Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction with other compounds, such as drugs or peptides. However SLB characteristics differ depending on the support used.
Commonly used techniques for SLB imaging and measurements are single molecule fluorescence microscopy, FCS and atomic force microscopy (AFM). Because most optical imaging studies are carried out on a glass support, while AFM requires an extremely flat surface (generally mica), results from these techniques cannot be compared directly, since the charge and smoothness properties of these materials strongly influence diffusion. Unfortunately, the high level of manual dexterity required for the cutting and gluing thin slices of mica to the glass slide presents a hurdle to routine use of mica for SLB preparation. Although this would be the method of choice, such prepared mica surfaces often end up being uneven (wavy) and difficult to image, especially with small working distance, high numerical aperture lenses. Here we present a simple and reproducible method for preparing thin, flat mica surfaces for lipid vesicle deposition and SLB preparation. Additionally, our custom made chamber requires only very small volumes of vesicles for SLB formation. The overall procedure results in the efficient, simple and inexpensive production of high quality lipid bilayer surfaces that are directly comparable to those used in AFM studies.
22 Related JoVE Articles!
High-throughput Crystallization of Membrane Proteins Using the Lipidic Bicelle Method
Institutions: University of California Los Angeles , David Geffen School of Medicine, UCLA.
Membrane proteins (MPs) play a critical role in many physiological processes such as pumping specific molecules across the otherwise impermeable membrane bilayer that surrounds all cells and organelles. Alterations in the function of MPs result in many human diseases and disorders; thus, an intricate understanding of their structures remains a critical objective for biological research. However, structure determination of MPs remains a significant challenge often stemming from their hydrophobicity.
MPs have substantial hydrophobic regions embedded within the bilayer. Detergents are frequently used to solubilize these proteins from the bilayer generating a protein-detergent micelle that can then be manipulated in a similar manner as soluble proteins. Traditionally, crystallization trials proceed using a protein-detergent mixture, but they often resist crystallization or produce crystals of poor quality. These problems arise due to the detergent′s inability to adequately mimic the bilayer resulting in poor stability and heterogeneity. In addition, the detergent shields the hydrophobic surface of the MP reducing the surface area available for crystal contacts. To circumvent these drawbacks MPs can be crystallized in lipidic media, which more closely simulates their endogenous environment, and has recently become a de novo technique for MP crystallization.
Lipidic cubic phase (LCP) is a three-dimensional lipid bilayer penetrated by an interconnected system of aqueous channels1
. Although monoolein is the lipid of choice, related lipids such as monopalmitolein and monovaccenin have also been used to make LCP2
. MPs are incorporated into the LCP where they diffuse in three dimensions and feed crystal nuclei. A great advantage of the LCP is that the protein remains in a more native environment, but the method has a number of technical disadvantages including high viscosity (requiring specialized apparatuses) and difficulties in crystal visualization and manipulation3,4
. Because of these technical difficulties, we utilized another lipidic medium for crystallization-bicelles5,6
). Bicelles are lipid/amphiphile mixtures formed by blending a phosphatidylcholine lipid (DMPC) with an amphiphile (CHAPSO) or a short-chain lipid (DHPC). Within each bicelle disc, the lipid molecules generate a bilayer while the amphiphile molecules line the apolar edges providing beneficial properties of both bilayers and detergents. Importantly, below their transition temperature, protein-bicelle mixtures have a reduced viscosity and are manipulated in a similar manner as detergent-solubilized MPs, making bicelles compatible with crystallization robots.
Bicelles have been successfully used to crystallize several membrane proteins5,7-11 (Table 1)
. This growing collection of proteins demonstrates the versatility of bicelles for crystallizing both alpha helical and beta sheet MPs from prokaryotic and eukaryotic sources. Because of these successes and the simplicity of high-throughput implementation, bicelles should be part of every membrane protein crystallographer′s arsenal. In this video, we describe the bicelle methodology and provide a step-by-step protocol for setting up high-throughput crystallization trials of purified MPs using standard robotics.
Molecular Biology, Issue 59, membrane proteins crystallization, bicelle, lipidic crystallization
Gramicidin-based Fluorescence Assay; for Determining Small Molecules Potential for Modifying Lipid Bilayer Properties
Institutions: Weill Cornell Medical College.
Many drugs and other small molecules used to modulate biological function are amphiphiles that adsorb at the bilayer/solution interface and thereby alter lipid bilayer properties. This is important because membrane proteins are energetically coupled to their host bilayer by hydrophobic interactions. Changes in bilayer properties thus alter membrane protein function, which provides an indirect way for amphiphiles to modulate protein function and a possible mechanism for "off-target" drug effects. We have previously developed an electrophysiological assay for detecting changes in lipid bilayer properties using linear gramicidin channels as probes 3,12
. Gramicidin channels are mini-proteins formed by the transbilayer dimerization of two non-conducting subunits. They are sensitive to changes in their membrane environment, which makes them powerful probes for monitoring changes in lipid bilayer properties as sensed by bilayer spanning proteins. We now demonstrate a fluorescence assay for detecting changes in bilayer properties using the same channels as probes. The assay is based on measuring the time-course of fluorescence quenching from fluorophore-loaded large unilamellar vesicles due to the entry of a quencher through the gramicidin channels. We use the fluorescence indicator/quencher pair 8-aminonaphthalene-1,3,6-trisulfonate (ANTS)/Tl+
that has been successfully used in other fluorescence quenching assays 5,13
permeates the lipid bilayer slowly 8
but passes readily through conducting gramicidin channels 1,14
. The method is scalable and suitable for both mechanistic studies and high-throughput screening of small molecules for bilayer-perturbing, and potential "off-target", effects. We find that results using this method are in good agreement with previous electrophysiological results 12
Microbiology, Issue 44, membrane properties, bilayer properties, gramicidin, fluorescence quenching, high throughput drug screening
Method for Measurement of Viral Fusion Kinetics at the Single Particle Level
Institutions: Harvard Medical School, Harvard Medical School.
Membrane fusion is an essential step during entry of enveloped viruses into cells. Conventional fusion assays typically report on a large number of fusion events, making it difficult to quantitatively analyze the sequence of the molecular steps involved. We have developed an in vitro,
two-color fluorescence assay to monitor kinetics of single virus particles fusing with a target bilayer on an essentially fluid support.
Influenza viral particles are incubated with a green lipophilic fluorophore to stain the membrane and a red hydrophilic fluorophore to stain the viral interior. We deposit a ganglioside-containing lipid bilayer on the dextran-functionilized glass surface of a flow cell, incubate the viral particles on the planar bilayer and image the fluorescence of a 100 x 100 μm2
area, containing several hundreds of particles, on a CCD camera. By imaging both the red and green fluorescence, we can simultaneously monitor the behavior of the membrane dye (green) and the aqueous content (red) of the particles.
Upon lowering the pH to a value below the fusion pH, the particles will fuse with the membrane. Hemifusion, the merging of the outer leaflet of the viral membrane with the outer leaflet of the target membrane, will be visible as a sudden change in the green fluorescence of a particle. Upon the subsequent fusion of the two remaining distal leaflets a pore will be formed and the red-emitting fluorophore in the viral particle will be released under the target membrane. This event will give rise to a decrease of the red fluorescence of individual particles. Finally, the integrated fluorescence from a pH-sensitive fluorophore that is embedded in the target membrane reports on the exact time of the pH drop.
From the three fluorescence-time traces, all the important events (pH drop, lipid mixing upon hemifusion, content mixing upon pore formation) can now be extracted in a straightforward manner and for every particle individually. By collecting the elapsed times for the various transitions for many individual particles in histograms, we can determine the lifetimes of the corresponding intermediates. Even hidden intermediates that do not have a direct fluorescent observable can be visualized directly from these histograms.
Biomedical Engineering, Issue 31, Viral fusion, membrane fusion, supported lipid bilayer, biophysics, single molecule
Determination of Lipid Raft Partitioning of Fluorescently-tagged Probes in Living Cells by Fluorescence Correlation Spectroscopy (FCS)
Institutions: Hôpital de la Pitié-Salpêtrière, Université Paris-Sud, Université Paris-Sud.
In the past fifteen years the notion that cell membranes are not homogenous and rely on microdomains to exert their functions has become widely accepted. Lipid rafts are membrane microdomains enriched in cholesterol and sphingolipids. They play a role in cellular physiological processes such as signalling, and trafficking1,2
but are also thought to be key players in several diseases including viral or bacterial infections and neurodegenerative diseases3
Yet their existence is still a matter of controversy4,5
. Indeed, lipid raft size has been estimated to be around 20 nm6
, far under the resolution limit of conventional microscopy (around 200 nm), thus precluding their direct imaging. Up to now, the main techniques used to assess the partition of proteins of interest inside lipid rafts were Detergent Resistant Membranes (DRMs) isolation and co-patching with antibodies. Though widely used because of their rather easy implementation, these techniques were prone to artefacts and thus criticized7,8
. Technical improvements were therefore necessary to overcome these artefacts and to be able to probe lipid rafts partition in living cells.
Here we present a method for the sensitive analysis of lipid rafts partition of fluorescently-tagged proteins or lipids in the plasma membrane of living cells. This method, termed Fluorescence Correlation Spectroscopy (FCS), relies on the disparity in diffusion times of fluorescent probes located inside or outside of lipid rafts. In fact, as evidenced in both artificial membranes and cell cultures, probes would diffuse much faster outside than inside dense lipid rafts9,10
. To determine diffusion times, minute fluorescence fluctuations are measured as a function of time in a focal volume (approximately 1 femtoliter), located at the plasma membrane of cells with a confocal microscope (Fig. 1
). The auto-correlation curves can then be drawn from these fluctuations and fitted with appropriate mathematical diffusion models11
FCS can be used to determine the lipid raft partitioning of various probes, as long as they are fluorescently tagged. Fluorescent tagging can be achieved by expression of fluorescent fusion proteins or by binding of fluorescent ligands. Moreover, FCS can be used not only in artificial membranes and cell lines but also in primary cultures, as described recently12
. It can also be used to follow the dynamics of lipid raft partitioning after drug addition or membrane lipid composition change12
Cellular Biology, Issue 62, Lipid rafts, plasma membrane, diffusion times, confocal microscopy, fluorescence correlation spectroscopy (FCS)
Crystallization of Membrane Proteins in Lipidic Mesophases
Institutions: The Scripps Research Institute.
Membrane proteins perform critical functions in living cells related to signal transduction, transport and energy transformations, and, as such, are implicated in a multitude of malfunctions and diseases. However, a structural and functional understanding of membrane proteins is strongly lagging behind that of their soluble partners, mainly, due to difficulties associated with their solubilization and generation of diffraction quality crystals. Crystallization in lipidic mesophases (also known as in meso
or LCP crystallization) is a promising technique which was successfully applied to obtain high resolution structures of microbial rhodopsins, photosynthetic proteins, outer membrane beta barrels and G protein-coupled receptors. In meso crystallization takes advantage of a native-like membrane environment and typically produces crystals with lower solvent content and better ordering as compared to traditional crystallization from detergent solutions. The method is not difficult, but requires an understanding of lipid phase behavior and practice in handling viscous mesophase materials. Here we demonstrate a simple and efficient way of making LCP and reconstituting a membrane protein in the lipid bilayer of LCP using a syringe mixer, followed by dispensing nanoliter portions of LCP into an assay or crystallization plate, conducting pre-crystallization assays and harvesting crystals from the LCP matrix. These protocols provide a basic guide for approaching in meso
crystallization trials; however, as with any crystallization experiment, extensive screening and optimization are required, and a successful outcome is not necessarily guaranteed.
Structural Biology, Issue 49, membrane protein, lipidic cubic phase, crystallization, Fluorescence recovery after photobleaching (FRAP) , G protein-coupled receptors
Giant Liposome Preparation for Imaging and Patch-Clamp Electrophysiology
Institutions: University of Washington.
The reconstitution of ion channels into chemically defined lipid membranes for electrophysiological recording has been a powerful technique to identify and explore the function of these important proteins. However, classical preparations, such as planar bilayers, limit the manipulations and experiments that can be performed on the reconstituted channel and its membrane environment. The more cell-like structure of giant liposomes permits traditional patch-clamp experiments without sacrificing control of the lipid environment.
Electroformation is an efficient mean to produce giant liposomes >10 μm in diameter which relies on the application of alternating voltage to a thin, ordered lipid film deposited on an electrode surface. However, since the classical protocol calls for the lipids to be deposited from organic solvents, it is not compatible with less robust membrane proteins like ion channels and must be modified. Recently, protocols have been developed to electroform giant liposomes from partially dehydrated small liposomes, which we have adapted to protein-containing liposomes in our laboratory.
We present here the background, equipment, techniques, and pitfalls of electroformation of giant liposomes from small liposome dispersions. We begin with the classic protocol, which should be mastered first before attempting the more challenging protocols that follow. We demonstrate the process of controlled partial dehydration of small liposomes using vapor equilibrium with saturated salt solutions. Finally, we demonstrate the process of electroformation itself. We will describe simple, inexpensive equipment that can be made in-house to produce high-quality liposomes, and describe visual inspection of the preparation at each stage to ensure the best results.
Physiology, Issue 76, Biophysics, Molecular Biology, Biochemistry, Genetics, Cellular Biology, Proteins, Membranes, Artificial, Lipid Bilayers, Liposomes, Phospholipids, biochemistry, Lipids, Giant Unilamellar Vesicles, liposome, electrophysiology, electroformation, reconstitution, patch clamp
Isolation and Chemical Characterization of Lipid A from Gram-negative Bacteria
Institutions: The University of Texas at Austin, The University of Texas at Austin, The University of Texas at Austin.
Lipopolysaccharide (LPS) is the major cell surface molecule of gram-negative bacteria, deposited on the outer leaflet of the outer membrane bilayer. LPS can be subdivided into three domains: the distal O-polysaccharide, a core oligosaccharide, and the lipid A domain consisting of a lipid A molecular species and 3-deoxy-D-manno-oct-2-ulosonic acid residues (Kdo). The lipid A domain is the only component essential for bacterial cell survival. Following its synthesis, lipid A is chemically modified in response to environmental stresses such as pH or temperature, to promote resistance to antibiotic compounds, and to evade recognition by mediators of the host innate immune response. The following protocol details the small- and large-scale isolation of lipid A from gram-negative bacteria. Isolated material is then chemically characterized by thin layer chromatography (TLC) or mass-spectrometry (MS). In addition to matrix-assisted laser desorption/ionization-time of flight (MALDI-TOF) MS, we also describe tandem MS protocols for analyzing lipid A molecular species using electrospray ionization (ESI) coupled to collision induced dissociation (CID) and newly employed ultraviolet photodissociation (UVPD) methods. Our MS protocols allow for unequivocal determination of chemical structure, paramount to characterization of lipid A molecules that contain unique or novel chemical modifications. We also describe the radioisotopic labeling, and subsequent isolation, of lipid A from bacterial cells for analysis by TLC. Relative to MS-based protocols, TLC provides a more economical and rapid characterization method, but cannot be used to unambiguously assign lipid A chemical structures without the use of standards of known chemical structure. Over the last two decades isolation and characterization of lipid A has led to numerous exciting discoveries that have improved our understanding of the physiology of gram-negative bacteria, mechanisms of antibiotic resistance, the human innate immune response, and have provided many new targets in the development of antibacterial compounds.
Chemistry, Issue 79, Membrane Lipids, Toll-Like Receptors, Endotoxins, Glycolipids, Lipopolysaccharides, Lipid A, Microbiology, Lipids, lipid A, Bligh-Dyer, thin layer chromatography (TLC), lipopolysaccharide, mass spectrometry, Collision Induced Dissociation (CID), Photodissociation (PD)
Multi-step Preparation Technique to Recover Multiple Metabolite Compound Classes for In-depth and Informative Metabolomic Analysis
Institutions: National Jewish Health, University of Colorado Denver.
Metabolomics is an emerging field which enables profiling of samples from living organisms in order to obtain insight into biological processes. A vital aspect of metabolomics is sample preparation whereby inconsistent techniques generate unreliable results. This technique encompasses protein precipitation, liquid-liquid extraction, and solid-phase extraction as a means of fractionating metabolites into four distinct classes. Improved enrichment of low abundance molecules with a resulting increase in sensitivity is obtained, and ultimately results in more confident identification of molecules. This technique has been applied to plasma, bronchoalveolar lavage fluid, and cerebrospinal fluid samples with volumes as low as 50 µl. Samples can be used for multiple downstream applications; for example, the pellet resulting from protein precipitation can be stored for later analysis. The supernatant from that step undergoes liquid-liquid extraction using water and strong organic solvent to separate the hydrophilic and hydrophobic compounds. Once fractionated, the hydrophilic layer can be processed for later analysis or discarded if not needed. The hydrophobic fraction is further treated with a series of solvents during three solid-phase extraction steps to separate it into fatty acids, neutral lipids, and phospholipids. This allows the technician the flexibility to choose which class of compounds is preferred for analysis. It also aids in more reliable metabolite identification since some knowledge of chemical class exists.
Bioengineering, Issue 89, plasma, chemistry techniques, analytical, solid phase extraction, mass spectrometry, metabolomics, fluids and secretions, profiling, small molecules, lipids, liquid chromatography, liquid-liquid extraction, cerebrospinal fluid, bronchoalveolar lavage fluid
In-vivo Centrifugation of Drosophila Embryos
Institutions: University of Rochester.
A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in buoyant density. However, when cells are disrupted prior to centrifugation, proteins and organelles in this non-native environment often inappropriately stick to each other. Here we describe a method to separate organelles by density in intact, living Drosophila
embryos. Early embryos before cellularization are harvested from population cages, and their outer egg shells are removed by treatment with 50% bleach. Embryos are then transferred to a small agar plate and inserted, posterior end first, into small vertical holes in the agar. The plates containing embedded embryos are centrifuged for 30 min at 3000g. The agar supports the embryos and keeps them in a defined orientation. Afterwards, the embryos are dug out of the agar with a blunt needle. Centrifugation separates major organelles into distinct layers, a stratification easily visible by bright-field microscopy. A number of fluorescent markers are available to confirm successful stratification in living embryos. Proteins associated with certain organelles will be enriched in a particular layer, demonstrating colocalization. Individual layers can be recovered for biochemical analysis or transplantation into donor eggs. This technique is applicable for organelle separation in other large cells, including the eggs and oocytes of diverse species.
Cellular Biology, Issue 40, Drosophila, embryo, centrifugation, organelle, lipid droplet, yolk, colocalization, transplantation
Imaging Plasma Membrane Deformations With pTIRFM
Institutions: Wayne State University.
To gain novel insights into the dynamics of exocytosis, our group focuses on the changes in lipid bilayer shape that must be precisely regulated during the fusion of vesicle and plasma membranes. These rapid and localized changes are achieved by dynamic interactions between lipids and specialized proteins that control membrane curvature. The absence of such interactions would not only have devastating consequences for vesicle fusion, but a host of other cellular functions that involve control of membrane shape. In recent years, the identity of a number of proteins with membrane-shaping properties has been determined. What remains missing is a roadmap of when, where, and how they act as fusion and content release progress.
Our understanding of the molecular events that enable membrane remodeling has historically been limited by a lack of analytical methods that are sensitive to membrane curvature or have the temporal resolution to track rapid changes. PTIRFM satisfies both of these criteria. We discuss how pTIRFM is implemented to visualize and interpret rapid, submicron changes in the orientation of chromaffin cell membranes during dense core vesicle (DCV) fusion. The chromaffin cells we use are isolated from bovine adrenal glands. The membrane is stained with a lipophilic carbocyanine dye,1,1'-dioctadecyl-3,3,3',3'-tetramethylindodicarbocyanine, 4-chlorobenzenesulfonate, or diD. DiD intercalates in the membrane plane with a "fixed" orientation and is therefore sensitive to the polarization of the evanescent field. The diD-stained cell membrane is sequentially excited with orthogonal polarizations of a 561 nm laser (p-pol, s-pol). A 488 nm laser is used to visualize vesicle constituents and time the moment of fusion. Exocytosis is triggered by locally perfusing cells with a depolarizing KCl solution. Analysis is performed offline using custom-written software to understand how diD emission intensity changes relate to fusion pore dilation.
Biochemistry, Issue 86, Chromaffin Cells, Lipid Bilayers, Microscopy, Fluorescence, Polarization, Exocytosis, membrane, TIRF, pTIRF, chromaffin, polarization, vesicle
Optimized Negative Staining: a High-throughput Protocol for Examining Small and Asymmetric Protein Structure by Electron Microscopy
Institutions: The Molecular Foundry.
Structural determination of proteins is rather challenging for proteins with molecular masses between 40 - 200 kDa. Considering that more than half of natural proteins have a molecular mass between 40 - 200 kDa1,2
, a robust and high-throughput method with a nanometer resolution capability is needed. Negative staining (NS) electron microscopy (EM) is an easy, rapid, and qualitative approach which has frequently been used in research laboratories to examine protein structure and protein-protein interactions. Unfortunately, conventional NS protocols often generate structural artifacts on proteins, especially with lipoproteins that usually form presenting rouleaux artifacts. By using images of lipoproteins from cryo-electron microscopy (cryo-EM) as a standard, the key parameters in NS specimen preparation conditions were recently screened and reported as the optimized NS protocol (OpNS), a modified conventional NS protocol 3
. Artifacts like rouleaux can be greatly limited by OpNS, additionally providing high contrast along with reasonably high‐resolution (near 1 nm) images of small and asymmetric proteins. These high-resolution and high contrast images are even favorable for an individual protein (a single object, no average) 3D reconstruction, such as a 160 kDa antibody, through the method of electron tomography4,5
. Moreover, OpNS can be a high‐throughput tool to examine hundreds of samples of small proteins. For example, the previously published mechanism of 53 kDa cholesteryl ester transfer protein (CETP) involved the screening and imaging of hundreds of samples 6
. Considering cryo-EM rarely successfully images proteins less than 200 kDa has yet to publish any study involving screening over one hundred sample conditions, it is fair to call OpNS a high-throughput method for studying small proteins. Hopefully the OpNS protocol presented here can be a useful tool to push the boundaries of EM and accelerate EM studies into small protein structure, dynamics and mechanisms.
Environmental Sciences, Issue 90, small and asymmetric protein structure, electron microscopy, optimized negative staining
Reconstitution of a Kv Channel into Lipid Membranes for Structural and Functional Studies
Institutions: University of Texas Southwestern Medical Center at Dallas.
To study the lipid-protein interaction in a reductionistic fashion, it is necessary to incorporate the membrane proteins into membranes of well-defined lipid composition. We are studying the lipid-dependent gating effects in a prototype voltage-gated potassium (Kv) channel, and have worked out detailed procedures to reconstitute the channels into different membrane systems. Our reconstitution procedures take consideration of both detergent-induced fusion of vesicles and the fusion of protein/detergent micelles with the lipid/detergent mixed micelles as well as the importance of reaching an equilibrium distribution of lipids among the protein/detergent/lipid and the detergent/lipid mixed micelles. Our data suggested that the insertion of the channels in the lipid vesicles is relatively random in orientations, and the reconstitution efficiency is so high that no detectable protein aggregates were seen in fractionation experiments. We have utilized the reconstituted channels to determine the conformational states of the channels in different lipids, record electrical activities of a small number of channels incorporated in planar lipid bilayers, screen for conformation-specific ligands from a phage-displayed peptide library, and support the growth of 2D crystals of the channels in membranes. The reconstitution procedures described here may be adapted for studying other membrane proteins in lipid bilayers, especially for the investigation of the lipid effects on the eukaryotic voltage-gated ion channels.
Molecular Biology, Issue 77, Biochemistry, Genetics, Cellular Biology, Structural Biology, Biophysics, Membrane Lipids, Phospholipids, Carrier Proteins, Membrane Proteins, Micelles, Molecular Motor Proteins, life sciences, biochemistry, Amino Acids, Peptides, and Proteins, lipid-protein interaction, channel reconstitution, lipid-dependent gating, voltage-gated ion channel, conformation-specific ligands, lipids
Isolation of Cellular Lipid Droplets: Two Purification Techniques Starting from Yeast Cells and Human Placentas
Institutions: University of Tennessee, University of Tennessee.
Lipid droplets are dynamic organelles that can be found in most eukaryotic and certain prokaryotic cells. Structurally, the droplets consist of a core of neutral lipids surrounded by a phospholipid monolayer. One of the most useful techniques in determining the cellular roles of droplets has been proteomic identification of bound proteins, which can be isolated along with the droplets. Here, two methods are described to isolate lipid droplets and their bound proteins from two wide-ranging eukaryotes: fission yeast and human placental villous cells. Although both techniques have differences, the main method - density gradient centrifugation - is shared by both preparations. This shows the wide applicability of the presented droplet isolation techniques.
In the first protocol, yeast cells are converted into spheroplasts by enzymatic digestion of their cell walls. The resulting spheroplasts are then gently lysed in a loose-fitting homogenizer. Ficoll is added to the lysate to provide a density gradient, and the mixture is centrifuged three times. After the first spin, the lipid droplets are localized to the white-colored floating layer of the centrifuge tubes along with the endoplasmic reticulum (ER), the plasma membrane, and vacuoles. Two subsequent spins are used to remove these other three organelles. The result is a layer that has only droplets and bound proteins.
In the second protocol, placental villous cells are isolated from human term placentas by enzymatic digestion with trypsin and DNase I. The cells are homogenized in a loose-fitting homogenizer. Low-speed and medium-speed centrifugation steps are used to remove unbroken cells, cellular debris, nuclei, and mitochondria. Sucrose is added to the homogenate to provide a density gradient and the mixture is centrifuged to separate the lipid droplets from the other cellular fractions.
The purity of the lipid droplets in both protocols is confirmed by Western Blot analysis. The droplet fractions from both preps are suitable for subsequent proteomic and lipidomic analysis.
Bioengineering, Issue 86, Lipid droplet, lipid body, fat body, oil body, Yeast, placenta, placental villous cells, isolation, purification, density gradient centrifugation
Formation of Biomembrane Microarrays with a Squeegee-based Assembly Method
Institutions: University of Minnesota, University of Minnesota, Mayo Clinic College of Medicine, Mayo Clinic College of Medicine.
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.
Bioengineering, Issue 87, supported lipid bilayer, beads, microarray, fluorescence, microfabrication, nanofabrication, atomic layer deposition, myelin, lipid rafts
Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans
Institutions: Massachusetts General Hospital and Harvard Medical School, Massachusetts Institute of Technology.
The nematode C. elegans
has emerged as an important model for the study of conserved genetic pathways regulating fat metabolism as it relates to human obesity and its associated pathologies. Several previous methodologies developed for the visualization of C. elegans
triglyceride-rich fat stores have proven to be erroneous, highlighting cellular compartments other than lipid droplets. Other methods require specialized equipment, are time-consuming, or yield inconsistent results. We introduce a rapid, reproducible, fixative-based Nile red staining method for the accurate and rapid detection of neutral lipid droplets in C. elegans
. A short fixation step in 40% isopropanol makes animals completely permeable to Nile red, which is then used to stain animals. Spectral properties of this lipophilic dye allow it to strongly and selectively fluoresce in the yellow-green spectrum only when in a lipid-rich environment, but not in more polar environments. Thus, lipid droplets can be visualized on a fluorescent microscope equipped with simple GFP imaging capability after only a brief Nile red staining step in isopropanol. The speed, affordability, and reproducibility of this protocol make it ideally suited for high throughput screens. We also demonstrate a paired method for the biochemical determination of triglycerides and phospholipids using gas chromatography mass-spectrometry. This more rigorous protocol should be used as confirmation of results obtained from the Nile red microscopic lipid determination. We anticipate that these techniques will become new standards in the field of C. elegans
Genetics, Issue 73, Biochemistry, Cellular Biology, Molecular Biology, Developmental Biology, Physiology, Anatomy, Caenorhabditis elegans, Obesity, Energy Metabolism, Lipid Metabolism, C. elegans, fluorescent lipid staining, lipids, Nile red, fat, high throughput screening, obesity, gas chromatography, mass spectrometry, GC/MS, animal model
Proton Transfer and Protein Conformation Dynamics in Photosensitive Proteins by Time-resolved Step-scan Fourier-transform Infrared Spectroscopy
Institutions: Freie Universität Berlin.
Monitoring the dynamics of protonation and protein backbone conformation changes during the function of a protein is an essential step towards understanding its mechanism. Protonation and conformational changes affect the vibration pattern of amino acid side chains and of the peptide bond, respectively, both of which can be probed by infrared (IR) difference spectroscopy. For proteins whose function can be repetitively and reproducibly triggered by light, it is possible to obtain infrared difference spectra with (sub)microsecond resolution over a broad spectral range using the step-scan Fourier transform infrared technique. With ~102
repetitions of the photoreaction, the minimum number to complete a scan at reasonable spectral resolution and bandwidth, the noise level in the absorption difference spectra can be as low as ~10-4
, sufficient to follow the kinetics of protonation changes from a single amino acid. Lower noise levels can be accomplished by more data averaging and/or mathematical processing. The amount of protein required for optimal results is between 5-100 µg, depending on the sampling technique used. Regarding additional requirements, the protein needs to be first concentrated in a low ionic strength buffer and then dried to form a film. The protein film is hydrated prior to the experiment, either with little droplets of water or under controlled atmospheric humidity. The attained hydration level (g of water / g of protein) is gauged from an IR absorption spectrum. To showcase the technique, we studied the photocycle of the light-driven proton-pump bacteriorhodopsin in its native purple membrane environment, and of the light-gated ion channel channelrhodopsin-2 solubilized in detergent.
Biophysics, Issue 88, bacteriorhodopsin, channelrhodopsin, attenuated total reflection, proton transfer, protein dynamics, infrared spectroscopy, time-resolved spectroscopy, step-scan, membrane proteins, singular value decomposition
Lipid Bilayer Vesicle Generation Using Microfluidic Jetting
Institutions: University of Michigan, University of Michigan, The University of Texas at Austin, University of California, Berkeley, Lawrence Berkeley National Laboratory.
Bottom-up synthetic biology presents a novel approach for investigating and reconstituting biochemical systems and, potentially, minimal organisms. This emerging field engages engineers, chemists, biologists, and physicists to design and assemble basic biological components into complex, functioning systems from the bottom up. Such bottom-up systems could lead to the development of artificial cells for fundamental biological inquiries and innovative therapies1,2
. Giant unilamellar vesicles (GUVs) can serve as a model platform for synthetic biology due to their cell-like membrane structure and size. Microfluidic jetting, or microjetting, is a technique that allows for the generation of GUVs with controlled size, membrane composition, transmembrane protein incorporation, and encapsulation3
. The basic principle of this method is the use of multiple, high-frequency fluid pulses generated by a piezo-actuated inkjet device to deform a suspended lipid bilayer into a GUV. The process is akin to blowing soap bubbles from a soap film. By varying the composition of the jetted solution, the composition of the encompassing solution, and/or the components included in the bilayer, researchers can apply this technique to create customized vesicles. This paper describes the procedure to generate simple vesicles from a droplet interface bilayer by microjetting.
Bioengineering, Issue 84, Microfluidic jetting, synthetic biology, vesicle encapsulation, lipid bilayer, biochemical reconstitution, giant unilamellar vesicles
From Fast Fluorescence Imaging to Molecular Diffusion Law on Live Cell Membranes in a Commercial Microscope
Institutions: Scuola Normale Superiore, Instituto Italiano di Tecnologia, University of California, Irvine.
It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the regulation of many cellular functions. However, due to the fast dynamics and the tiny structures involved, a very high spatio-temporal resolution is required to catch the real behavior of molecules. Here we present the experimental protocol for studying the dynamics of fluorescently-labeled plasma-membrane proteins and lipids in live cells with high spatiotemporal resolution. Notably, this approach doesn’t need to track each molecule, but it calculates population behavior using all molecules in a given region of the membrane. The starting point is a fast imaging of a given region on the membrane. Afterwards, a complete spatio-temporal autocorrelation function is calculated correlating acquired images at increasing time delays, for example each 2, 3, n repetitions. It is possible to demonstrate that the width of the peak of the spatial autocorrelation function increases at increasing time delay as a function of particle movement due to diffusion. Therefore, fitting of the series of autocorrelation functions enables to extract the actual protein mean square displacement from imaging (iMSD), here presented in the form of apparent diffusivity vs average displacement. This yields a quantitative view of the average dynamics of single molecules with nanometer accuracy. By using a GFP-tagged variant of the Transferrin Receptor (TfR) and an ATTO488 labeled 1-palmitoyl-2-hydroxy-sn
-glycero-3-phosphoethanolamine (PPE) it is possible to observe the spatiotemporal regulation of protein and lipid diffusion on µm-sized membrane regions in the micro-to-milli-second time range.
Bioengineering, Issue 92, fluorescence, protein dynamics, lipid dynamics, membrane heterogeneity, transient confinement, single molecule, GFP
Simultaneous Multicolor Imaging of Biological Structures with Fluorescence Photoactivation Localization Microscopy
Institutions: University of Maine.
Localization-based super resolution microscopy can be applied to obtain a spatial map (image) of the distribution of individual fluorescently labeled single molecules within a sample with a spatial resolution of tens of nanometers. Using either photoactivatable (PAFP) or photoswitchable (PSFP) fluorescent proteins fused to proteins of interest, or organic dyes conjugated to antibodies or other molecules of interest, fluorescence photoactivation localization microscopy (FPALM) can simultaneously image multiple species of molecules within single cells. By using the following approach, populations of large numbers (thousands to hundreds of thousands) of individual molecules are imaged in single cells and localized with a precision of ~10-30 nm. Data obtained can be applied to understanding the nanoscale spatial distributions of multiple protein types within a cell. One primary advantage of this technique is the dramatic increase in spatial resolution: while diffraction limits resolution to ~200-250 nm in conventional light microscopy, FPALM can image length scales more than an order of magnitude smaller. As many biological hypotheses concern the spatial relationships among different biomolecules, the improved resolution of FPALM can provide insight into questions of cellular organization which have previously been inaccessible to conventional fluorescence microscopy. In addition to detailing the methods for sample preparation and data acquisition, we here describe the optical setup for FPALM. One additional consideration for researchers wishing to do super-resolution microscopy is cost: in-house setups are significantly cheaper than most commercially available imaging machines. Limitations of this technique include the need for optimizing the labeling of molecules of interest within cell samples, and the need for post-processing software to visualize results. We here describe the use of PAFP and PSFP expression to image two protein species in fixed cells. Extension of the technique to living cells is also described.
Basic Protocol, Issue 82, Microscopy, Super-resolution imaging, Multicolor, single molecule, FPALM, Localization microscopy, fluorescent proteins
Supported Planar Bilayers for the Formation of Study of Immunological Synapses and Kinapse
Institutions: New York University - NYU.
Supported planar bilayers are powerful tools that can be used to model the molecular interactions in an immunological synapse. To mimic the interactions between lymphocytes and antigen presenting cells, we use Ni2+-chelating lipids to anchor recombinant cell adhesion and MHC proteins to the upper leaflet of a bilayer with poly-histidine tags. Planar bilayers are generated by preparing lipid, treating the glass surfaces where the bilayer will form, and then forming the bilayer in a specialized chamber containing a flow-cell where the lymphocytes will be added. Then, bilayers are charged with Ni and his-tagged recombinant proteins are added. Finally, lymphocytes are injected into the flow cell and TIRF microscopy can be used to image synapse formation and the mechanisms that control T cell locomotion, sites of receptor sorting, and sites of receptor degradation.
Immunology, Issue 19, Annual Review, Immunological Synapse, Planar Lipid Bilayers, ICAM-1,
Single Molecule Methods for Monitoring Changes in Bilayer Elastic Properties
Institutions: Weill Cornell Medical College, Weill Cornell Medical College of Cornell University.
Membrane protein function is regulated by the cell membrane lipid composition. This regulation is due to a combination of specific lipid-protein interactions and more general lipid bilayer-protein interactions. These interactions are particularly important in pharmacological research, as many current pharmaceuticals on the market can alter the lipid bilayer material properties, which can lead to altered membrane protein function. The formation of gramicidin channels are dependent on conformational changes in gramicidin subunits which are in turn dependent on the properties of the lipid. Hence the gramicidin channel current is a reporter of altered properties of the bilayer due to certain compounds.
Cellular Biology, Issue 21, Springer Protocols, Membrane Biophysics, Gramicidin Channels, Artificial Bilayers, Bilayer Elastic Properties,
Preparation of Artificial Bilayers for Electrophysiology Experiments
Institutions: Weill Cornell Medical College of Cornell University.
Planar lipid bilayers, also called artificial lipid bilayers, allow you to study ion-conducting channels in a well-defined environment. These bilayers can be used for many different studies, such as the characterization of membrane-active peptides, the reconstitution of ion channels or investigations on how changes in lipid bilayer properties alter the function of bilayer-spanning channels. Here, we show how to form a planar bilayer and how to isolate small patches from the bilayer, and in a second video will also demonstrate a procedure for using gramicidin channels to determine changes in lipid bilayer elastic properties. We also demonstrate the individual steps needed to prepare the bilayer chamber, the electrodes and how to test that the bilayer is suitable for single-channel measurements.
Cellular Biology, Issue 20, Springer Protocols, Artificial Bilayers, Bilayer Patch Experiments, Lipid Bilayers, Bilayer Punch Electrodes, Electrophysiology