RAW macrophages were grown in Dulbecco modified Eagle medium containing l-glutamine and sodium pyruvate (PAA Laboratories) supplemented with 10% fetal bovine serum. About 2.5 × 10⁴ cells were seeded into each well of a 48-well plate and were infected with green fluorescent protein (GFP)-expressing M. tuberculosis (MTB_NDT1) at a multiplicity of infection (MOI) of 10:1 for 4 h. After infection, the monolayers were washed three times with fresh medium to remove nonadherent bacteria and then exposed to fresh medium with or without isoniazid (0.75 μg/ml), faropenem (7, 28, or 56 μg/ml), or meropenem (5, 25, or 50 μg/ml) plus clavulanate (2.5 μg/ml). Medium (containing antibiotics) was changed every 24 h. At various time points, macrophages were washed once with 1× PBS and then lysed using 0.5% Triton X-100. The macrophage lysates were diluted in PBS containing 0.05% Tween 80, serial dilutions were plated on 7H10 agar, and bacterial CFU were enumerated after incubation of the plates at 37°C for 3 to 4 weeks. For analysis of the infected macrophages by flow cytometry, the cells were washed once with 1× PBS before being resuspended in PBS. Cells were analyzed by flow cytometry using a BD Accuri C6 flow cytometer. The threshold was set at 80,000, gates were drawn to exclude clumps and debris, and 50,000 gated events were acquired for each sample. RAW macrophages infected with nonfluorescent M. tuberculosis were used to determine the background fluorescence gates. The fraction of infected cells was calculated from the number of events exhibiting fluorescence above this background level.