Experimental Overview
The real-time PCR antibiogram utilizes antimicrobial exposure, preanalytic removal of heme and human background DNA, and colony PCR to assess pathogen susceptibility [20], [21]. The optimized protocol is described below in the Materials and Methods Section. Briefly, 1 mL of spiked blood (∼100 CFU/mL) is added to 9 mL of growth medium and incubated for 9 hours in various antibiotic environments. The sample is fractionated to separate red blood cells that contain heme, a PCR inhibitor. The supernatant, which consists of bacteria and mammalian cells, is pelleted and decanted. The pellet is then resuspended in mammalian lysis buffer and treated with DNase. This technique removes human DNA found in white blood cells from the sample, thus enhancing the sensitivity of detection. This is an essential part of this protocol as excess background human DNA can saturate the PCR amplification curves when using intercalating fluorophores. The sample is spun-down and the unseen bacterial cell pellet is washed in reticulocyte saline (RS) buffer. Preparation concludes by adding 2 µL of the sample directly to the PCR plate as template. This bacterial isolation method takes approximately 2–3 hours of manual labor, but could be automated to decrease sample preparation time and multiplexed for high-throughput testing in a clinical diagnostic laboratory setting.
A target pathogen concentration of ∼100 CFU/mL was chosen to emulate levels found clinically in sepsis cases [22]–[24]. For bacteremia, greater than 50% of cases are low-grade (100 CFU/mL), thus necessitating a detection sensitivity of at least 100 CFU/mL.