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Sphingolipids play important roles in physiology and cell biology, but a systematic examination of their functions is lacking. We performed a genome-wide CRISPRi screen in sphingolipid-depleted human cells and identified hypersensitive mutants in genes of membrane trafficking and lipid biosynthesis, including ether lipid synthesis. Systematic lipidomic analysis showed a coordinate regulation of ether lipids with sphingolipids, suggesting an adaptation and functional compensation. Biophysical experiments on model membranes show common properties of these structurally diverse lipids that also share a known function as glycosylphosphatidylinositol (GPI) anchors in different kingdoms of life. Molecular dynamics simulations show a selective enrichment of ether phosphatidylcholine around p24 proteins, which are receptors for the export of GPI-anchored proteins and have been shown to bind a specific sphingomyelin species. Our results support a model of convergent evolution of proteins and lipids, based on their physico-chemical properties, to regulate GPI- anchored protein transport and maintain homeostasis in the early secretory pathway.

The target of rapamycin complex 2 (TORC2) plays a key role in maintaining the homeostasis of plasma membrane (PM) tension. TORC2 activation following increased PM tension involves redistribution of the Slm1 and 2 paralogues from PM invaginations known as eisosomes into membrane compartments containing TORC2. How Slm1/2 relocalization is triggered, and if/how this plays a role in TORC2 inactivation with decreased PM tension, is unknown. Using osmotic shocks and palmitoylcarnitine as orthogonal tools to manipulate PM tension, we demonstrate that decreased PM tension triggers spontaneous, energy-independent reorganization of pre-existing phosphatidylinositol-4,5-bisphosphate into discrete invaginated membrane domains, which cluster and inactivate TORC2. These results demonstrate that increased and decreased membrane tension are sensed through different mechanisms, highlighting a role for membrane lipid phase separation in mechanotransduction.

Cells and organelles are delimited by lipid bilayers in which high deformability is essential to many cell processes, including motility, endocytosis and cell division. Membrane tension is therefore a major regulator of the cell processes that remodel membranes, albeit one that is very hard to measure in vivo. Here we show that a planarizable push-pull fluorescent probe called FliptR (fluorescent lipid tension reporter) can monitor changes in membrane tension by changing its fluorescence lifetime as a function of the twist between its fluorescent groups. The fluorescence lifetime depends linearly on membrane tension within cells, enabling an easy quantification of membrane tension by fluorescence lifetime imaging microscopy. We further show, using model membranes, that this linear dependency between lifetime of the probe and membrane tension relies on a membrane-tension-dependent lipid phase separation. We also provide calibration curves that enable accurate measurement of membrane tension using fluorescence lifetime imaging microscopy.

Membrane shape is a key feature of many cellular processes, including cell differentiation, division, migration, and trafficking. The development of nanostructured surfaces allowing for the in situ manipulation of membranes in living cells is crucial to understand these processes, but this requires complicated and limited-access technologies. Here, we investigate the self-organization of cellular membranes by using a customizable and bench top method allowing to engineer 1D SiO2 nanopillar arrays of defined sizes and shapes on high-performance glass compatible with advanced microscopies. As a result of this original combination, we provide a mapping of the morphology-induced modulation of the cell membrane mechanics, dynamics and steady-state organization of key protein complexes implicated in cellular trafficking and signal transduction.

Measuring forces inside cells is particularly challenging. With the development of quantitative microscopy, fluorophores which allow the measurement of forces became highly desirable. We have previously introduced a mechanosensitive flipper probe, which responds to the change of plasma membrane tension by changing its fluorescence lifetime and thus allows tension imaging by FLIM. Herein, we describe the design, synthesis, and evaluation of flipper probes that selectively label intracellular organelles, i.e., lysosomes, mitochondria, and the endoplasmic reticulum. The probes respond uniformly to osmotic shocks applied extracellularly, thus confirming sensitivity toward changes in membrane tension. At rest, different lifetimes found for different organelles relate to known differences in membrane organization rather than membrane tension and allow colabeling in the same cells. At the organelle scale, lifetime heterogeneity provides unprecedented insights on ER tubules and sheets, and nuclear membranes. Examples on endosomal trafficking or increase of tension at mitochondrial constriction sites outline the potential of intra-cellularly targeted fluorescent tension probes to address essential questions that were previously beyond reach.

The Target of Rapamycin Complex 2 (TORC2) plays a key role in maintaining the homeostasis of plasma membrane (PM) tension. TORC2 activation upon increased PM tension involves redistribution of the Slm1 and 2 paralogs from PM invaginations known as eisosomes into membrane compartments containing TORC2. How Slm1/2 relocalization is triggered, and if/how this plays a role in TORC2 inactivation upon decreased PM tension is unknown. Using osmotic shocks and Palmitoylcarnitine (PalmC) as orthogonal tools to manipulate PM tension, we demonstrate that decreased PM tension triggers spontaneous, energy-independent reorganization of pre-existing phosphatidylinositol 4,5-bisphosphate (PtdIns(4,5)P2) into discrete invaginated membrane domains which cluster and inactivate TORC2. These results demonstrate that an increase and a decrease in membrane tension are sensed through different mechanisms and highlight a role for membrane lipid phase separation in mechanotransduction.

International audience; Dynamin is a dimeric GTPase that assembles into a helix around the neck of endocytic buds. Upon GTP hydrolysis, dynamin breaks these necks, a reaction called membrane fission. Fission requires dynamin to first constrict the membrane. It is unclear, however, how dynamin helix constriction works. Here we undertake a direct high-speed atomic force microscopy imaging analysis to visualize the constriction of single dynamin-coated membrane tubules. We show GTP-induced dynamic rearrangements of the dynamin helix turns: the average distances between turns reduce with GTP hydrolysis. These distances vary, however, over time because helical turns were observed to transiently pair and dissociate. At fission sites, these cycles of association and dissociation were correlated with relative lateral displacement of the turns and constriction. Our findings show relative longitudinal and lateral displacements of helical turns related to constriction. Our work highlights the potential of high-speed atomic force microscopy for the observation of mechanochemical proteins onto membranes during action at almost molecular resolution.

In this report, “fluorescent flippers” are introduced to create planarizable push–pull probes with the mechanosensitivity and fluorescence lifetime needed for practical use in biology. Twisted push–pull scaffolds with large and bright dithienothiophenes and their S,S-dioxides as the first “fluorescent flippers” are shown to report on the lateral organization of lipid bilayers with quantum yields above 80% and lifetimes above 4 ns. Their planarization in liquid-ordered (Lo) and solid-ordered (So) membranes results in red shifts in excitation of up to +80 nm that can be transcribed into red shifts in emission of up to +140 nm by Förster resonance energy transfer (FRET). These unique properties are compatible with multidomain imaging in giant unilamellar vesicles (GUVs) and cells by confocal laser scanning or fluorescence lifetime imaging microscopy. Controls indicate that strong push–pull macrodipoles are important, operational probes do not relocate in response to lateral membrane reorganization, and two flippers are indeed needed to “really swim,” i.e., achieve high mechanosensitivity.

International audience; Keywords: High-speed atomic force microscopy High-speed force spectroscopy Membrane protein Membrane structure Titin Actin cortex a b s t r a c t Atomic force microscopy (AFM) is the type of scanning probe microscopy that is probably best adapted for imaging biological samples in physiological conditions with submolecular lateral and vertical resolution. In addition, AFM is a method of choice to study the mechanical unfolding of proteins or for cellular force spectroscopy. In spite of 28 years of successful use in biological sciences, AFM is far from enjoying the same popularity as electron and fluorescence microscopy. The advent of high-speed atomic force microscopy (HS-AFM), about 10 years ago, has provided unprecedented insights into the dynamics of membrane proteins and molecular machines from the single-molecule to the cellular level. HS-AFM imaging at nanometer-resolution and sub-second frame rate may open novel research fields depicting dynamic events at the single bio-molecule level. As such, HS-AFM is complementary to other structural and cellular biology techniques, and hopefully will gain acceptance from researchers from various fields. In this review we describe some of the most recent reports of dynamic bio-molecular imaging by HS-AFM, as well as the advent of high-speed force spectroscopy (HS-FS) for single protein unfolding.

International audience; High-speed atomic force microscopy is a powerful tool for studying structure and dynamics of proteins. So far, however, high-speed atomic force microscopy was restricted to well-controlled molecular systems of purified proteins. Here we integrate an optical microscopy path into high-speed atomic force microscopy, allowing bright field and fluorescence microscopy, without loss of high-speed atomic force microscopy performance. This hybrid high-speed atomic force microscopy/optical microscopy setup allows positioning of the high-speed atomic force microscopy tip with high spatial precision on an optically identified zone of interest on cells. We present movies at 960 ms per frame displaying aquaporin-0 array and single molecule dynamics in the plasma membrane of intact eye lens cells. This hybrid setup allows high-speed atomic force microscopy imaging on cells about 1,000 times faster than conventional atomic force microscopy/optical microscopy setups, and allows first time visualization of unlabelled membrane proteins on a eukaryotic cell under physiological conditions. This development advances high-speed atomic force microscopy from molecular to cell biology to analyse cellular processes at the membrane such as signalling, infection, transport and diffusion. H igh-speed atomic force microscopy (HS-AFM) 1 has been proven to be a unique and powerful tool for the concomitant analysis of the structure and dynamics of single biomolecules 2. HS-AFM was used to visualize myosin-V walking 3 , cellulase cellulose degradation 4 , F 1-ATPase rotary catalysis 5 , bacteriorhodopsin photocycle 6 , and OmpF diffusion and interaction 7. However, HS-AFM has been restricted so far to well-controlled molecular systems of purified proteins under well-controlled conditions. These systems were characterized by a limited number of pure molecular species with small corrugation, mainly because the fast z-piezo that follows the topography profile has a small extension range (typically 400 nm; ref. 8). Recently, the development of a wide-range HS-AFM scanner allowed the first nanoscale observation of molecular movements at two frames per second on living bacteria 9. Historically, AFM debuted as early as 1990 for cell imaging applications 10 , prompted by the need to perform cell structural analysis at a resolution superior to light microscopy. However, AFM images of cells revealed rather the cell interior, like actin filaments, than the membrane structure of the cells. Later, AFM and optical microscopy (OM) were combined in order to take advantage of OM's large-scale overview imaging capacities and its power to analyse fluorescence signal targets of proteins of interest 11. For this purpose, AFMs were built in a table-top configuration and mounted on inverted optical microscopes 12. In order to avoid modifications of the conventional inverted OM, two types of table-top AFM configurations were built. In one of them, the AFM tip and the scanner is combined into a single moving component, and in the other a large optical microscope sample stage on which the AFM sample is mounted needs to be moved. In both cases, there is a loss of AFM performance. In the tip-scanner configuration, the laser detection must be coupled to the moving cantilever, while for the second case, the sample stage that needs to be moved is complex and heavy. Both types of structures are prone to capture environmental noise and feature innate resonance frequencies. Furthermore, massive objects do not allow sub-millisecond mechanical response, thus precluding individual protein imaging at high resolution and high speed. In this work, we integrate an OM path into our HS-AFM, maintaining the structure of the HS-AFM setup 1 , and hence not compromising HS-AFM performance in terms of speed and resolution. To achieve this, we choose a completely different approach compared with most (if not all) AFM/OM integration developments: instead of building a table-top AFM mountable on an inverted optical microscope, accepting loss of AFM performance, we build an optical path into our HS-AFM setup, accepting minimal trade-offs in OM performance. We show that our setup can acquire bright field and fluorescence OM images of biological samples, and that it allows HS-AFM tip positioning with high precision guided by OM. Finally, we show that it achieves HS-AFM imaging of individual membrane proteins on eukaryotic cells and record their dynamics at an imaging rate of 960 ms per frame. This accomplishment opens the door to a wide range of real-time studies of molecular dynamics in membrane processes on living cells. Results Development of hybrid HS-AFM and fluorescence microscope. In order to integrate an OM path into the HS-AFM (Fig. 1 and Fig. 2a,b), we added and exchanged several elements to and from the HS-AFM setup, previously designed by Ando et al 1. The original laser diode of the HS-AFM system that reads the cantilever position was changed to a far-red superluminescent diode (SLD) with 750 nm wavelength and low coherence (Fig. 1, label 1). The use of the SLD 750 nm allowed the optical separation

International audience; Junctional microdomains, paradigm for membrane protein segregation in functional assemblies, in eye lens fiber cell membranes are constituted of lens-specific aquaporin-0 tetramers (AQP0(4)) and connexin (Cx) hexamers, termed connexons. Both proteins have double function to assure nutrition and mediate adhesion of lens cells. Here we use high-speed atomic force microscopy to examine microdomain protein dynamics at the single-molecule level. We found that the adhesion function of head-to-head associated AQP0(4) and Cx is cooperative. This finding provides first experimental evidence for the mechanistic importance for junctional microdomain formation. From the observation of lateral association-dissociation events of AQP0(4), we determine that the enthalpic energy gain of a single AQP0(4)-AQP0(4) interaction in the membrane plane is -2.7 k(B)T, sufficient to drive formation of microdomains. Connexon association is stronger as dynamics are rarely observed, explaining their rim localization in junctional microdomains.