The authors thank Libby Perry (Augusta University) for her help with the SEM sample preparation. This work was supported by the National Institutes of Health [R01HL139562 (G.C.) and K99HL146954 (B.S.)] and American Heart Association [17POST33661254 (B.S.)].

NOTE: The following is a general protocol used to quantify membrane ruffle formation in RAW 264.7 macrophages using SEM microscopy. Optimization may be required for different cell types.
1. Cell line and cell culture
<ol>
	<li>Grow RAW 264.7 macrophages in Dulbecco&#39;s Modified Eagle Medium (DMEM) supplemented with 10% (vol/vol) heat-inactivated Fetal Bovine Serum (FBS), 100 IU/mL of penicillin and 100 &#181;g/mL streptomycin in a humidified incubator at 37 &#176;C and 5% CO2. Replace growth media every other day.</li>
	<li>When cells become 80% confluent, wash the plate two times using sterile PBS.</li>
	<li>Detach cells by adding 1,000 &#181;L of 0.5% trypsin-EDTA and incubate the cell culture plate for 3-5 min in a humidified incubator at 37 &#176;C.</li>
	<li>Examine the plate under a light microscope to confirm cell dissociation. Add 10 mL of growth media containing 10% FBS to inactivate trypsin.</li>
	<li>Collect the cell suspension in a 50 mL conical tube and centrifuge at 400 x g for 5 min at room temperature. Gently resuspend cells in the complete cell culture medium and determine the cell count and viability using Trypan Blue (0.4%) staining. 
	NOTE: The recommended minimum cell viability for this experiment is &#62;90%.</li>
	<li>Place sterile glass coverslips in wells of a 24-well plate using autoclaved forceps. Seed cells onto coverslips at a density of 1 x 106 cells/mL and incubate the plate overnight in a humidified incubator at 37 &#176;C and 5% CO2.</li>
	<li>Change media of each well with fresh 500 &#181;L complete media before treatment. Pretreat macrophages with vehicle (DMSO or other solvents used to dissolve EIPA) or the macropinocytosis inhibitor 5-(N-ethyl-N-isopropyl)-Amiloride (EIPA, 25 &#181;M21) for 30 min.</li>
	<li>Treat cells with vehicle and stimulators of macropinocytosis to promote membrane ruffling: phorbol 12-myristate 13-acetate (PMA, 1 &#181;M, 30 min21) and macrophage colony-stimulating factor (M-CSF, 100 ng/mL, 30 min3). 
	NOTE: Alternative inhibitors [e.g., latrunculin A (1 &#181;M, 30 min) or cytochalasin D (1 &#181;M, 30 min)] and stimulators [e.g., epidermal growth factor (EGF, 100 ng/mL, 10 min) or platelet-derived growth factor (PDGF, 100 ng/mL, 15 min)] of macropinocytosis can be also used to characterize macropinocytosis in macrophages and other cell types16,27,28,29. Cells may require overnight serum starvation prior to the treatment with physiological stimulators of macropinocytosis. It is important to note that the most effective concentrations and incubation times will have to be determined to stimulate macropinocytosis in other cell types.</li>
</ol>
2. SEM fixation
<ol>
	<li>After the treatment, aspirate the media from wells and wash coverslips with ice-cold PBS twice.</li>
	<li>Fix cells (4% paraformaldehyde and 2% glutaraldehyde in 0.1 M sodium cacodylate) for 30 min at room temperature, followed by overnight incubation in the fixative at 4 &#176;C.</li>
	<li>Gently wash and incubate coverslips with 500 &#181;L of following reagents without disturbing the cell monolayer.
	<ol>
		<li>Wash once in 0.1 M sodium cacodylate and incubate for 15 min.</li>
		<li>Wash twice with dH2O with 10 min incubation in each wash.</li>
		<li>Wash twice with 25% ethanol with 10 min incubation in each wash.</li>
		<li>Wash twice with 50% ethanol with 10 min incubation in each wash.</li>
		<li>Wash twice with 75% ethanol with 10 min incubation in each wash.</li>
		<li>Wash twice with 80% ethanol with 10 min incubation in each wash.</li>
		<li>Wash twice with 95% ethanol with 10 min incubation in each wash.</li>
		<li>Wash twice with 100% ethanol. Perform 10 min of incubation in each wash. 
		NOTE: Changing/replacing of reagents should be done fast to prevent air drying of cells. Coverslips can be left in 100% ethanol for several days at 4 &#176;C. For long-term storage, add additional 100% ethanol and cover wells with parafilm to prevent air drying of the sample.</li>
	</ol>
	</li>
</ol>
3.&#160;Critical point drying
<ol>
	<li>Place coverslips in a Critical Point Dryer and cover with 100% ethanol. Press the Power button and open the CO2 tank.</li>
	<li>Press the Cool button for approximately 30 s until the temperature decreases to 0 &#176;C. Press the Fill button until a bubble appears in the chamber window.</li>
	<li>Press the Purge button until the smell of ethanol from the purge exhaust disappears. Press the Cool button again until the temperature decreases to 0 &#176;C.</li>
	<li>Repress the Fill and Purge buttons to turn them off and close the CO2 tank. Repress the Cool button to turn it off and press the Heat button.</li>
	<li>Set the temperature at 42 &#176;C and pressure at 1,200 psi. Once the pressure and temperature stabilize, press the Bleed button to allow the pressure to decrease slowly.</li>
	<li>Once the chamber pressure reaches 150 psi, press the Vent button and wait until the pressure decreases to 0 psi. Turn off the critical point dryer and remove the coverslips.</li>
	<li>Mount coverslips on SEM aluminum specimen mounts using Carbon Adhesive tabs and subject to sputter coating using gold/palladium in a sputter coater.</li>
	<li>Turn on the Power button and wait until the vacuum reaches 30 mTorr. Flush the chamber to remove humidity and air by turning off the gas switch and turning the Fine gas valve counterclockwise.</li>
	<li>Once the vacuum increases to 200 mTorr turn off the gas switch and wait until the vacuum reaches 30 mTorr. Repeat this step 3 times.</li>
	<li>Push the Timer button and adjust the Voltage knob until the gauge reads 10 mA. Remove coated coverslips from the chamber. 
	NOTE: Follow manufacturer&#39;s instruction to perform critical point drying and sputter coating of the sample.</li>
</ol>
4. Imaging and quantification
<ol>
	<li>Insert sample coverslips into the chamber of a scanning electron microscope. Close the door and press the Evac button.</li>
	<li>Open the SEM operation software. Set the accelerating voltage to 15 kv and the working distance to 10 mm.</li>
	<li>Press the Coordinates button and move around the controller until the cells appear in the center of the observation screen. Set the magnification to 3,500x and image the sample by clicking the Photo button. 
	NOTE: Use a higher level of magnification (8,500x to 16,000x) to show the plasma membrane at greater details.</li>
	<li>Take images from at least 10 random locations on the coverslips, making sure that each microscopic field contains multiple cells. Save images as .tif files.</li>
	<li>From the images, locate membrane ruffles, defined as protruding blanket-like folds of the plasma membrane, with a size greater than 500 nm (Figure 1). Count the number of ruffles per cell. 
	NOTE: C-shaped ruffles were identified as curved membrane protrusions that did not fuse on their lateral ends [(Figure 1 (M-CSF treatment) and Figure 2B]. Fused circular membrane ruffles, prior to their closure, were identified as macropinocytotic cups (Figure 2C).</li>
	<li>Normalize the number of membrane ruffles to the total number of cells in the microscopic field evaluated. Repeat this analysis for the samples from each group.</li>
</ol>

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The authors declare no conflicts of interest.

The present SEM imaging protocol provides a tool to visualize and quantify membrane ruffle formation, circular protrusions, and macropinocytic cups on the cell surface in vitro. Although the current protocol focuses on macrophages, studies have shown that membrane ruffle formation also occurs on various other cell types including dendritic cells, fibroblasts, neurons, and cancer cells11,12,14,<sup class='xref...

Macropinocytosis is an endocytic process responsible for internalizing a large amount of extracellular fluid and its content via the formation of dynamic and actin-driven plasma membrane protrusions called membrane ruffles1. Many of these membrane ruffles form cups that close and fuse back onto the cell and separate from the plasma membrane as large, heterogeneous intracellular endosomes also known as macropinosomes1. Although macropinocytosis is induced by growth factors such as macrophage colony-stimulating factor (M-CSF) and epidermal growth factor (EGF) in a wide range of cell types, an additional unique, calcium-dependent process known as constitutive macropinocytosis has been also observed in innate immune cells2,3,4,5,6,7,8.
The ability of cells to internalize extracellular material via macropinocytosis has been shown to play an important role in a variety of physiological processes ranging from nutrient uptake to pathogen capture and antigen presentation9,10,11. However, because this process is non-selective and inducible, it has also been implicated in a number of pathological conditions. Indeed, previous studies suggested that macropinocytosis plays an important role in Alzheimer&#39;s disease, Parkinson&#39;s disease, cancer, nephrolithiasis and atherosclerosis12,13,14,15,16. Moreover, certain bacteria and viruses have shown to utilize macropinocytosis to gain entry into host cells and induce infection17,18. Interestingly, stimulation of macropinocytosis can be also exploited for targeted delivery of therapeutic agents in various disease conditions19,20.
Previous studies have explored macropinocytosis by quantifying internalized fluorescently-tagged fluid-phase markers in the absence and presence of pharmacological agents that inhibit macropinocytosis using flow cytometry and confocal imaging21,22. Currently available pharmacological tools that inhibit macropinocytosis are limited to and comprise of 1) actin polymerization inhibitors (cytochalasin D and latrunculins), 2) PI3K blockers (LY-290042 and wortmannin) and 3) inhibitors of sodium hydrogen exchangers (NHE) (amiloride and EIPA)5,14,15,23,24,25. However, because these inhibitors have endocytosis independent effects, it is difficult to selectively determine the contribution of macropinocytosis to solute uptake and disease pathogenesis especially in vivo21.
Scanning electron microscopy (SEM) is a type of electron microscope that produces ultra-high-resolution images of cells using a focused beam of electrons26. In macropinocytosis research, SEM imaging is regarded as the gold standard technique to visualize topographical and morphological characteristics of the plasma membrane, quantify membrane ruffle formation, and investigate their progression towards macropinosome internalization. Furthermore, scanning electron microscopy combined with the quantification of solute uptake, in the presence and absence of macropinocytosis blockers, provides a reliable strategy to examine macropinocytotic solute internalization in vitro. This paper provides a detailed protocol on how to prepare cells for SEM, visualize the cell surface, quantify ruffle formation, and examine their progress towards cup closure and macropinosome internalization.

<table><tbody><tr><td>0.5% Trypsin-EDTA</td><td>Gibco</td><td>15400-054</td><td></td></tr><tr><td>2% Glutaraldehyde</td><td>Electron Microscopy Sciences</td><td>16320</td><td></td></tr><tr><td>4% Paraformaldehyde</td><td>Santa Cruz Biotechnology</td><td>281692</td><td></td></tr><tr><td>5-(N-ethyl-N-isopropyl)-Amiloride</td><td>Sigma Life Science</td><td>A3085</td><td></td></tr><tr><td>Accuri C6 Flow Cytometer</td><td></td><td></td><td></td></tr><tr><td>Carbon Adhesive Tabs</td><td>Electron Microscopy Sciences</td><td>77825-09</td><td></td></tr><tr><td>Dimethyl Sulfoxide</td><td>Corning</td><td>25-950-CQC</td><td></td></tr><tr><td>Dulbecco&#039;s Modified Eagle Medium</td><td>Cytiva Life Sciences</td><td>SH30022.01</td><td></td></tr><tr><td>Falcon 24-well Clear Flat Bottom TC-treated Multiwell Cell Culture Plate</td><td>Falcon</td><td>353047</td><td></td></tr><tr><td>Fetal Bovine Serum</td><td>Gemini Bio</td><td>900-108</td><td></td></tr><tr><td>FitC-dextran</td><td>Thermo Fisher Scientific</td><td>D1823</td><td></td></tr><tr><td>FM 4-64</td><td>Thermo Fisher Scientific</td><td>T13320</td><td></td></tr><tr><td>HERAcell 150i CO2 incubator</td><td>Thermo Fisher Scientific</td><td>51026282</td><td></td></tr><tr><td>Hummer Model 6.2 Sputter Coater</td><td>Anatech USA</td><td>58565</td><td></td></tr><tr><td>JSM-IT500HR scanning electron microscope</td><td></td><td></td><td></td></tr><tr><td>Microscope Cover Glass</td><td>Thermo Fisher Scientific</td><td>12-545-82</td><td></td></tr><tr><td>Pen Strep</td><td>Gibco</td><td>15140-122</td><td></td></tr><tr><td>phorbol 12-myristate 13-acetate</td><td>Millipore Sigma</td><td>524400</td><td></td></tr><tr><td>RAW 264.7 macrophage</td><td>ATCC</td><td>ATCC TIB-71</td><td></td></tr><tr><td>Recombinant Human M-CSF</td><td>Peprotech</td><td>300-25</td><td></td></tr><tr><td>Samdri-790 Critical Point Dryer</td><td>Tousimis Research Corporation</td><td>8778B</td><td></td></tr><tr><td>SEM Aluminum Specimen Mounts</td><td>Electron Microscopy Sciences</td><td>75220</td><td></td></tr><tr><td>Sodium Cacodylate</td><td>Electron Microscopy Sciences</td><td>12300</td><td></td></tr><tr><td>Texas red-dextra</td><td>Thermo Fisher Scientific</td><td>D1864</td><td></td></tr><tr><td>Trypan Blue Solution</td><td>Thermo Fisher Scientific</td><td>15250061</td><td></td></tr><tr><td>Zeiss LSM 780 confocal microscope</td><td></td><td></td><td></td></tr></tbody></table>

visualizing membrane ruffle formation using scanning electron microscopy

Membrane ruffling is the formation of motile plasma membrane protrusions containing a meshwork of newly polymerized actin filaments. Membrane ruffles may form spontaneously or in response to growth factors, inflammatory cytokines, and phorbol esters. Some of the membrane protrusions may reorganize into circular membrane ruffles that fuse at their distal margins and form cups that close and separate into the cytoplasm as large, heterogeneous vacuoles called macropinosomes. During the process, ruffles trap extracellular fluid and solutes that internalize within macropinosomes. High-resolution scanning electron microscopy (SEM) is a commonly used imaging technique to visualize and quantify membrane ruffle formation, circular protrusions, and closed macropinocytic cups on the cell surface. The following protocol describes the cell culture conditions, stimulation of the membrane ruffle formation in vitro, and how to fix, dehydrate, and prepare cells for imaging using SEM. Quantification of membrane ruffling, data normalization, and stimulators and inhibitors of membrane ruffle formation are also described. This method can help answer key questions about the role of macropinocytosis in physiological and pathological processes, investigate new targets that regulate membrane ruffle formation, and identify yet uncharacterized physiological stimulators as well as novel pharmacological inhibitors of macropinocytosis.

Here, we describe the results from the presented technique. Representative SEM images shown in Figure 1 demonstrate membrane ruffle formation in RAW 264.7 macrophages following treatment with PMA and M-CSF. Images were first captured at a magnification of 3,500x for quantification purposes and then at higher levels of magnification (8,500x to 16,000x) to show the plasma membrane at greater details. Pretreatment of macrophages with the macropinocytosis inhibitor EIPA attenuated membrane ruffl...

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Visualizing Membrane Ruffle Formation using Scanning Electron Microscopy

Macropinocytosis is a highly conserved endocytic process initiated by the formation of F-actin-rich sheet-like membrane projections, also known as membrane ruffles. Increased rate of macropinocytotic solute internalization has been implicated in various pathological conditions. This protocol presents a method to quantify membrane ruffle formation in vitro using scanning electron microscopy.

Membrane ruffling is the formation of motile plasma membrane protrusions containing a meshwork of newly polymerized actin filaments. Membrane ruffles may ...

visualizing-membrane-ruffle-formation-using-scanning-electron

Research

JoVE Journal

Medicine

2.5K Views.  Medical College of Georgia at Augusta University. Het huidige SEM-beeldvormingsprotocol biedt een hulpmiddel om membraanrommelvorming, cirkelvormige uitsteeksels en macropinocytische cups op het celoppervlak in vitro te visualiseren en te kwantificeren. SEM biedt beelden met een hoge resolutie van het celoppervlak die kunnen worden gebruikt om macropinocytische membraanactiviteiten te visualiseren. Deze techniek kan worden gebruikt om nieuwe signaalroutes te ontdekken die macropinocytose reguleren en nieuwe stimulatoren en remmers van microp...

Video: Visualizing Membrane Ruffle Formation using Scanning Electron Microscopy

Method to Visualize and Analyze Membrane Interacting Proteins by Transmission Electron Microscopy

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the availability of enough area to perform the function. Nanodiscs are used to provide a membrane surface of controlled size and lipid content. In the absence of bound extrinsic proteins, sodium phosphotungstate-stained nanodiscs appear as stacks of coins when viewed from the side by transmission electron microscopy (TEM). This protocol is therefore designed to intentionally promote stacking; consequently, the prevention of stacking can be interpreted as the binding of the membrane-binding protein to the nanodisc. In a further step, the TEM images of the protein-nanodisc complexes can be processed with standard single-particle methods to yield low-resolution structures as a basis for higher resolution cryoEM work. Furthermore, the nanodiscs provide samples suitable for either TEM or non-denaturing gel electrophoresis. To illustrate the method, Ca2+-induced binding of 5-lipoxygenase on nanodiscs is presented.

Many proteins perform their function when attached to membrane surfaces. The binding of extrinsic proteins on nanodisc membranes can be indirectly imaged by transmission electron microscopy. We show that the characteristic stacking (rouleau) of nanodiscs induced by the negative stain sodium phosphotungstate is prevented by the binding of extrinsic protein.

Monotopic proteins exert their function when attached to a membrane surface, and such interactions depend on the specific lipid composition and on the ...

Biochemistry

Formation of Biomembrane Microarrays with a Squeegee-based Assembly Method

Lipid bilayer membranes form the plasma membranes of cells and define the boundaries of subcellular organelles. In nature, these membranes are heterogeneous mixtures of many types of lipids, contain membrane-bound proteins and are decorated with carbohydrates. In some experiments, it is desirable to decouple the biophysical or biochemical properties of the lipid bilayer from those of the natural membrane. Such cases call for the use of model systems such as giant vesicles, liposomes or supported lipid bilayers (SLBs). Arrays of SLBs are particularly attractive for sensing applications and mimicking cell-cell interactions. Here we describe a new method for forming SLB arrays. Submicron-diameter SiO2 beads are first coated with lipid bilayers to form spherical SLBs (SSLBs). The beads are then deposited into an array of micro-fabricated submicron-diameter microwells. The preparation technique uses a "squeegee" to clean the substrate surface, while leaving behind SSLBs that have settled into microwells. This method requires no chemical modification of the microwell substrate, nor any particular targeting ligands on the SSLB. Microwells are occupied by single beads because the well diameter is tuned to be just larger than the bead diameter. Typically, more 75% of the wells are occupied, while the rest remain empty. In buffer SSLB arrays display long-term stability of greater than one week. Multiple types of SSLBs can be placed in a single array by serial deposition, and the arrays can be used for sensing, which we demonstrate by characterizing the interaction of cholera toxin with ganglioside GM1. We also show that phospholipid vesicles without the bead supports and biomembranes from cellular sources can be arrayed with the same method and cell-specific membrane lipids can be identified.

Supported lipid bilayers and natural membrane particles are convenient systems that can approximate the properties of cell membranes and be incorporated in a variety of analytical strategies. Here we demonstrate a method for preparing microarrays composed of supported lipid bilayer-coated SiO2 beads, phospholipid vesicles or natural membrane particles.

Lipid bilayer membranes form the plasma membranes of cells and define the boundaries of subcellular organelles. In nature, these membranes are ...

Bioengineering

Mitochondria and Endoplasmic Reticulum Imaging by Correlative Light and Volume Electron Microscopy

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved structures are folded into various shapes to form a dynamic network depending on the cellular conditions. Visualization of this network between mitochondria and ER has been attempted using super-resolution fluorescence imaging and light microscopy; however, the limited resolution is insufficient to observe the membranes between the mitochondria and ER in detail. Transmission electron microscopy provides good membrane contrast and nanometer-scale resolution for the observation of cellular organelles; however, it is exceptionally time-consuming when assessing the three-dimensional (3D) structure of highly curved organelles. Therefore, we observed the morphology of mitochondria and ER via correlative light-electron microscopy (CLEM) and volume electron microscopy techniques using enhanced ascorbate peroxidase 2 and horseradish peroxidase staining. An en bloc staining method, ultrathin serial sectioning (array tomography), and volume electron microscopy were applied to observe the 3D structure. In this protocol, we suggest a combination of CLEM and 3D electron microscopy to perform detailed structural studies of mitochondria and ER.

We present a protocol to study the distribution of mitochondria and endoplasmic reticulum in whole cells after genetic modification using correlative light and volume electron microscopy including ascorbate peroxidase 2 and horseradish peroxidase staining, serial sectioning of cells with and without the target gene in the same section, and serial imaging via electron microscopy.

Cellular organelles, such as mitochondria and endoplasmic reticulum (ER), create a network to perform a variety of functions. These highly curved ...

Freeze-Fracture Electron Microscopy for Extracellular Vesicle Analysis

Extracellular vesicles (EVs) are membrane-limited structures released from the cells into the extracellular space and are implicated in intercellular communication. EVs consist of three populations of vesicles, namely microvesicles (MVs), exosomes, and apoptotic bodies. The limiting membrane of EVs is crucially involved in the interactions with the recipient cells, which could lead to the transfer of biologically active molecules to the recipient cells and, consequently, affect their behavior. The freeze-fracture electron microscopy technique is used to study the internal organization of biological membranes. Here, we present a protocol for MV isolation from cultured cancerous urothelial cells and the freeze-fracture of MVs in the steps of rapid freezing, fracturing, making and cleaning the replicas, and analyzing them with transmission electron microscopy. The results show that the protocol for isolation yields a homogenous population of EVs, which correspond to the shape and size of MVs. Intramembrane particles are found mainly in the protoplasmic face of the limiting membrane. Hence, freeze-fracture is the technique of choice to characterize the MVs' diameter, shape, and distribution of membrane proteins. The presented protocol is applicable to other populations of EVs.

We present a protocol for the isolation and freeze-fracturing of extracellular vesicles (EVs) originating from cancerous urothelial cells. The freeze-fracture technique revealed the EVs' diameter and shape and-as a unique feature-the internal organization of the EV membranes. These are of immense importance in understanding how EVs interact with the recipient membranes.

Extracellular vesicles (EVs) are membrane-limited structures released from the cells into the extracellular space and are implicated in intercellular ...

Cancer Research

Preparing Lamellae from Vitreous Biological Samples Using a Dual-Beam Scanning Electron Microscope for Cryo-Electron Tomography

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could easily be adapted for other biological samples. The basic principles for preparing samples, milling, and viewing lamellae are common to all instruments and the protocol can be followed as a general guide to on-grid cryo-lamella preparation for cryo-electron microscopy (cryoEM) and cryo-electron tomography (cryoET). Electron microscopy grids supporting the cells are plunge-frozen into liquid nitrogen-cooled liquid ethane using a manual or automated plunge freezer, then screened on a light microscope equipped with a cryo-stage. Frozen grids are transferred into a cryo-scanning electron microscope equipped with a focused ion beam (cryoFIB-SEM). Grids are routinely sputter coated prior to milling, which aids dispersal of charge build-up during milling. Alternatively, an e-beam rotary coater can be used to apply a layer of carbon-platinum to the grids, the exact thickness of which can be more precisely controlled. Once inside the cryoFIB-SEM an additional coating of an organoplatinum compound is applied to the surface of the grid via a gas injection system (GIS). This layer protects the front edge of the lamella as it is milled, the integrity of which is critical for achieving uniformly thin lamellae. Regions of interest are identified via SEM and milling is carried out in a step-wise fashion, reducing the current of the ion beam as the lamella reaches electron transparency, in order to avoid excessive heat generation. A grid with multiple lamellae is then transferred to a transmission electron microscope (TEM) under cryogenic conditions for tilt-series acquisition. A robust and contamination-free workflow for lamella preparation is an essential step for downstream techniques, including cellular cryoEM, cryoET, and sub-tomogram averaging. Development of these techniques, especially for lift-out and milling of high-pressure frozen samples, is of high-priority in the field.

Using focused ion beam milling to produce vitreous on-grid lamellae from plunge frozen biological samples for cryo-electron tomography.

Presented here is a protocol for preparing cryo-lamellae from plunge-frozen grids of Plasmodium falciparum-infected&#160;human erythrocytes, which could ...

Reconstitution of Septin Assembly at Membranes to Study Biophysical Properties and Functions

Most cells can sense and change their shape to carry out fundamental cell processes. In many eukaryotes, the septin cytoskeleton is an integral component in coordinating shape changes like cytokinesis, polarized growth, and migration. Septins are filament-forming proteins that assemble to form diverse higher-order structures and, in many cases, are found in different areas of the plasma membrane, most notably in regions of micron-scale positive curvature. Monitoring the process of septin assembly in vivo is hindered by the limitations of light microscopy in cells, as well as the complexity of interactions with both membranes and cytoskeletal elements, making it difficult to quantify septin dynamics in living systems. Fortunately, there has been substantial progress in the past decade in reconstituting the septin cytoskeleton in a cell-free system to dissect the mechanisms controlling septin assembly at high spatial and temporal resolutions. The core steps of septin assembly include septin heterooligomer association and dissociation with the membrane, polymerization into filaments, and the formation of higher-order structures through interactions between filaments. Here, we present three methods to observe septin assembly in different contexts: planar bilayers, spherical supports, and rod supports. These methods can be used to determine the biophysical parameters of septins at different stages of assembly: as single octamers binding the membrane, as filaments, and as assemblies of filaments. We use these parameters paired with measurements of curvature sampling and preferential adsorption to understand how curvature sensing operates at a variety of length and time scales.

Cell-free reconstitution has been a key tool to understand the cytoskeleton assembly, and work in the last decade has established approaches to study septin dynamics in minimal systems. Presented here are three complementary methods to observe septin assembly in different membrane contexts: planar bilayers, spherical supports, and rod supports.

Most cells can sense and change their shape to carry out fundamental cell processes. In many eukaryotes, the septin cytoskeleton is an integral component ...

Spontaneous Formation and Rearrangement of Artificial Lipid Nanotube Networks as a Bottom-Up Model for Endoplasmic Reticulum

We present a convenient method to form a bottom-up structural organelle model for the endoplasmic reticulum (ER). The model consists of highly dense lipidic nanotubes that are, in terms of morphology and dynamics, reminiscent of ER. The networks are derived from phospholipid double bilayer membrane patches adhering to a transparent Al2O3 substrate. The adhesion is mediated by Ca2+ in the ambient buffer. Subsequent depletion of Ca2+ by means of BAPTA/EDTA causes retraction of the membrane, resulting in spontaneous lipid nanotube network formation. The method only comprises phospholipids and microfabricated surfaces for simple formation of an ER model and does not require the addition of proteins or chemical energy (e.g., GTP or ATP). In contrast to the 3D morphology of the cellular endoplasmic reticulum, the model is two-dimensional (albeit the nanotube dimensions, geometry, structure, and dynamics are maintained). This unique in vitro ER model consists of only a few components, is easy to construct, and can be observed under a light microscope. The resulting structure can be further decorated for additional functionality, such as the addition of ER-associated proteins or particles to study transport phenomena among the tubes. The artificial networks described here are suitable structural models for the cellular ER, whose unique characteristic morphology has been shown to be related to its biological function, whereas details regarding formation of the tubular domain and rearrangements within are still not completely understood. We note that this method uses Al2O3 thin-film-coated microscopy coverslips, which are commercially available but require special orders. Therefore, it is advisable to have access to a microfabrication facility for preparation.

Solid-supported, protein-free, double phospholipid bilayer membranes (DLBM) can be transformed into complex and dynamic lipid nanotube networks and can serve as 2D bottom-up models of the endoplasmic reticulum.

We present a convenient method to form a bottom-up structural organelle model for the endoplasmic reticulum (ER). The model consists of highly dense ...

Visualization of Endoplasmic Reticulum Subdomains in Cultured Cells

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and functions despite the intense interorganelle molecular traffic. The techniques described in this paper are powerful means of studying protein and lipid mobility and trafficking in vivo and in their physiological environment. Fluorescence recovery after photobleaching (FRAP) and fluorescence loss in photobleaching (FLIP) are widely used live-cell imaging techniques for studying intracellular trafficking through the exo-endocytic pathway, the continuity between organelles or subcompartments, the formation of protein complexes, and protein localization in lipid microdomains, all of which can be observed under physiological and pathological conditions. The limitations of these approaches are mainly due to the use of fluorescent fusion proteins, and their potential drawbacks include artifactual over-expression in cells and the possibility of differences in the folding and localization of tagged and native proteins. Finally, as the limit of resolution of optical microscopy (about 200 nm) does not allow investigation of the fine structure of the ER or the specific subcompartments that can originate in cells under stress (i.e. hypoxia, drug administration, the over-expression of transmembrane ER resident proteins) or under pathological conditions, we combine live-cell imaging of cultured transfected cells with ultrastructural analyses based on transmission electron microscopy.

We describe the imaging approaches we use to investigate the distribution and mobility of the transfected fluorescent proteins resident in the endoplasmic reticulum (ER) by means of the confocal imaging of living cells. We also ultrastructurally analyze the effect of their expression on the architecture of this subcellular compartment.

The lipids and proteins in eukaryotic cells are continuously exchanged between cell compartments, although these retain their distinctive composition and ...

Biology

Array Tomography Workflow for the Targeted Acquisition of Volume Information using Scanning Electron Microscopy

Electron microscopy is applied in biology and medicine for imaging of cellular and structural details at nanometer resolution. Historically, Transmission Electron Microscopy (TEM) provided insight into cell ultrastructure, but in the recent decade, the development of modern Scanning Electron Microscopes (SEM) has changed the way of looking inside the cells. Even though the resolution of TEM is superior when protein-level structural details are needed, SEM-resolution is sufficient for the majority of the organelle-level cell biology-related questions. The advancement in technology enabled automatic volume acquisition solutions such as Serial block-face imaging (SBF-SEM) and Focused ion beam SEM (FIB-SEM). Nevertheless, to this day, these methods remain inefficient when the identification and navigation to areas of interest are crucial. Without the means for precise localization of target areas before imaging, operators need to acquire much more data than they need (in SBF-SEM), or, even worse, prepare many grids and image them all (in TEM). We propose the strategy of "lateral screening" using Array Tomography in SEM, which facilitates the localization of areas of interest, followed by automated imaging of the relevant fraction of the total sample volume. Array tomography samples are conserved during imaging, and they can be arranged into section libraries ready for repeated imaging. Several examples are shown in which lateral screening enables us to analyze structural details that are incredibly challenging to access with any other method.

We describe the preparation of ribbons of serial sections and their collection on large transfer support for use as Array Tomography samples, along with automated imaging procedures in a scanning electron microscope. The protocol allows screening, retrieval, and targeted imaging of local, rare events, and the acquisition of large data volumes.

Electron microscopy is applied in biology and medicine for imaging of cellular and structural details at nanometer resolution. Historically, Transmission ...

Bottom-Up In Vitro Methods to Assay the Ultrastructural Organization, Membrane Reshaping, and Curvature Sensitivity Behavior of Septins

Membrane remodeling occurs constantly at the plasma membrane and within cellular organelles. To fully dissect the role of the environment (ionic conditions, protein and lipid compositions, membrane curvature) and the different partners associated with specific membrane reshaping processes, we undertake in vitro bottom-up approaches. In recent years, there has been keen interest in revealing the role of septin proteins associated with major diseases. Septins are essential and ubiquitous cytoskeletal proteins that interact with the plasma membrane. They are implicated in cell division, cell motility, neuro-morphogenesis, and spermiogenesis, among other functions. It is, therefore, important to understand how septins interact and organize at membranes to subsequently induce membrane deformations and how they can be sensitive to specific membrane curvatures. This article aims to decipher the interplay between the ultra-structure of septins at a molecular level and the membrane remodeling occurring at a micron scale. To this end, budding yeast, and mammalian septin complexes were recombinantly expressed and purified. A combination of in vitro assays was then used to analyze the self-assembly of septins at the membrane. Supported lipid bilayers (SLBs), giant unilamellar vesicles (GUVs), large unilamellar vesicles (LUVs), and wavy substrates were used to study the interplay between septin self-assembly, membrane reshaping, and membrane curvature.

Bottom-Up In Vitro Methods to Assay the Ultrastructural Organization, Membrane Reshaping, and Curvature Sensitivity Behavior of Septins

Septins are cytoskeletal proteins. They interact with lipid membranes and can sense but also generate membrane curvature at the micron scale. We describe in this protocol bottom-up in vitro methodologies for analyzing membrane deformations, curvature-sensitive septin binding, and septin filament ultrastructure.

Visualizing Membrane Ruffle Formation using Scanning Electron Microscopy

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