Modular DNA tile-based self-assembly is a versatile way to engineer basic tessellation patterns on the nanometer scale, but it remains challenging to achieve high levels of structural complexity. We introduce a set of general design principles to create intricate DNA tessellations by employing multi-arm DNA motifs with low symmetry. We achieved two novel Archimedean tiling patterns, (4.8.8) and (3.6.3.6), and one pattern with higher-order structures beyond the complexity observed in Archimedean tiling. Our success in assembling complicated DNA tessellations demonstrates the broad design space of DNA structural motifs, enriching the toolbox of DNA tile-based self-assembly and expanding the complexity boundaries of DNA tile-based tessellation.
Phosphatidylinositol kinases (PIKs) are key enzymatic regulators of membrane phospholipids and membrane environments that control many aspects of cellular function, from signal transduction to secretion, through the Golgi apparatus. Here, we have developed a photoreactive "clickable" probe, PIK-BPyne, to report the activity of PIKs. We investigated the selectivity and efficiency of the probe to both inhibit and label PIKs, and we compared PIK-BPyne to a wortmannin activity-based probe also known to target PIKs. We found that PIK-BPyne can act as an effective in situ activity-based probe, and for the first time, report changes in PI4K-IIIβ activity induced by the hepatitis C virus. These results establish the utility of PIK-BPyne for activity-based protein profiling studies of PIK function in native biological systems.
Measuring molecular binding to membrane proteins is critical for understanding cellular functions, validating biomarkers, and screening drugs. Despite the importance, developing such a capability has been a difficult challenge, especially for small-molecule binding to membrane proteins in their native cellular environment. Here we show that the binding of both large and small molecules to membrane proteins can be quantified on single cells by trapping single cells with a microfluidic device and detecting binding-induced cellular membrane deformation on the nanometer scale with label-free optical imaging. We develop a thermodynamic model to describe the binding-induced membrane deformation, validate the model by examining the dependence of membrane deformation on cell stiffness, membrane protein expression level, and binding affinity, and study four major types of membrane proteins, including glycoproteins, ion channels, G-protein coupled receptors, and tyrosine kinase receptors. The single-cell detection capability reveals the importance of local membrane environment on molecular binding and variability in the binding kinetics of different cell lines and heterogeneity of different cells within the same cell line.
Most drugs work by binding to receptors on the cell surface. Quantification of binding kinetics between drug and membrane protein is an essential step in drug discovery. Current methods for measuring binding kinetics involve extracting the membrane protein and labeling, and both have issues. Surface plasmon resonance (SPR) imaging has been demonstrated for quantification of protein binding to cells with single-cell resolution, but it only senses the bottom of the cell and the signal diminishes with the molecule size. We have discovered that ligand binding to the cell surface is accompanied by a small cell membrane deformation, which can be used to measure the binding kinetics by tracking the cell edge deformation. Here, we report the first integration of SPR imaging and cell edge tracking methods in a single device, and we use lectin interaction as a model system to demonstrate the capability of the device. The integration enables the simultaneous collection of complementary information provided by both methods. Edge tracking provides the advantage of small molecule binding detection capability, while the SPR signal scales with the ligand mass and can quantify membrane protein density. The kinetic constants from the two methods were cross-validated and found to be in agreement at the single-cell level. The variation of observed rate constant between the two methods is about 0.009 s−1, which is about the same level as the cell-to-cell variations. This result confirms that both methods can be used to measure whole-cell binding kinetics, and the integration improves the reliability and capability of the measurement.
scite is a Brooklyn-based organization that helps researchers better discover and understand research articles through Smart Citations–citations that display the context of the citation and describe whether the article provides supporting or contrasting evidence. scite is used by students and researchers from around the world and is funded in part by the National Science Foundation and the National Institute on Drug Abuse of the National Institutes of Health.
hi@scite.ai
10624 S. Eastern Ave., Ste. A-614
Henderson, NV 89052, USA
Copyright © 2024 scite LLC. All rights reserved.
Made with 💙 for researchers
Part of the Research Solutions Family.