Quantitative Reverse Transcriptase PCR (RT-PCR)

Total RNA (5 μg) was used as the template in the cDNA-synthesis reactions with random primers and Superscript III reverse transcriptase (Applied Biosystems). The resultant cDNAs were used (at a 1:20 dilution) to detect the level of endogenous BRCA1, BRCA2, and PD-L1 mRNA expression by quantitative PCR (qPCR). Accurate quantitation was achieved using standard curves generated by serially diluting a known quantity of RNA from an in vitro transcription reaction and performing TaqMan qPCR with the dilution along with the cell samples. Quantitative analysis of mRNA expression was performed with the StepOne^TM Real-Time PCR System (ABI). The primers and TaqMan probes used for the analyses were designed using the manufacturer’s software, Primer Express. The following primers were used: BRCA1 (HS01556193), BRCA2 (HS00609073), and GAPDH (HS99999905). No-reverse-transcription (no-RT) control reactions were performed with 100 ng of total RNA from each individual sample as a template to ensure that amplification was not caused by DNA contamination. No signal was detected in the no-RT controls. Target gene mRNA expression was assessed by real-time RT-PCR. The reference gene GAPDH was used as the internal control for RNA quality. All quantitative analyses were performed in duplicate to assess the consistency of the results. The relative expression levels of the target gene, normalized to GAPDH expression, were calculated as ΔC_t = C_t (target) – C_t (GAPDH). The ratio of the number of copies of the target gene mRNA to the number of copies of GAPDH was then calculated as 2^–Ct × K (K = 10⁶, a constant).