 44. von Hippel, P. H. & Delagoutte, E. (2001). A general model for nucleic acid helicases and their ``coupling'' within macromolecular machines. Cell, 104, 177-190.

45. Jarvis, T. C., Ring, D. M., Daube, S. S. & von Hippel, P. H. (1990). ``Macromolecular crowding'': thermodynamic consequences for protein-protein interactions within the T4 DNA replication complex. J. Biol. Chem. 265, 15160-15167. 46. Johnson, K. A. (1986). Rapid kinetic analysis of mechanochemical adenosinetriphosphatases. Methods Enzymol. 134, 677-705. 47. Latham, G. J., Dong, F., Pietroni, P., Dozono, J. M., Bacheller, D. J. & von Hippel, P. H. (1999). Opening of a monomer-monomer interface of the trimeric bacteriophage T4-coded GP45 sliding clamp is required for clamp-loading onto DNA. Proc. Natl Acad. Sci. USA, 96, 12448-12453. 48. Barshop, B. A., Wrenn, R. F. & Frieden, C. (1983). Analysis of numerical methods for computer simulation of kinetic processes: development of KINSIM± a ¯exible, portable system. Anal. Biochem. 130, 134-145.

Appendix We have used the program ProFit (QuantumSoft Software) to obtain a simultaneous ®t of the results of our steady-state ATPase titrations to two equations that were created on the basis of the model shown in Figure 7 of the main text. This model can be represented as:

binding of DNA to the gp44/62-gp45 complex, kt.c.(t.c., ternary complex) is the ®rst-order rate constant for the steps that involve the ternary gp44/ 62-gp45-DNA complex (ATP hydrolysis and gp45 and DNA dissociation from gp44/62), and k44c. (44c., gp44/62 complex) is the ®rst-order rate constant for the post-hydrolysis conformational change that involves only the gp44/62 complex,

In this scheme, ES is the bound enzyme-substrate complex (gp44/62-ATP), EP is the bound enzyme-product complex (gp44/62-ADP), k45bind is the second-order rate constant for binding of gp45 to gp44/62, kb.c.(b.c., binary complex) is the ®rstorder rate constant for the pre-hydrolysis conformational change in the binary gp44/62-gp45 complex, kDNAbind is the second-order rate constant for

and which we speculate corresponds to ADP release. As stated in Discussion in the main text, all the rate constants we have used are apparent, in that they represent lower limits for the forward reactions of the various steps. Data from the titration of gp44/62 with p/t DNA in the presence of gp45 were ®tted to the following equation:

…1†

888

Kinetics of Loading the T4 Processivity Clamp

in which the concentration of free p/t DNA ([DNA]) was obtained from:

…2†

and the concentration calculated from:

of gp45

([45max]) was

…3†

Here [Etot] is the total concentration of the enzyme gp44/62, [45max] is the concentration of free gp45 at the point of maximal stimulation of the ATPase activity with p/t DNA, [DNA] is the concentration of free p/t DNA, and [DNAtot] is the total concentration of p/t DNA. [E
DNA] is the concentration of the gp44/62-DNA bound species (with or without gp45), [45tot] is the total concentration of gp45, and [E
45max] is the concentration of the gp44/62-gp45 bound species (with or without DNA) at the maximal stimulation of the ATPase activity by DNA. Similarly, data from the titration of gp44/62 with gp45 in the presence of p/t DNA were ®t to the following equation: …4†

where the concentration of free gp45 ([45]) was derived from:

…5†

and [DNAmax] was calculated from:

…6†

889

Kinetics of Loading the T4 Processivity Clamp

Here again, [45] represents the concentration of free gp45, [DNAmax] is the concentration of free p/t DNA at the point of maximal stimulation of the ATPase activity by gp45, [E
45] is the concentration of the gp44/62-gp45 bound species (with or without DNA), [DNAtot] is the total concentration of p/t DNA, and [E
DNAmax] is the concentration of the gp44/62-DNA bound species (with or without gp45) at the point of maximal stimulation of the ATPase activity by gp45. The equations were derived as follows. The catalytic rate of a substrate-saturated enzyme that is active only upon the binding of a cofactor (or where the basal rate of the enzyme in the absence of cofactor is not signi®cant) is given by: rate ˆ ‰Etot Š

1

1 kmax

‡

1 ‰CŠkbind

…7†

where [Etot] is the total concentration of enzyme, kmax is the maximal rate observed per unit of enzyme when cofactor binding is no longer ratelimiting (i.e. kmax is the ®rst-order rate constant for the unimolecular steps in the cycle), [C] is the concentration of free cofactor, and kbind is the relative (apparent) second-order rate constant for the binding of cofactor to the enzyme. The concentration of free cofactor can be derived from:

for the steps in which gp45 is involved, which, based on the model, include the pre-hydrolysis conformational change in the binary gp44/62gp45, the binding of p/t DNA to the binary gp44/ 62-gp45 complex, ATP hydrolysis, gp45 dissociation from gp44/62 and gp45 sliding off the p/ t DNA (Figure A1(a)). If we assume: (i) that p/t DNA and gp45 dissociation from gp44/62 are fast relative to the ATP hydrolysis step (then kt.c. will represent the rate of the hydrolysis step) or occur at the same time (then kt.c. represents the rate constant of the ATP hydrolysis step plus the rate constant for the dissociation of both p/t DNA and gp45 from gp44/ 62); and (ii) gp45 sliding off DNA is fast relative to the pre-hydrolysis conformational change; then the rate constants describing the two gp44/62 ATPase titrations with one of the two cofactors (in the presence of a constant excess of the other cofactor), can be written as follows (Figure A1(b)). For the titration of the ATPase activity of gp44/ 62 with p/t DNA in the presence of excess gp45 we may write: 1 1 ˆ kEC kt:c:

…10†

1 1 1 1 1 ˆ ‡ ‡ ‡ kmax ‰45Šk45bind kb:c: kt:c: k44c:

…11†

and:

‰CŠ ˆ ‰Ctot Š ÿ ‰ECŠ 1 ˆ ‰Ctot Š ÿ ‰Etot Š

1 kmax

kEC ‡

…8†

1 ‰CŠkbind

where [Ctot] is the total concentration of cofactor (free and bound), [EC] is the concentration of all the enzyme-cofactor bound species, and kEC is the ®rst-order rate constant for the unimolecular steps that involve the enzyme species that are bound to cofactor. If we substitute [C] derived from equation (8) into equation (7), the ®tting of a plot of ATPase rate as a function of the concentration of the titrating cofactor ([Ctot]) will provide the values of the ®tting parameters kmax, kEC, and kbind. We can relate these kinetic constants to speci®c steps based on a model derived from the results presented in the main text and elsewhere. The model is shown in Figure A1. For the titration of the ATPase activity of gp44/ 62 with p/t DNA in the presence of excess gp45, kEC represents the ®rst-order rate constant for the steps in which the p/t DNA is involved. These steps, based on the model shown in Figure A1, must include ATP hydrolysis and the dissociation of the p/t DNA from the gp44/62 loading complex. Similarly, when gp44/62 is titrated with gp45 in the presence of excess p/t DNA, kEC represents the ®rst-order rate constant

For the titration of the ATPase activity of gp44/ 62 with gp45 in the presence of excess p/t DNA we may write: 1 1 1 1 ˆ ‡ ‡ kEC kb:c: ‰DNAŠkDNAbind kt:c:

…12†

and: 1 kmax

ˆ

1 kb:c:

‡

1 1 1 ‡ ‡ ‰DNAŠkDNAbind kt:c: k44c:

…13†

In the above equations, we have included the rate constant for the binding of the non-titrating cofactor (the cofactor present in excess and at constant concentration during the titration), because the concentration of this cofactor that was used (especially where the non-titrating cofactor was gp45) was such that its rate of binding to the enzyme becomes partially rate-determing, at least under conditions where the titration approaches the ATPase rate plateau level. The rate of binding of the non-titrating cofactor to the enzyme is different at each titration point, depending on the concentration of the cofactor free in solution, which depends in turn on the concentration of the titrating cofactor. However, as stated above, this rate becomes signi®cant only when the titration approaches the plateau. Therefore, we can assume that the rate of binding of the non-titrating cofactor

890

Kinetics of Loading the T4 Processivity Clamp

Figure A1. A representation of the ®ve rate constants used in the ®tting of the steady-state ATPase titrations relative to the proposed model. kEC is the rate constant for the steps in which the enzyme-titrating cofactor-bound species are involved. kmax is the maximal rate observed per unit of enzyme when the titrating cofactor binding is no longer rate-limiting. The steps described by kEC for the titration of the ATPase activity of gp44/62 with gp45 in the presence of excess p/t DNA (‡gp45) are outlined in red. These steps involve the gp44/62-gp45 species (with or without DNA). The steps described by kEC for the titration of the ATPase activity of gp44/62 with p/t DNA in the presence of excess gp45 (‡DNA) are outlined in blue. These steps involve the gp44/62-p/t DNA species (with or without gp45). The steps described by kmax for both titrations, as indicated, are outlined in black. In (a) we show that some of the steps may occur simultaneaously (i.e. p/t DNA dissociation, gp45 sliding off p/t DNA and ADP release) and that the rate constants for these single steps cannot be distinguished. In (b) we show that if we make the indicated assumptions, the mechanism becomes the sum of ®ve discrete steps, with rate constants that can be de®ned by rewriting kEC and kmax in the ®tting equations appropriately for the two titrations. (See the text for further details.)

is constant during the titration and it is determined by the concentration of the free non-titrating cofactor present at the plateau of the titration. We have found, by ®tting simulated data, that this approximation does not affect the result appreciably; i.e. the ®tting parameters differ from the rate values used to simulate the data by only a few percentage points, which difference is less than the experimental error of the experiment (calculations not shown). The maximal concentration of the free non-titrating cofactor at the plateau of the titration is calcu-

lated as follows. For the titration with p/t DNA, the concentration of free gp45 present at the maximal stimulation of ATPase activity ([45max]) is derived from:

…14†

Kinetics of Loading the T4 Processivity Clamp

where [45tot] is the total concentration of gp45, [b.c.max] and [t.c.max] are the concentrations of the binary (gp44/62-gp45) complex and the ternary (gp44/62-gp45-p/t DNA) complex, respectively, at the maximal ATPase activity. For the titration with gp45, the concentration of free DNA present at the maximal stimulation of ATPase activity ([DNAmax]) is derived from:

891 where [DNAtot] is the total concentration of DNA. By substituting [45max] and [DNAmax] derived from equations (14) and (15) into [45] and [DNA] in equations (10) through (13), and then substituting these equations into equations (7) and (8), we obtained equations (1) through (6).

…15†

Edited by R. Ebright (Received 10 January 2001; received in revised form 4 April 2001; accepted 4 April 2001)