Deregulation in uterine contractility can cause common pathological disorders of the female reproductive system, including preterm labor, infertility, inappropriate implantation, and irregular menstrual cycle. A better understanding of human myometrium contractility is essential to designing and testing interventions for these important clinical problems. Robust studies on the physiology of human uterine contractions require in vitro models, utilizing a human source. Importantly, uterine contractility is a three-dimensionally (3D)-coordinated phenomenon and should be studied in a 3D environment. Here, we propose and assess for the first time a 3D in vitro model for the evaluation of human uterine contractility. Magnetic 3D bioprinting is applied to pattern human myometrium cells into rings, which are then monitored for contractility over time and as a function of various clinically relevant agents. Commercially available and patient-derived myometrium cells were magnetically bioprinted into rings in 384-well formats for throughput uterine contractility analysis. The bioprinted uterine rings from various cell origins and patients show different patterns of contractility and respond differently to clinically relevant uterine contractility inhibitors, indomethacin and nifedipine. We believe that the novel system will serve as a useful tool to evaluate the physiology of human parturition while enabling high-throughput testing of multiple agents and conditions.

584 Background: Tumor genotyping using fluid samples such as blood can potentially allow tracking of dynamic changes in mutational profiles over time and allow better access than biopsies. We present a method to detect somatic mutations from a blood draw, where circulating tumor cell (CTC) enrichment above 10% of total cell numbers allows the use of standard gene panels typically used to analyze tissue-based biopsies. Methods: Analytical samples were obtained from 9 prostate cancer (PC) patients and 6 renal cell cancer (RCC) patients, followed by CTC enrichment using the IsoFlux System. Cells were lysed and DNA amplified by whole genome amplification (WGA) using the NGS Kit (Fluxion Biosciences) and quantified via qPCR. Samples were enumerated to determine CTC load, with CTCs defined as CK+, CD45- nucleated cells (DAPI+). Next-generation sequencing was performed using 3 targeted cancer panels on the Ion torrent PGM platform: the Ion ampliseq cancer hotspot panel (50 genes; 6 PC samples), the Oncomine (143 genes; 3 PC samples), and a 29-gene panel of actionable mutations in RCC (6 samples). Data was analyzed using a customized variant calling/filtering pipeline based on standard alignment and variant calling tools. Variant filtering and functional interpretation was performed using VarSeq. All data was analyzed in a blinded manner. Results: Our method was able to isolate CTCs from all patient samples. Whole genome amplified DNA concentration was at a range of 25-164 ng/µL (median, 69) in PC and 29-180 ng/µL (median, 69) in RCC. CTCs were recovered in 2.9-33.7% (median, 10.5%) of PC samples and 1.9-33% (median, 14.5%) of RCC samples After WGA, we found 1 variant/sample using hotpot, 12/sample using Oncomine, and 3/sample using the RCC panel. Conclusions: Our assay consistently detected somatic variants from blood draw using standard gene panels in both PC and RCC. Obtaining repeat tumor biopsies from patients during treatment and/or at time of progression is both challenging and impractical from a clinical perspective. Our assay provides molecular characterization using standard blood draws and will be prospectively validated in clinical trials.