Piattaforme di colture ingegnerizzate per ricerca e sviluppo farmaceutico

Engineered cell-culture platforms are increasingly expected to serve two goals at once: reproduce key biophysical aspects of native microenvironments (outside-in mechanobiology) and remain compatible with the throughput, automation, and imaging demands of pharmaceutical screening. This thesis aims to address that gap by developing and validating a set of practical, imaging-forward platforms that enable (i) stiffness-controlled culture in standard multiwell plates, (ii) scalable, geometry-controlled collective migration assays, and (iii) multiscale intracellular rheology readouts that can be integrated with traction force microscopy (inside-out mechanics). First, we introduce HYDRA, a robotic liquid-handling strategy that casts thin fish-gelatin hydrogel films directly inside commercial high-throughput plates while minimizing meniscus-driven curvature. HYDRA leverages a dispense–reaspirate operation to leave a micrometric boundary layer that can be enzymatically crosslinked, yielding substrates within a physiologically relevant stiffness range while remaining compatible with high-resolution microscopy. The method produces hydrated films of ~10–30 µm, a range chosen to balance mechanical relevance with imaging quality. HYDRA scales to imaging-grade 384-well plates using ~1 µL working volumes and includes an automated plate-wide morphology quality-control workflow. In 384-well plates, this QC pipeline supports rapid classification of gel shape and wall-contact artifacts, enabling high usable yields and demonstrating practical feasibility for imaging-based high-content screening, including long-term live imaging and subcellular-resolution confocal readouts. Second, to expand beyond static phenotyping, we develop a scalable migration assay based on two-step photopatterning to create reproducible “wound” geometries without physical barriers. By optimizing exposure conditions and pattern design, this approach aims to increase reproducibility and format flexibility compared with traditional scratch and insert-based assays, while remaining compatible with automated microscopy and systematic variation of wound geometry and boundary conditions. Third, we advance next-generation tools for multiscale intracellular rheology using single-particle tracking microrheology (SPT) in engineered HaCaT keratinocytes. We first established a coupled workflow that co-registers genetically encoded multimeric nanoparticles (GEMs)-based intracellular nanorheology with traction force microscopy on compliant substrates, enabling intracellular viscoelastic proxies to be interpreted alongside bead displacements and reconstructed traction maps within a TFM-compatible imaging design. We then implement a dual-tracer strategy combining GEMs and fluorescent lipid droplets (LDs) to probe complementary length scales while preserving a tight fluorescence channel budget. We introduce a cytometry-inspired, control-anchored classification framework: a transparent size-based gate (≈3 µm) and a probabilistic classifier separate GEM-like and LD-like trajectories, preventing mixed-population artifacts and maintaining interpretability. Finally, we demonstrate that MSD-derived rheology descriptors are acquisition-conditioned, and we define probe-specific stability regimes and QC-informed reporting strategies to improve reproducibility and cross-condition comparability. Together, these platforms provide a coherent toolkit for mechanobiology-oriented pharmaceutical R&D: HYDRA enables stiffness-controlled, imaging-compatible screening substrates; photopatterning enables scalable migration phenotyping with controllable geometry; and probe- and acquisition-aware SPT microrheology extends intracellular mechanical readouts in a way that can be integrated with traction-based force context.