dCMP removal from correctly paired or frayed DNA.
To examine the kinetics of the exonuclease reaction, we prepared synthetic oligonucleotides to create defined primer and template strands of 26 and 45 bases, respectively. We designed one primer-template duplex to be complementary throughout the double-stranded region and one with eight mismatches on the 3′ end of the primer to create a frayed duplex. The rationale for using a frayed duplex is based upon observations suggesting that the rate of excision is usually limited by the rate of transfer of the primer strand from the polymerase site to the exonuclease site and that a frayed duplex favors partitioning into the exonuclease site so that the observed rate more closely approaches the intrinsic rate of hydrolysis (5
). We chose a length of eight mismatched base pairs at the end of the frayed duplex because previous work had shown that the effects of fraying on the kinetics of excision were saturated after six to eight nucleotides.
We first examined the rates of removal of the natural nucleotide (dCMP) from the primer 3′ terminus in reactions with mixtures containing either a correctly paired or frayed DNA substrate. The wild-type Pol γ holoenzyme was preincubated with either correctly paired or frayed DNA, under conditions in which the enzyme was in molar excess relative to the DNA, and rapidly mixed with Mg2+
in a chemical quench-flow to start the reaction. The exonuclease reaction was quenched at various times with 0.5 M EDTA. The concentration of the starting material (full-length primer) was plotted as a function of time (Fig. 1
). The removal of dCMP from correctly base-paired DNA was biphasic, with rates of 0.86 ± 0.35 s−1
and 0.016 ± 0.004 s−1
and amplitudes of 21 ± 3 nM and 70 ± 3 nM, respectively. Because the experiment was performed with the enzyme in excess compared to the DNA, we can eliminate models invoking multiple turnovers to account for the slow phase. Rather, the amplitude of the fast phase may represent the fraction of primer strand that was in the exonuclease site of Pol γ when the reaction was initiated, while the slow, larger-amplitude phase may represent the rate at which the remaining primer partitions into the exonuclease site. The exonuclease reaction observed using the frayed DNA substrate was also biphasic, with rates of 1.6 ± 0.5 s−1
and 0.1 ± 0.07 s−1
and amplitudes of 60 ± 10 nM and 25 ± 10 nM, respectively.
In comparing the results for the frayed duplex to those for the fully complementary duplex, the rate of the fast phase was found to increase only slightly but the amplitude for the fast phase increased significantly, consistent with our working model. The rate of excision for the slow phase of the reaction with frayed DNA was an order of magnitude higher than the rate of excision for the slow phase with the correctly paired DNA, suggesting that the primer strand melting and/or transfer was faster with DNA comprising a region of 3′ mismatches. We interpret these data to mean that the intrinsic rate of hydrolysis of dCMP from the 3′ end of a primer is greater than or equal to 1.6 s−1, while the rate of excision of dCMP from properly base-paired DNA is limited by the transfer of the primer from the polymerase site at a rate 100-fold lower (0.016 s−1).
ddCMP removal from correctly paired or frayed DNA.
The removal of ddCMP was examined using a DNA sequence identical to the one in the procedure described above except that the primer was terminated with ddCMP at the 3′ end. As described above, two reactions were performed, one with correctly paired DNA and the other with frayed DNA, both with the enzyme in excess relative to the DNA. The concentration of the substrate was plotted as a function of time (Fig. 2
). Unlike the removal of the natural nucleotide, the removal of ddCMP occurred in a single phase and could be fitted to a single exponential equation. The rate of the removal of ddCMP from correctly paired DNA was (3 ± 0.4) × 10−4
, about 50-fold lower than the rate of dCMP removal in the slow phase. The removal of ddCMP from frayed DNA was only 15-fold faster than the removal from correctly paired DNA and occurred at a rate of (4.4 ± 0.5) × 10−3
. Both reactions went to completion at rates much lower than the rate of dissociation of DNA from the enzyme active site (∼0.02 s−1
) measured previously (16
), implying that the vast majority of DNA dissociates from the enzyme and rebinds multiple times during the time course of excision. Nonetheless, by examining the reaction with the enzyme in excess, the observed rate is limited only by the reactions occurring at the active site. Our data suggest that at equilibrium, the majority of the correctly paired ddCMP-terminated primer remains in the polymerase active site. Moreover, the rate of hydrolysis to excise ddCMP, defined by the removal from the frayed primer-template combinations, is about 360-fold lower than that seen for dCMP. Therefore, it appears that the 3′-hydroxyl group on the primer strand plays a significant role in the binding and/or alignment of the terminal nucleotide at the exonuclease site. Moreover, as described previously, ddCTP binds more tightly than dCTP at the polymerase site, and it is reasonable to suggest that similar tighter binding at the polymerase site after incorporation shifts the partitioning away from the exonuclease site. However, the binding of the next correct nucleotide will alter the distribution among various DNA states bound to the enzyme.
Removal of ddC in the presence of dNTPs.
The binding of the next correct deoxynucleoside triphosphate (dNTP) to a primer-template lacking a 3′-OH molecule forms a ternary complex, stabilizing the primer at the polymerase site (17
), thereby attenuating the migration of the primer to the exonuclease site and reducing the observed hydrolysis rate (5
). Therefore, after ddCMP is incorporated, the rate of excision may be lowered by the presence of dNTPs. As shown in Fig. 3
, ddCMP removal from correctly paired DNA in the presence of 100 μM dNTPs was slowed by almost fourfold to a rate of (8.0 ± 0.7) × 10−5
. The rate of removal of ddCMP from frayed DNA in the presence of dNTPs [(6.3 ± 0.3) × 10−3
] was comparable to that seen in their absence [(4.4 ± 0.5) × 10−3
]. Therefore, the tight ternary complex cannot be formed by the binding of the next correct nucleotide to a frayed duplex. The rationale for adding all four dNTPs at 100 μM was based upon the fact that the inclusion of only the next correct base would have made the data analysis unnecessarily complex. For example, if only the next correct nucleotide was present during the excision reaction, the removal of ddCMP would have likely been followed by the misincorporation of the single dNTP present in the solution and would have led to the same length of primer with which the reaction was initiated. The presence of the other three nucleotides allowed rapid polymerization of the primer following the slow excision of ddCMP. Therefore, the disappearance of the primer was easily quantified in this format. In any event, the presence of all four nucleotides would be a more accurate representation of the true physiological circumstance than the presence of only one nucleotide and therefore provides the best estimate for the in vivo rate of excision to date. Results of previous studies suggest that the effect is specific to the next correct nucleotide (13
). Table 1
provides a summary of the data obtained for the removal of dCMP and ddCMP under the various conditions examined.
Of the eight FDA-approved nucleoside analogs, ddC is the best substrate for incorporation by Pol γ and is removed with the least efficiency by the proofreading exonuclease, and these combined effects lead to the marked toxicity seen clinically. The individual contributions from incorporation and exonuclease removal to the in vivo toxicity of ddC can be evaluated by computing a toxicity index as described previously (15
). The toxicity index expresses the increase in time required to replicate a given segment of DNA by comparing the rate of incorporation of the analog leading to chain termination to the rate of excision leading to rescue and the resumption of synthesis. The toxicity index is defined by the following equation: toxicity index = (kpol/kexo
), where D
is the discrimination (the ratio of specificity constants) against the analog triphosphate (AnaTP), kpol
is the rate of incorporation of the normal correct nucleotide (dNTP), and kexo
is the rate of excision of the analog monophosphate from the primer. The toxicity index for ddC is approximately 50,000 when [AnaTP] equals [dNTP], implying that it would take 50,000-fold longer to replicate the human mitochondrial genome if the concentration of ddCTP were equal to that of dCTP in the mitochondria. Although the toxicity in vivo will be attenuated by the actual [AnaTP]/[dNTP] ratio in the mitochondria, this high calculated toxicity index clearly points to the high potential for mitochondrial toxicity and explains the toxicity of ddC seen clinically.
These data correct our previous attempt to measure the rate of excision of ddC (7
), in which we failed to observe any hydrolysis after 12 h of incubation and therefore placed only a lower limit on the rate of excision. It is not known why the previous measurements failed, but it is possible that the enzyme, used at a lower concentration than that in the present study, degraded over the long time course of the experiment or that the large excess of DNA may have bound to the enzyme such that hydrolysis was inhibited. Although the steady-state approach should have provided a valid measurement in this case, the single-turnover experiments reported here with the enzyme in slight excess relative to the DNA circumvent many of the possible difficulties.
From our new data, it appears that both the transfer of the primer from the polymerase to the exonuclease site and the low rate of hydrolysis at the exonuclease site contribute to the slow removal of ddCMP. The rate of hydrolysis of ddCMP is about 360-fold lower than that of the natural nucleotide, suggesting that the 3′-hydroxyl plays a major role in the binding and/or alignment of the 3′ terminus at the exonuclease active site. In vivo, ddC would be correctly base paired in most instances; therefore, the rate of excision would be hindered due to the fact that the rate of primer strand melting and/or transfer from the polymerase site to the exonuclease site was also low. Moreover, the addition of the next correct nucleotide stabilized binding at the polymerase site and lowered the rate of excision fourfold to yield a half-life of about 2.4 h.
The rate of removal of the eight FDA-approved analogs varies from 0.0003 s−1
for ddC to 0.015 s−1
for (−)3TC, as summarized in Fig. 4
. For comparison, the rate of removal of correctly paired dCMP is 0.016 s−1
. Conclusions about structure-function relationships pertaining to the rates of removal of these nucleoside analogs are highly speculative because there currently is no crystal structure available for Pol γ, but several notable generalizations can be drawn. There is only a 2.3-fold difference between the highest and lowest rates of excision of ddC, d4T, AZT, ddATP, and PMPA, but the rates of removal of CBV, (−)FTC, and (−)3TC are significantly higher. The analogs (−)FTC and (−)3TC are the most efficiently removed; each has a sulfur atom in the ribose ring and an unnatural l
-(−) nucleoside configuration, which may destabilize binding at the polymerase site. For example, it is known that mutant forms of HIV reverse transcriptase resistant to 3TC have a structural substitution that is thought to lead to steric blocking of binding at the polymerase site (9
It is notable that FIAU, which does contain a 3′-hydroxyl, was removed at a rate of 0.06 s−1
, six times faster even than (−)3TC (7
). Unfortunately, FIAU caused severe lactic acidosis and the deaths of several patients during the initial clinical trial (19
). Although the presence of a fluorine atom in the ribose ring inhibits the extension on top of FIAU approximately 500-fold, the rate of polymerization of the next correct nucleotide is still higher than the excision rate (15
), leading to the stable incorporation of FIAU.
Efforts at designing more effective nucleoside analogs can incorporate the notion that the rate of removal of these chain terminators by Pol γ may play a significant role in their overall toxicities. Ideally, nucleoside analogs would be designed to mimic a mismatch so that they would be well discriminated against by Pol γ but also be removed at a high rate because primer strand melting and/or transfer would be faster. Furthermore, it appears that a better 3′-OH mimic could increase the actual rate of hydrolysis. It is noteworthy that each of the three most toxic nucleotide analogs, ddCTP, ddATP (the active metabolite of dideoxyinosine), and d4TTP, bind more tightly during polymerization than the corresponding natural nucleotides, and this tighter binding contributes significantly to their specificity constants governing incorporation, leading to increased toxicity (15
). Moreover, these three analogs are the slowest to be excised by the proofreading exonuclease. It is reasonable to suppose that their greater affinities at the polymerase site reduce the rate and/or equilibrium constants governing partitioning to the exonuclease site. If so, this one feature, tighter binding at the polymerase site, contributes doubly to their net toxicity by increasing rates of incorporation while decreasing rates of excision. Correlations of toxicity with structure imply that discrimination by the mitochondrial DNA polymerase requires a more substantial steric handle than that afforded by these three simple nucleoside analogs. Other less toxic analogs differ substantially enough in their structural mimic of the deoxyribose moiety and allow significant discrimination by the mitochondrial polymerase, while retaining sufficient similarity to allow efficient incorporation by HIV reverse transcriptase. The ongoing search for new drugs must continue to find a balance between these two opposing structural constraints underlying effectiveness versus toxicity. We remain confident that the assays developed here and in our earlier work (15
) provide accurate, quantifiable parameters to assess the effectiveness and toxicity and thus the therapeutic index of new drugs under development.