Mechanical Stimulation of Piezo1 Receptors Depends on Extracellular Matrix Proteins and Directionality of Force

Piezo receptors convert mechanical forces into electrical signals. In mammals, they play important roles in basic physiological functions including proprioception, sensation of touch, and vascular development. However, basic receptor properties like the gating mechanism, the interaction with extracellular matrix (ECM) proteins, and the response to mechanical stimulation, remain poorly understood. Here, we establish an atomic force microscopy (AFM)-based assay to mechanically stimulate Piezo1 receptors in living animal cells, while monitoring receptor activation in real-time using functional calcium imaging. Our experiments show that in the absence of ECM proteins Piezo1 receptors are relatively insensitive to mechanical forces pushing the cellular membrane, whereas they can hardly be activated by mechanically pulling the membrane. Yet, if conjugated with Matrigel, a mix of ECM proteins, the receptors become sensitized. Thereby, forces pulling the cellular membrane activate the receptor much more efficiently compared to pushing forces. Finally, we found that collagen IV, a component of the basal lamina, which forms a cohesive network and mechanical connection between cells, sensitizes Piezo1 receptors to mechanical pulling

Nanomechanical Properties of Proteins and Membranes Depend on Loading Rate and Electrostatic Interactions

Knowing the dynamic mechanical response of tissue, cells, membranes, proteins, nucleic acids, and carbohydrates to external perturbations is important to understand various biological and biotechnological problems. Atomic force microscopy (AFM)-based approaches are the most frequently used nanotechnologies to determine the mechanical properties of biological samples that range in size from microscopic to (sub)nanoscopic. However, the dynamic nature of biomechanical properties has barely been addressed by AFM imaging. In this work, we characterizethe viscoelastic properties of the native light-driven proton pump bacteriorhodopsin of the purple membrane of <i>Halobacterium salinarum</i>. Using force–distance curve (<i>F</i>–<i>D</i>)-based AFM we imaged purple membranes while force probing their mechanical response over a wide range of loading rates (from ∼0.5 to 100 μN/s). Our results show that the mechanical stiffness of protein and membrane increases with the loading rate up to a factor of 10 (from ∼0.3 to 3.2 N/m). In addition, the electrostatic repulsion between AFM tip and sample can alter the mechanical stiffness measured by AFM up to ∼60% (from ∼0.8 to 1.3 N/m).These findings indicate that the mechanical response of membranes and proteins and probably of other biomolecular systems should be determined at different loading rates to fully understand their properties

Quantitative Imaging of the Electrostatic Field and Potential Generated by a Transmembrane Protein Pore at Subnanometer Resolution

Elucidating the mechanisms by which proteins translocate small molecules and ions through transmembrane pores and channels is of great interest in biology, medicine, and nanotechnology. However, the characterization of pore forming proteins in their native state lacks suitable methods that are capable of high-resolution imaging (∼1 nm) while simultaneously mapping physical and chemical properties. Here we report how force–distance (FD) curve-based atomic force microscopy (AFM) imaging can be applied to image the native pore forming outer membrane protein F (OmpF) at subnanometer resolution and to quantify the electrostatic field and potential generated by the transmembrane pore. We further observe the electrostatic field and potential of the OmpF pore switching “on” and “off” in dependence of the electrolyte concentration. Because electrostatic field and potential select for charged molecules and ions and guide them to the transmembrane pore the insights are of fundamental importance to understand the pore function. These experimental results establish FD-based AFM as a unique tool to image biological systems to subnanometer resolution and to quantify their electrostatic properties

Pull-and-Paste of Single Transmembrane Proteins

How complex cytoplasmic membrane proteins insert and fold into cellular membranes is not fully understood. One problem is the lack of suitable approaches that allow investigating the process by which polypeptides insert and fold into membranes. Here, we introduce a method to mechanically unfold and extract a single polytopic α-helical membrane protein, the lactose permease (LacY), from a phospholipid membrane, transport the fully unfolded polypeptide to another membrane and insert and refold the polypeptide into the native structure. Insertion and refolding of LacY is facilitated by the transmembrane chaperone/insertase YidC in the absence of the SecYEG translocon. Insertion into the membrane occurs in a stepwise, stochastic manner employing multiple coexisting pathways to complete the folding process. We anticipate that our approach will provide new means of studying the insertion and folding of membrane proteins and to mechanically reconstitute membrane proteins at high spatial precision and stoichiometric control, thus allowing the functional programming of synthetic and biological membranes

Detecting Ligand-Binding Events and Free Energy Landscape while Imaging Membrane Receptors at Subnanometer Resolution

Force–distance curve-based atomic force microscopy has emerged into a sophisticated technique for imaging cellular membranes and for detecting specific ligand-binding events of native membrane receptors. However, so far the resolution achieved has been insufficient to structurally map ligand-binding sites onto membrane proteins. Here, we introduce experimental and theoretical approaches for overcoming this limitation. To establish a structurally and functionally well-defined reference sample, we engineer a ligand-binding site to the light-driven proton pump bacteriorhodopsin of purple membrane. Functionalizing the AFM stylus with an appropriate linker-system tethering the ligand and optimizing the AFM conditions allows for imaging the engineered bacteriorhodopsin at subnanometer resolution while structurally mapping the specific ligand–receptor binding events. Improved data analysis allows reconstructing the ligand-binding free energy landscape from the experimental data, thus providing thermodynamic and kinetic insight into the ligand-binding process. The nanoscopic method introduced is generally applicable for imaging receptors in native membranes at subnanometer resolution and for systematically mapping and quantifying the free energy landscape of ligand binding

Unraveling the Pore-Forming Steps of Pneumolysin from <i>Streptococcus pneumoniae</i>

Pneumolysin (PLY) is the main virulence factor of <i>Streptococcus pneumoniae</i> that causes pneumonia, meningitis, and invasive pneumococcal infection. PLY is produced as monomers, which bind to cholesterol-containing membranes, where they oligomerize into large pores. To investigate the pore-forming mechanism, we determined the crystal structure of PLY at 2.4 Å and used it to design mutants on the surface of monomers. Electron microscopy of liposomes incubated with PLY mutants revealed that several mutations interfered with ring formation. Mutants that formed incomplete rings or linear arrays had strongly reduced hemolytic activity. By high-resolution time-lapse atomic force microscopy of wild-type PLY, we observed two different ring-shaped complexes. Most of the complexes protruded ∼8 nm above the membrane surface, while a smaller number protruded ∼11 nm or more. The lower complexes were identified as pores or prepores by the presence or absence of a lipid bilayer in their center. The taller complexes were side-by-side assemblies of monomers of soluble PLY that represent an early form of the prepore. Our observations suggest a four-step mechanism of membrane attachment and pore formation by PLY, which is discussed in the context of recent structural models. The functional separation of these steps is necessary for the understanding how cholesterol-dependent cytolysins form pores and lyse cells

Localizing Chemical Groups while Imaging Single Native Proteins by High-Resolution Atomic Force Microscopy

Simultaneous high-resolution imaging and localization of chemical interaction sites on single native proteins is a pertinent biophysical, biochemical, and nanotechnological challenge. Such structural mapping and characterization of binding sites is of importance in understanding how proteins interact with their environment and in manipulating such interactions in a plethora of biotechnological applications. Thus far, this challenge remains to be tackled. Here, we introduce force–distance curve-based atomic force microscopy (FD-based AFM) for the high-resolution imaging of SAS-6, a protein that self-assembles into cartwheel-like structures. Using functionalized AFM tips bearing Ni²⁺-<i>N</i>-nitrilotriacetate groups, we locate specific interaction sites on SAS-6 at nanometer resolution and quantify the binding strength of the Ni²⁺-NTA groups to histidine residues. The FD-based AFM approach can readily be applied to image any other native protein and to locate and structurally map histidine residues. Moreover, the surface chemistry used to functionalize the AFM tip can be modified to map other chemical interaction sites

Directly Observing the Lipid-Dependent Self-Assembly and Pore-Forming Mechanism of the Cytolytic Toxin Listeriolysin O

Listeriolysin O (LLO) is the major virulence factor of <i>Listeria monocytogenes</i> and a member of the cholesterol-dependent cytolysin (CDC) family. Gram-positive pathogenic bacteria produce water-soluble CDC monomers that bind cholesterol-dependent to the lipid membrane of the attacked cell or of the phagosome, oligomerize into prepores, and insert into the membrane to form transmembrane pores. However, the mechanisms guiding LLO toward pore formation are poorly understood. Using electron microscopy and time-lapse atomic force microscopy, we show that wild-type LLO binds to membranes, depending on the presence of cholesterol and other lipids. LLO oligomerizes into arc- or slit-shaped assemblies, which merge into complete rings. All three oligomeric assemblies can form transmembrane pores, and their efficiency to form pores depends on the cholesterol and the phospholipid composition of the membrane. Furthermore, the dynamic fusion of arcs, slits, and rings into larger rings and their formation of transmembrane pores does not involve a height difference between prepore and pore. Our results reveal new insights into the pore-forming mechanism and introduce a dynamic model of pore formation by LLO and other CDC pore-forming toxins

High-Resolution Imaging and Multiparametric Characterization of Native Membranes by Combining Confocal Microscopy and an Atomic Force Microscopy-Based Toolbox

To understand how membrane proteins function requires characterizing their structure, assembly, and inter- and intramolecular interactions in physiologically relevant conditions. Conventionally, such multiparametric insight is revealed by applying different biophysical methods. Here we introduce the combination of confocal microscopy, force–distance curve-based (FD-based) atomic force microscopy (AFM), and single-molecule force spectroscopy (SMFS) for the identification of native membranes and the subsequent multiparametric analysis of their membrane proteins. As a well-studied model system, we use native purple membrane from <i>Halobacterium salinarum</i>, whose membrane protein bacteriorhodopsin was His-tagged to bind nitrilotriacetate (NTA) ligands. First, by confocal microscopy we localize the extracellular and cytoplasmic surfaces of purple membrane. Then, we apply AFM to image single bacteriorhodopsins approaching sub-nanometer resolution. Afterwards, the binding of NTA ligands to bacteriorhodopsins is localized and quantified by FD-based AFM. Finally, we apply AFM-based SMFS to characterize the (un)­folding of the membrane protein and to structurally map inter- and intramolecular interactions. The multimethodological approach is generally applicable to characterize biological membranes and membrane proteins at physiologically relevant conditions

Single-Molecule Force Spectroscopy from Nanodiscs: An Assay to Quantify Folding, Stability, and Interactions of Native Membrane Proteins

Single-molecule force spectroscopy (SMFS) can quantify and localize inter- and intramolecular interactions that determine the folding, stability, and functional state of membrane proteins. To conduct SMFS the membranes embedding the membrane proteins must be imaged and localized in a rather time-consuming manner. Toward simplifying the investigation of membrane proteins by SMFS, we reconstituted the light-driven proton pump bacteriorhodopsin into lipid nanodiscs. The advantage of using nanodiscs is that membrane proteins can be handled like water-soluble proteins and characterized with similar ease. SMFS characterization of bacteriorhodopsin in native purple membranes and in nanodiscs reveals no significant alterations of structure, function, unfolding intermediates, and strengths of inter- and intramolecular interactions. This demonstrates that lipid nanodiscs provide a unique approach for <i>in vitro</i> studies of native membrane proteins using SMFS and open an avenue to characterize membrane proteins by a wide variety of SMFS approaches that have been established on water-soluble proteins