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The cells responsible for the second phase decay of HIV-1 viremia following the initiation of antiretroviral therapy have yet to be identified. A dynamic model that considers where drugs act in the virus life cycle places constraints on candidate cell types. In this regard, the rapid drop in viremia in patients starting regimens containing the integrase inhibitor raltegravir is of particular interest. We show here that the time delay between reverse transcription and integration is short in differentiated macrophages, making these cells poor candidates for the second phase compartment under the assumptions of standard models of viral dynamics.
Pioneering studies of viral dynamics1–5 established that the decay in plasma HIV-1 levels following initiation of treatment depends on the turnover rate of productively infected cells. The classic model assumes that antiretroviral drugs completely stop new infection, revealing the decay rates of previously infected cells. The rapid initial decay in viremia reflects the short half-life (1 day) of productively infected CD4+ T lymphoblasts, the primary target cells for HIV-1. When most of these cells have died, a second slower phase becomes apparent, reflecting virus production by another as yet unidentified population of infected cells with a half-life of ~14 days.
Differences in decay rates of viremia in patients starting different regimens can provide insight into the nature of the second phase compartment. A general model developed by Sedaghat et al. describes how decay dynamics are influenced by the stage in the virus life cycle at which an antiviral drug acts, termed a stage effect (Fig. 1A).6 Assuming full efficacy, drugs acting later after virus entry can produce a more rapid decay rate in viremia than drugs acting earlier in the life cycle. This effect is observed if the sum of the early stage infected cell death rate and the rate of conversion of early stage infected cells to late stage, productively infected cells is less than the decay rate of productively infected cells. If not, decay dynamics may still be altered by the fact that for drugs acting earlier in the life cycle there is a shoulder period before decay begins (Fig. 1B), reflecting the decay of early stage infected cells, either through death or progression to late stages of infection.6,7 The length of this shoulder can be estimated as 1/(δM1+kM), where δM1 is the decay rate of the early stage cells and kM is the rate of conversion to late stage cells.
This model directly reflects the stepwise nature of viral replication, and as long as a fundamental assumption of complete inhibition is met, it can be used to evaluate candidate second phase cell types in situations in which decay dynamics differ with different drug regimens. In treatment-naive patients, viremia falls below the limit of detection more quickly in the patients starting regimens including the integrase inhibitor raltegravir (RAL) than in patients starting comparable regimens including the reverse transcriptase (RT) inhibitor efavirenz (EFV) in place of RAL.8 Murray and colleagues9 suggested that the rapid drop in viremia in patients on RAL is due to a smaller second phase compartment in the presence of RAL rather than a change in the first or second phase decay rates. In this situation, the rapid first phase decay continues for longer before virus production from the second phase compartment becomes quantitatively dominant. The end result is that viremia falls below the limit of detection more quickly in patients on RAL.
As is illustrated in Fig. 1B, the lack of a shoulder effect could contribute to lower apparent second phase viremia in patients starting RAL. According to the model described in Fig. 1A, second phase viremia on RAL is suppressed by a factor of δM2/(δM1+kM) relative to second phase viremia on EFV, where δM2 is the decay rate of productively infected second phase cells. The derivation of this effect involves considering the limit of the ratio between the model predictions of second phase viremia on RAL and second phase viremia on EFV as time increases.6,7 Essentially the reduced second phase viremia on RAL reflects the fact that second phase decay at rate δM2 begins as soon as an RAL-based regimen is started while decay is delayed by the shoulder period equal to 1/(δM1+kM) in patients starting an EFV-based regimen. By this analysis, the observed 70% reduction in second phase viremia9 can be completely explained by a stage effect if the transition from early to late stages of infection in second phase cells (kM) is slow (t1/2 >6 days), so that infected cells can accumulate in an EFV-resistant, RAL-susceptible state (M1 cells, Fig. 1A).
Based on susceptibility to infection, turnover rate, and resistance to HIV-1 cytopathic effects, the macrophage represents a reasonable candidate for the second phase compartment. To determine whether terminally differentiated macrophages meet the kinetic criteria described above, we measured kM in monocyte-derived macrophages (MDM) using time of addition experiments.10 The completion of reverse transcription was slower in MDM than in CD4+ T lymphoblasts (Fig. 1C). However, the interval between completion of reverse transcription, measured by loss of EFV-mediated inhibition, and completion of integration, measured by loss of RAL-mediated inhibition, was short in both cell types (~0.14 days). If this interval is an accurate reflection of the transition from M1 to M2 cells, then kM for macrophages is approximately 7.14 day–1. Thus for reasonable values of δM1 (0.69–0.05 day–1),7,10 the shoulder is short (3.1–3.3h), and the reduction in second phase viremia on RAL due to the stage effect is small (0.6–0.7%).
If the fundamental assumption of viral dynamics models is correct (complete or near complete inhibition of ongoing replication) and if macrophages derived from monocytes by in vitro differentiation accurately model infection in vivo, then differentiated macrophages do not meet the kinetic criteria for the second phase compartment. It is possible that the fundamental assumption is not satisfied and that the virus continues to replicate in patients on HAART. However, this assumption is strongly supported by recent pharmacodynamic11 and treatment intensification studies,12 and thus alternatives for the second phase compartment should be considered. The second phase may reflect infection of monocytes, which differentiate into macrophages after leaving the circulation. Integration is delayed in monocytes, occurring days not hours after integration.13 In resting CD4+ T cells, integration may be delayed by a block in nuclear import.9,10,14–18 Both cell types thus represent candidates for the second phase compartment. Interestingly, the length of the shoulder (Fig. 1B) is also influenced by the decay of early stage infected cells (δM1). In in vitro studies,10 the decay rate of unintegrated viral genomes in resting CD4+ T cells is too rapid (t1/2=1 day) to be consistent with the observed second phase decay. However, in vivo studies of the loss of recoverable virus from resting CD4+ T cells following initiation of therapy suggest a slower decay.17
These results highlight the importance of definitively identifying the cells responsible for the second phase decay. While macrophages have often been considered as the source, the measured kinetics of replication in differentiated macrophages are not consistent with predictions from standard models of viral dynamics, and thus other cell types should be considered.
This work was supported by the Doris Duke Charitable Foundation and by NIH Grants AI51178 and AI081600. A.S. was supported by NIH Grants T32 AI07247 and T32 AI007291.
No competing financial interests exist.