Mechanism of membrane pore formation by human gasdermin-D.

Gasdermin-D (GSDMD), a member of the gasdermin protein family, mediates pyroptosis in human and murine cells. Cleaved by inflammatory caspases, GSDMD inserts its N-terminal domain (GSDMDNterm) into cellular membranes and assembles large oligomeric complexes permeabilizing the membrane. So far, the mechanisms of GSDMDNterm insertion, oligomerization, and pore formation are poorly understood. Here, we apply high-resolution (≤ 2 nm) atomic force microscopy (AFM) to describe how GSDMDNterm inserts and assembles in membranes. We observe GSDMDNterm inserting into a variety of lipid compositions, among which phosphatidylinositide (PI(4,5)P2) increases and cholesterol reduces insertion. Once inserted, GSDMDNterm assembles arc-, slit-, and ring-shaped oligomers, each of which being able to form transmembrane pores. This assembly and pore formation process is independent on whether GSDMD has been cleaved by caspase-1, caspase-4, or caspase-5. Using time-lapse AFM, we monitor how GSDMDNterm assembles into arc-shaped oligomers that can transform into larger slit-shaped and finally into stable ring-shaped oligomers. Our observations translate into a mechanistic model of GSDMDNterm transmembrane pore assembly, which is likely shared within the gasdermin protein family.

High-Resolution Imaging of Maltoporin LamB while Quantifying the Free-Energy Landscape and Asymmetry of Sugar Binding.

Maltoporins are a family of membrane proteins that facilitate the diffusion of hydrophilic molecules and maltosaccharides across the outer membrane of Gram-negative bacteria. Two contradicting models propose the sugar binding, uptake, and transport by maltoporins to be either symmetric or asymmetric. Here, we address this contradiction and introduce force-distance-based atomic force microscopy to image single maltoporin LamB trimers in the membrane at sub-nanometer resolution and simultaneously quantify the binding of different malto-oligosaccharides. We assay subtle differences of the binding free-energy landscape of maltotriose, maltotetraose, and maltopentaose, which quantifies how binding strength and affinity increase with the malto-oligosaccharide chain length. The ligand-binding parameters change considerably by mutating the extracellular loop 3, which folds into and constricts the transmembrane pore of LamB. By recording LamB topographs and structurally mapping binding events at sub-nanometer resolution, we observe LamB to preferentially bind maltodextrin from the periplasmic side, which shows sugar binding and uptake to be asymmetric. The study introduces atomic force microscopy as an analytical nanoscopic tool that can differentiate among the factors modulating and models describing the binding and uptake of substrates by membrane proteins.

Imaging G protein-coupled receptors while quantifying their ligand-binding free-energy landscape.

Imaging native membrane receptors and testing how they interact with ligands is of fundamental interest in the life sciences but has proven remarkably difficult to accomplish. Here, we introduce an approach that uses force-distance curve-based atomic force microscopy to simultaneously image single native G protein-coupled receptors in membranes and quantify their dynamic binding strength to native and synthetic ligands. We measured kinetic and thermodynamic parameters for individual protease-activated receptor-1 (PAR1) molecules in the absence and presence of antagonists, and these measurements enabled us to describe PAR1's ligand-binding free-energy landscape with high accuracy. Our nanoscopic method opens an avenue to directly image and characterize ligand binding of native membrane receptors.

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.

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 Ni(2+)-N-nitrilotriacetate groups, we locate specific interaction sites on SAS-6 at nanometer resolution and quantify the binding strength of the Ni(2+)-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.

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.

A robust CETSA data analysis automation workflow for routine screening.

The Cellular Thermal Shift Assay (CETSA) enables the study of protein-ligand interactions in a cellular context. It provides valuable information on the binding affinity and specificity of both small and large molecule ligands in a relevant physiological context, hence forming a unique tool in drug discovery. Though high-throughput lab protocols exist for scaling up CETSA, subsequent data analysis and quality control remain laborious and limit experimental throughput. Here, we introduce a scalable and robust data analysis workflow which allows integration of CETSA into routine high throughput screening (HT-CETSA). This new workflow automates data analysis and incorporates quality control (QC), including outlier detection, sample and plate QC, and result triage. We describe the workflow and show its robustness against typical experimental artifacts, show scaling effects, and discuss the impact of data analysis automation by eliminating manual data processing steps.

Structural Properties of the Human Protease-Activated Receptor 1 Changing by a Strong Antagonist.

The protease-activated receptor 1 (PAR1), a G protein-coupled receptor (GPCR) involved in hemostasis, thrombosis, and inflammation, is activated by thrombin or other coagulation proteases. This activation is inhibited by the irreversible antagonist vorapaxar used for anti-platelet therapy. Despite detailed structural and functional information, how vorapaxar binding alters the structural properties of PAR1 to prevent activation is hardly known. Here we apply dynamic single-molecule force spectroscopy to characterize how vorapaxar binding changes the mechanical, kinetic, and energetic properties of human PAR1 under physiologically relevant conditions. We detect structural segments stabilizing PAR1 and quantify their properties in the unliganded and the vorapaxar-bound state. In the presence of vorapaxar, most structural segments increase conformational variability, lifetime, and free energy, and reduce mechanical rigidity. These changes highlight a general trend in how GPCRs are affected by strong antagonists.

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 Halobacterium salinarum, 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.

SAS-6 engineering reveals interdependence between cartwheel and microtubules in determining centriole architecture.

Centrioles are critical for the formation of centrosomes, cilia and flagella in eukaryotes. They are thought to assemble around a nine-fold symmetric cartwheel structure established by SAS-6 proteins. Here, we have engineered Chlamydomonas reinhardtii SAS-6-based oligomers with symmetries ranging from five- to ten-fold. Expression of a SAS-6 mutant that forms six-fold symmetric cartwheel structures in vitro resulted in cartwheels and centrioles with eight- or nine-fold symmetries in vivo. In combination with Bld10 mutants that weaken cartwheel-microtubule interactions, this SAS-6 mutant produced six- to eight-fold symmetric cartwheels. Concurrently, the microtubule wall maintained eight- and nine-fold symmetries. Expressing SAS-6 with analogous mutations in human cells resulted in nine-fold symmetric centrioles that exhibited impaired length and organization. Together, our data suggest that the self-assembly properties of SAS-6 instruct cartwheel symmetry, and lead us to propose a model in which the cartwheel and the microtubule wall assemble in an interdependent manner to establish the native architecture of centrioles.

Identifying and quantifying two ligand-binding sites while imaging native human membrane receptors by AFM.

A current challenge in life sciences is to image cell membrane receptors while characterizing their specific interactions with various ligands. Addressing this issue has been hampered by the lack of suitable nanoscopic methods. Here we address this challenge and introduce multifunctional high-resolution atomic force microscopy (AFM) to image human protease-activated receptors (PAR1) in the functionally important lipid membrane and to simultaneously localize and quantify their binding to two different ligands. Therefore, we introduce the surface chemistry to bifunctionalize AFM tips with the native receptor-activating peptide and a tris-N-nitrilotriacetic acid (tris-NTA) group binding to a His10-tag engineered to PAR1. We further introduce ways to discern between the binding of both ligands to different receptor sites while imaging native PAR1s. Surface chemistry and nanoscopic method are applicable to a range of biological systems in vitro and in vivo and to concurrently detect and localize multiple ligand-binding sites at single receptor resolution.

Organization of the Escherichia coli aerobic enzyme complexes of oxidative phosphorylation in dynamic domains within the cytoplasmic membrane.

The Escherichia coli cytoplasmic membrane contains the enzyme complexes of oxidative phosphorylation (OXPHOS). Not much is known about their supramolecular organization and their dynamics within the membrane in this model organism. In mitochondria and other bacteria, it was demonstrated by nondenaturing electrophoretic methods and electron microscopy that the OXPHOS complexes are organized in so-called supercomplexes, stable assemblies with a defined number of the individual enzyme complexes. To investigate the organization of the E. coli enzyme complexes of aerobic OXPHOS in vivo, we established fluorescent protein fusions of the NADH:ubiquinone oxidoreductase, the succinate:ubiquinone oxidoreductase, the cytochrome bd-I, and the cytochrome bo3 terminal oxidases, and the FoF1 ATP-synthase. The fusions were integrated in the chromosome to prevent artifacts caused by protein overproduction. Biochemical analysis revealed that all modified complexes were fully assembled, active, and stable. The distribution of the OXPHOS complexes in living cells was determined using total internal reflection fluorescence microscopy. The dynamics within the membrane were detected by fluorescence recovery after photobleaching. All aerobic OXPHOS complexes showed an uneven distribution in large mobile patches within the E. coli cytoplasmic membrane. It is discussed whether the individual OXPHOS complexes are organized as clustered individual complexes, here called "segrazones."

Multiparametric high-resolution imaging of native proteins by force-distance curve-based AFM.

A current challenge in the life sciences is to understand how the properties of individual molecular machines adjust in order to meet the functional requirements of the cell. Recent developments in force-distance (FD) curve-based atomic force microscopy (FD-based AFM) enable researchers to combine sub-nanometer imaging with quantitative mapping of physical, chemical and biological properties. Here we present a protocol to apply FD-based AFM to the multiparametric imaging of native proteins under physiological conditions. We describe procedures for experimental FD-based AFM setup, high-resolution imaging of proteins in the native unperturbed state with simultaneous quantitative mapping of multiple parameters, and data interpretation and analysis. The protocol, which can be completed in 1-3 d, enables researchers to image proteins and protein complexes in the native unperturbed state and to simultaneously map their biophysical and biochemical properties at sub-nanometer resolution.

Out but not in: the large transmembrane ß-barrel protein FhuA unfolds but cannot refold via ß-hairpins.

How transmembrane ß-barrel proteins insert and fold into membranes and by which factors they destabilize, unfold, and misfold represents a field of intense studies. Here, we use single-molecule force spectroscopy to characterize the un- and refolding of the ferric hydroxamate uptake receptor (FhuA), which is one of the largest ß-barrel proteins of the outer membrane of Escherichia coli. Applied to mechanical stress, FhuA undergoes a complex unfolding pathway in which each of the 11 ß-hairpins unfolds one after the other until the entire ß-barrel has unfolded. Once unfolded and relaxed, the FhuA polypeptide cannot fold back into the lipid membrane and adopts various misfolded conformations. Such misfolding is in contrast to the reversible refolding behavior of much smaller ß-barrel outer membrane proteins OmpA and OmpG that occurs at similar experimental conditions. The results suggest that large ß-barrel proteins that show more complex (un-)folding pathways require cofactors for proper insertion and folding into the membrane.