Quantifying single-cell biomechanics: a comparative analysis of experimental methods and theoretical models

This review presents a comprehensive overview of methodologies employed to measure and quantify the biomechanical properties of single cells, focusing on techniques that provide insights into cellular deformation and force response. Understanding cell mechanics is crucial for advancing our knowledge of cellular behavior, disease progression, and therapeutic innovation in contexts such as cancer metastasis, tissue engineering, and regenerative medicine. We critically examine five primary experimental platforms: atomic force microscopy (AFM), dielectrophoresis (DEP), optical tweezers, micropipette aspiration, and hydrodynamic-induced deformation. For each method, we detail the underlying theoretical frameworks, experimental configurations, advantages, limitations, and application scope. Contact-based techniques (AFM, micropipette aspiration) provide high-resolution measurements of localized stiffness and viscoelasticity, while non-contact approaches (DEP, hydrodynamic stretching, optical tweezers) enable high-throughput analysis with minimal physical disruption. Furthermore, we evaluate the mechanical models used to extract biomechanical parameters and highlight modern data extraction processes, including automated image analysis. Crucially, this review addresses the significant discrepancies in measured properties that arise across different methods due to varying loading rates, boundary conditions, and model assumptions. By synthesizing these experimental and theoretical considerations, this review aims to guide the standardization and refinement of single-cell biomechanics for both fundamental research and translational biomedical applications.