生物物理学报:英文版最新文献

Liquid-liquid phase separation (LLPS) is an emerging phenomenon involved in various biological processes. The formation of phase-separated condensates is crucial for many intrinsically disordered proteins to fulfill their biological functions. Using the recombinant protein to reconstitute the formation of condensates in vitro has become the standard method to investigate the behavior and function of LLPS. Meanwhile, there is an urgent need to characterize the LLPS in living cells. Importantly, condensates formed through LLPS at physical relevant concentrations are often smaller than the optical diffraction limit, which makes precise characterization and quantification inaccurate due to the scatter of light. The booming development of super-resolution optical microscopy enables the visualization of multiple obscured subcellular components and processes, which is also suitable for the LLPS research. In this protocol, we provide step-by-step instructions to help users take advantage of super-resolution imaging to depict the morphology and quantify the molecule number of endogenous condensates in living cells using RNA Pol II as an example. This streamlined workflow offers exceptional robustness, sensitivity, and precision, which could be easily implemented in any laboratory with an inverted total internal reflection microscope. We expect that super-resolution microscopy will contribute to the investigation of both large and tiny condensates under physiological and pathological conditions and lead our understanding of the mechanism of LLPS to a higher and deeper layer.

Liquid-liquid phase separation (LLPS) causes the formation of membraneless condensates, which play important roles in diverse cellular processes. Currently, optical microscopy is the most commonly used method to visualize micron-scale phase-separated condensates. Because the optical spatial resolution is restricted by the diffraction limit (~200 nm), dynamic formation processes from individual biomolecules to micron-scale condensates are still mostly unknown. Herein, we provide a detailed protocol applying dual-color fluorescence cross-correlation spectroscopy (dcFCCS) to detect and quantify condensates at the nanoscale, including their size, growth rate, molecular stoichiometry, and the binding affinity of client molecules within condensates. We expect that the quantitative dcFCCS method can be widely applied to investigate many other important phase separation systems.

Despite the importance of studying nucleoprotein complexes, no appropriate method for quantifying them is available. Here, a UV absorbance method using the formula "C_mg/mL = 1.55A₂₈₀ - 0.76A₂₆₀" were applied to quantify nucleoprotein complexes. After modification using two paired A₂₆₀ and A₂₈₀ values, the UV-derived formula-based method could accurately quantify proteins in nucleoprotein complexes. Otherwise, by taking the target protein as a standard, the Bradford assay can accurately quantify proteins in nucleoprotein complexes without interference by nucleic acids. The above methods were successfully applied to measure the concentration of MtuP49-CTG complexes of Mycobacterium tuberculosis. In conclusion, both the Bradford assay and the UV-derived formula-based method were appropriate for quantifying proteins in nucleoprotein complexes, which may make contributions to explore the interactions between proteins and nucleic acids at the molecular level.

With the biological relevance of the whole cells, low cost compared with animal experiments, a wide variety of cell-based screening platforms (cell-based assay, cell-based microfluidics, cell-based biosensor, cell-based chromatography) have been developed to address the challenges of drug discovery. In this review, we conclude the current advances in cell-based screening and summary the pros and cons of the platforms for different applications. Challenges and improvement strategies associated with cell-based methods are also discussed.

Cas9 is an RNA-guided endonuclease from the type II CRISPR-Cas system that employs RNA-DNA base pairing to target and cleave foreign DNA in bacteria. Due to its robust and programmable activity, Cas9 has been repurposed as a revolutionary technology for wide-ranging biological and medical applications. A comprehensive understanding of Cas9 mechanisms at the molecular level would aid in its better usage as a genome tool. Over the past few years, single-molecule techniques, such as fluorescence resonance energy transfer, DNA curtains, magnetic tweezers, and optical tweezers, have been extensively applied to characterize the detailed molecular mechanisms of Cas9 proteins. These techniques allow researchers to monitor molecular dynamics and conformational changes, probe essential DNA-protein interactions, detect intermediate states, and distinguish heterogeneity along the reaction pathway, thus providing enriched functional and mechanistic perspectives. This review outlines the single-molecule techniques that have been utilized for the investigation of Cas9 proteins and discusses insights into the mechanisms of the widely used Streptococcus pyogenes (Sp) Cas9 revealed through these techniques.

Life science is often focused on the microscopic level. Single-molecule technology has been used to observe components at the micro- or nanoscale. Single-molecule imaging provides unprecedented information about the behavior of individual molecules in contrast to the information from ensemble methods that average the information of many molecules in various states. A typical feature of living systems is motion. The lack of synchronicity of motion biomachines in living systems makes it challenging to image the motion process with high resolution. Thus, single-molecule technology is especially useful for real-time study on motion mechanism of biomachines, such as viral DNA packaging motor, or other ATPases. The most common optical instrumentations in single-molecule studies are optical tweezers and single molecule total internal refection fluorescence microscopy (smTIRF). Optical tweezers are the force-based technique. The analysis of RNA using optical tweezer has led to the discovery of the rubbery or amoeba property of RNA nanoparticles for compelling vessel extravasation to enhance tumor targeting and fast renal excretion. The rubbery property of RNA lends mechanistic evidence for RNAs use as an ideal reagent in cancer treatment with undetectable toxicity. Single molecule photobleaching allows for the direct counting of biomolecules. This technique was invented for single molecule counting of RNA in the phi29 DNA packaging motor to resolve the debate between five and six copies of RNA in the motor. The technology has subsequently extended to counting components in biological machines composed of protein, DNA, and other macromolecules. In combination with statistical analysis, it reveals biomolecular mechanisms in detail and leads to the development of ultra-sensitive sensors in diagnosis and forensics. This review focuses on the applications of optical tweezers and fluorescence-based techniques as single-molecule technologies to resolve mechanistic questions related to RNA and DNA nanostructures.

Tracking the transmembrane topology and conformational dynamics of membrane proteins is key to understand their functions. It is however challenging to monitor position changes of individual proteins in cell membranes with high sensitivity and high resolution. We review on three single-molecule fluorescence imaging methods - SIFA, LipoFRET and QueenFRET - recently developed in our lab for studying the dynamics of membrane proteins. They can be applied, progressively, to investigate membrane proteins in solid-supported lipid bilayers, artificial liposome membranes and live-cell plasma membranes. The techniques take advantage of the energy transfer from a fluorophore to a cloud of quenchers and are able to extract in real time positions and position changes of a single fluorophore-labeled protein in the direction normal to the membrane surface. The methods have sub-nanometer precision and have proved powerful to investigate biomolecules interacting with bio-membranes.

Single-molecule methods have been applied to study the mechanisms of many bio-physical systems that occur on the nanometer scale. To probe the dynamics of the such systems including vesicle docking, tethering, fusion, trafficking, protein-membrane interactions, etc., and to obtain reproducible experimental data; proper methodology and framework are crucial. Here, we address this need by developing a protocol for immobilization of vesicles composed of synthetic lipids and measurement using total internal reflection fluorescence (TIRF) microscopy. Furthermore, we demonstrate applications including vesicle clustering mediated by proteins such as alpha-Synuclein (αSyn) and the influence of external ions by using TIRF microscopy. Moreover, we use this method to quantify the dependence of lipid composition and charge on vesicle clustering mediated by αSyn which is based on the methods previously reported.