Luminescence Resonance Energy Transfer, or LRET, is a powerful technique used to measure distances between two sites in proteins within the distance range of 10-100 Å. By measuring the distances under various ligated conditions, conformational changes of the protein can be easily assessed. With LRET, a lanthanide, most often chelated terbium, is used as the donor fluorophore, affording advantages such as a longer donor-only emission lifetime, the flexibility to use multiple acceptor fluorophores, and the opportunity to detect sensitized acceptor emission as an easy way to measure energy transfer without the risk of also detecting donor-only signal. Here, we describe a method to use LRET on membrane proteins expressed and assayed on the surface of intact mammalian cells. We introduce a protease cleavage site between the LRET fluorophore pair. After obtaining the original LRET signal, cleavage at that site removes the specific LRET signal from the protein of interest allowing us to quantitatively subtract the background signal that remains after cleavage. This method allows for more physiologically relevant measurements to be made without the need for purification of protein.
22 Related JoVE Articles!
Fluorescence detection methods for microfluidic droplet platforms
Institutions: Imperial College London , Chungbuk National University, Institute for Chemical and Bioengineering, ETH Zurich.
The development of microfluidic platforms for performing chemistry and biology has in large part been driven by a range of potential benefits that accompany system miniaturisation. Advantages include the ability to efficiently process nano- to femoto- liter volumes of sample, facile integration of functional components, an intrinsic predisposition towards large-scale multiplexing, enhanced analytical throughput, improved control and reduced instrumental footprints.1
In recent years much interest has focussed on the development of droplet-based (or segmented flow) microfluidic systems and their potential as platforms in high-throughput experimentation.2-4
Here water-in-oil emulsions are made to spontaneously form in microfluidic channels as a result of capillary instabilities between the two immiscible phases. Importantly, microdroplets of precisely defined volumes and compositions can be generated at frequencies of several kHz. Furthermore, by encapsulating reagents of interest within isolated compartments separated by a continuous immiscible phase, both sample cross-talk and dispersion (diffusion- and Taylor-based) can be eliminated, which leads to minimal cross-contamination and the ability to time analytical processes with great accuracy. Additionally, since there is no contact between the contents of the droplets and the channel walls (which are wetted by the continuous phase) absorption and loss of reagents on the channel walls is prevented.
Once droplets of this kind have been generated and processed, it is necessary to extract the required analytical information. In this respect the detection method of choice should be rapid, provide high-sensitivity and low limits of detection, be applicable to a range of molecular species, be non-destructive and be able to be integrated with microfluidic devices in a facile manner. To address this need we have developed a suite of experimental tools and protocols that enable the extraction of large amounts of photophysical information from small-volume environments, and are applicable to the analysis of a wide range of physical, chemical and biological parameters. Herein two examples of these methods are presented and applied to the detection of single cells and the mapping of mixing processes inside picoliter-volume droplets. We report the entire experimental process including microfluidic chip fabrication, the optical setup and the process of droplet generation and detection.
Bioengineering, Issue 58, Droplet Microfluidics, Single Cell Assays, Single Molecule Assays, Fluorescence Spectroscopy, Fluorescence Lifetime Imaging
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
Analysis of Targeted Viral Protein Nanoparticles Delivered to HER2+ Tumors
Institutions: University of Southern California, Cedars-Sinai Medical Center, University of California, Los Angeles.
The HER2+ tumor-targeted nanoparticle, HerDox, exhibits tumor-preferential accumulation and tumor-growth ablation in an animal model of HER2+ cancer. HerDox is formed by non-covalent self-assembly of a tumor targeted cell penetration protein with the chemotherapy agent, doxorubicin, via a small nucleic acid linker. A combination of electrophilic, intercalation, and oligomerization interactions facilitate self-assembly into round 10-20 nm particles. HerDox exhibits stability in blood as well as in extended storage at different temperatures. Systemic delivery of HerDox in tumor-bearing mice results in tumor-cell death with no detectable adverse effects to non-tumor tissue, including the heart and liver (which undergo marked damage by untargeted doxorubicin). HER2 elevation facilitates targeting to cells expressing the human epidermal growth factor receptor, hence tumors displaying elevated HER2 levels exhibit greater accumulation of HerDox compared to cells expressing lower levels, both in vitro
and in vivo
. Fluorescence intensity imaging combined with in situ
confocal and spectral analysis has allowed us to verify in vivo
tumor targeting and tumor cell penetration of HerDox after systemic delivery. Here we detail our methods for assessing tumor targeting via multimode imaging after systemic delivery.
Biomedical Engineering, Issue 76, Cancer Biology, Medicine, Bioengineering, Molecular Biology, Cellular Biology, Biochemistry, Nanotechnology, Nanomedicine, Drug Delivery Systems, Molecular Imaging, optical imaging devices (design and techniques), HerDox, Nanoparticle, Tumor, Targeting, Self-Assembly, Doxorubicin, Human Epidermal Growth Factor, HER, HER2+, Receptor, mice, animal model, tumors, imaging
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)
Examining the Conformational Dynamics of Membrane Proteins in situ with Site-directed Fluorescence Labeling
Institutions: Worcester Polytechnic Institute.
Two electrode voltage clamp electrophysiology (TEVC) is a powerful tool to investigate the mechanism of ion transport1 for a wide variety of membrane proteins including ion channels2
, ion pumps3
, and transporters4
. Recent developments have combined site-specific fluorophore labeling alongside TEVC to
concurrently examine the conformational dynamics at specific residues and function of these proteins on the surface of single cells.
We will describe a method to study the conformational dynamics of membrane proteins by simultaneously monitoring fluorescence and current changes using voltage-clamp fluorometry. This approach can be used to examine the molecular motion of membrane proteins site-specifically following cysteine replacement and site-directed fluorophore labeling5,6
. Furthermore, this method provides an approach to determine distance constraints between specific residues7,8
This is achieved by selectively attaching donor and acceptor fluorophores to two mutated cysteine residues of interest.
In brief, these experiments are performed following functional expression of the desired protein on the surface of Xenopus leavis
oocytes. The large surface area of these oocytes enables facile functional measurements and a robust fluorescence signal5
. It is also possible to readily change the extracellular conditions such as pH, ligand or cations/anions, which can provide further information on the mechanism of membrane proteins4
. Finally, recent developments
have also enabled the manipulation of select internal ions following co-expression with a second protein9
Our protocol is described in multiple parts. First, cysteine scanning mutagenesis proceeded by fluorophore labeling is completed at residues located at the interface of the transmembrane and extracellular domains. Subsequent experiments are designed to identify residues which demonstrate large changes in fluorescence intensity (<5%)3
upon a conformational change of the protein. Second, these changes in fluorescence intensity are compared to the kinetic parameters of
the membrane protein in order to correlate the conformational dynamics to the function of the protein10
. This enables a rigorous biophysical analysis of the molecular motion of the target protein. Lastly, two residues of the holoenzyme can be labeled with a donor and acceptor fluorophore in order to determine distance constraints using donor photodestruction methods. It is also possible to monitor the relative movement of protein subunits following labeling with a donor and acceptor fluorophore.
Cellular Biology, Issue 51, membrane protein, two electrode voltage-clamp, biophysics, site-specific fluorophore labeling, microscopy, conformational dynamics
In vivo Neuronal Calcium Imaging in C. elegans
Institutions: Boston University School of Medicine, Boston University Photonics Center.
The nematode worm C. elegans
is an ideal model organism for relatively simple, low cost neuronal imaging in vivo
. Its small transparent body and simple, well-characterized nervous system allows identification and fluorescence imaging of any neuron within the intact animal. Simple immobilization techniques with minimal impact on the animal's physiology allow extended time-lapse imaging. The development of genetically-encoded calcium sensitive fluorophores such as cameleon 1
and GCaMP 2
allow in vivo
imaging of neuronal calcium relating both cell physiology and neuronal activity. Numerous transgenic strains expressing these fluorophores in specific neurons are readily available or can be constructed using well-established techniques. Here, we describe detailed procedures for measuring calcium dynamics within a single neuron in vivo
using both GCaMP and cameleon. We discuss advantages and disadvantages of both as well as various methods of sample preparation (animal immobilization) and image analysis. Finally, we present results from two experiments: 1) Using GCaMP to measure the sensory response of a specific neuron to an external electrical field and 2) Using cameleon to measure the physiological calcium response of a neuron to traumatic laser damage. Calcium imaging techniques such as these are used extensively in C. elegans
and have been extended to measurements in freely moving animals, multiple neurons simultaneously and comparison across genetic backgrounds. C. elegans
presents a robust and flexible system for in vivo
neuronal imaging with advantages over other model systems in technical simplicity and cost.
Developmental Biology, Issue 74, Physiology, Biophysics, Neurobiology, Cellular Biology, Molecular Biology, Anatomy, Developmental Biology, Biomedical Engineering, Medicine, Caenorhabditis elegans, C. elegans, Microscopy, Fluorescence, Neurosciences, calcium imaging, genetically encoded calcium indicators, cameleon, GCaMP, neuronal activity, time-lapse imaging, laser ablation, optical neurophysiology, neurophysiology, neurons, animal model
Fluorescence Imaging with One-nanometer Accuracy (FIONA)
Institutions: University of Illinois at Urbana-Champaign, University of Illinois at Urbana-Champaign, University of Illinois at Urbana-Champaign.
Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in the x-y plane. Here a summary of the FIONA technique is reported and examples of research that have been performed using FIONA are briefly described. First, how to set up the required equipment for FIONA experiments, i.e.
, a total internal reflection fluorescence microscopy (TIRFM), with details on aligning the optics, is described. Then how to carry out a simple FIONA experiment on localizing immobilized Cy3-DNA single molecules using appropriate protocols, followed by the use of FIONA to measure the 36 nm step size of a single truncated myosin Va motor labeled with a quantum dot, is illustrated. Lastly, recent effort to extend the application of FIONA to thick samples is reported. It is shown that, using a water immersion objective and quantum dots soaked deep in sol-gels and rabbit eye corneas (>200 µm), localization precision of 2-3 nm can be achieved.
Molecular Biology, Issue 91, FIONA, fluorescence imaging, nanometer precision, myosin walking, thick tissue
High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy
Institutions: University of Würzburg, Germany.
Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g
., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e
-adrenergic and γ-aminobutyric acid type B (GABAB
) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.
Bioengineering, Issue 89, pharmacology, microscopy, receptor, live-cell imaging, single-molecule, total internal reflection fluorescence, tracking, dimerization, protein-protein interactions
Monitoring Intraspecies Competition in a Bacterial Cell Population by Cocultivation of Fluorescently Labelled Strains
Institutions: Georg-August University.
Many microorganisms such as bacteria proliferate extremely fast and the populations may reach high cell densities. Small fractions of cells in a population always have accumulated mutations that are either detrimental or beneficial for the cell. If the fitness effect of a mutation provides the subpopulation with a strong selective growth advantage, the individuals of this subpopulation may rapidly outcompete and even completely eliminate their immediate fellows. Thus, small genetic changes and selection-driven accumulation of cells that have acquired beneficial mutations may lead to a complete shift of the genotype of a cell population. Here we present a procedure to monitor the rapid clonal expansion and elimination of beneficial and detrimental mutations, respectively, in a bacterial cell population over time by cocultivation of fluorescently labeled individuals of the Gram-positive model bacterium Bacillus subtilis
. The method is easy to perform and very illustrative to display intraspecies competition among the individuals in a bacterial cell population.
Cellular Biology, Issue 83, Bacillus subtilis, evolution, adaptation, selective pressure, beneficial mutation, intraspecies competition, fluorophore-labelling, Fluorescence Microscopy
Real-time Monitoring of Ligand-receptor Interactions with Fluorescence Resonance Energy Transfer
Institutions: Southern Illinois University.
FRET is a process whereby energy is non-radiatively transferred from an excited donor molecule to a ground-state acceptor molecule through long-range dipole-dipole interactions1
. In the present sensing assay, we utilize an interesting property of PDA: blue-shift in the UV-Vis electronic absorption spectrum of PDA (Figure 1
) after an analyte interacts with receptors attached to PDA2,3,4,7
. This shift in the PDA absorption spectrum provides changes in the spectral overlap (J
) between PDA (acceptor) and rhodamine (donor) that leads to changes in the FRET efficiency. Thus, the interactions between analyte (ligand) and receptors are detected through FRET between donor fluorophores and PDA. In particular, we show the sensing of a model protein molecule streptavidin. We also demonstrate the covalent-binding of bovine serum albumin (BSA) to the liposome surface with FRET mechanism. These interactions between the bilayer liposomes and protein molecules can be sensed in real-time. The proposed method is a general method for sensing small chemical and large biochemical molecules. Since fluorescence is intrinsically more sensitive than colorimetry, the detection limit of the assay can be in sub-nanomolar range or lower8
. Further, PDA can act as a universal acceptor in FRET, which means that multiple sensors can be developed with PDA (acceptor) functionalized with donors and different receptors attached on the surface of PDA liposomes.
Biochemistry, Issue 66, Molecular Biology, Chemistry, Physics, Fluorescence Resonance Energy Transfer (FRET), Polydiacetylene (PDA), Biosensor, Liposome, Sensing
Synthesis and Calibration of Phosphorescent Nanoprobes for Oxygen Imaging in Biological Systems
Institutions: University of Pennsylvania .
Oxygen measurement by phosphorescence quenching [1, 2] consists of the following steps: 1) the probe is delivered into the medium of interest (e.g. blood or interstitial fluid); 2) the object is illuminated with light of appropriate wavelength in order to excite the probe into its triplet state; 3) the emitted phosphorescence is collected, and its time course is analyzed to yield the phosphorescence lifetime, which is converted into the oxygen concentration (or partial pressure, pO2
). The probe must not interact with the biological environment and in some cases to be 4) excreted from the medium upon the measurement completion. Each of these steps imposes requirements on the molecular design of the phosphorescent probes, which constitute the only invasive component of the measurement protocol. Here we review the design of dendritic phosphorescent nanosensors for oxygen measurements in biological systems. The probes consist of Pt or Pd porphyrin-based polyarylglycine (AG) dendrimers, modified peripherally with polyethylene glycol (PEG's) residues. For effective two-photon excitation, termini of the dendrimers may be modified with two-photon antenna chromophores, which capture the excitation energy and channel it to the triplet cores of the probes via intramolecular FRET (Förster Resonance Energy Transfer). We describe the key photophysical properties of the probes and present detailed calibration protocols.
Cellular Biology, Issue 37, oxygen, phosphorescence, porphyrin, dendrimer, imaging, nanosensor, two-photon
Fluorescence Lifetime Imaging of Molecular Rotors in Living Cells
Institutions: King's College London, Imperial College London , PhotoBiotics Ltd.
Diffusion is often an important rate-determining step in chemical reactions or biological processes and plays a role in a wide range of intracellular events. Viscosity is one of the key parameters affecting the diffusion of molecules and proteins, and changes in viscosity have been linked to disease and malfunction at the cellular level.1-3
While methods to measure the bulk viscosity are well developed, imaging microviscosity remains a challenge. Viscosity maps of microscopic objects, such as single cells, have until recently been hard to obtain. Mapping viscosity with fluorescence techniques is advantageous because, similar to other optical techniques, it is minimally invasive, non-destructive and can be applied to living cells and tissues.
Fluorescent molecular rotors exhibit fluorescence lifetimes and quantum yields which are a function of the viscosity of their microenvironment.4,5
Intramolecular twisting or rotation leads to non-radiative decay from the excited state back to the ground state. A viscous environment slows this rotation or twisting, restricting access to this non-radiative decay pathway. This leads to an increase in the fluorescence quantum yield and the fluorescence lifetime. Fluorescence Lifetime Imaging (FLIM) of modified hydrophobic BODIPY dyes that act as fluorescent molecular rotors show that the fluorescence lifetime of these probes is a function of the microviscosity of their environment.6-8
A logarithmic plot of the fluorescence lifetime versus the solvent viscosity yields a straight line that obeys the Förster Hoffman equation.9
This plot also serves as a calibration graph to convert fluorescence lifetime into viscosity.
Following incubation of living cells with the modified BODIPY fluorescent molecular rotor, a punctate dye distribution is observed in the fluorescence images. The viscosity value obtained in the puncta in live cells is around 100 times higher than that of water and of cellular cytoplasm.6,7
Time-resolved fluorescence anisotropy measurements yield rotational correlation times in agreement with these large microviscosity values. Mapping the fluorescence lifetime is independent of the fluorescence intensity, and thus allows the separation of probe concentration and viscosity effects.
In summary, we have developed a practical and versatile approach to map the microviscosity in cells based on FLIM of fluorescent molecular rotors.
Bioengineering, Issue 60, fluorescence, microscopy, FLIM, fluorescent molecular rotors
Synthesis, Cellular Delivery and In vivo Application of Dendrimer-based pH Sensors
Institutions: Eindhoven University of Technology & NEST, Scuola Normale Superiore and Istituto Nanoscienze-CNR, Center for Nanotechnology Innovation, NEST, Scuola Normale Superiore and Istituto Nanoscienze-CNR, Center for Nanotechnology Innovation.
The development of fluorescent indicators represented a revolution for life sciences. Genetically encoded and synthetic fluorophores with sensing abilities allowed the visualization of biologically relevant species with high spatial and temporal resolution. Synthetic dyes are of particular interest thanks to their high tunability and the wide range of measureable analytes. However, these molecules suffer several limitations related to small molecule behavior (poor solubility, difficulties in targeting, often no ratiometric imaging allowed). In this work we introduce the development of dendrimer-based sensors and present a procedure for pH measurement in vitro
, in living cells and in vivo
. We choose dendrimers as ideal platform for our sensors for their many desirable properties (monodispersity, tunable properties, multivalency) that made them a widely used scaffold for several biomedical devices. The conjugation of fluorescent pH indicators to the dendrimer scaffold led to an enhancement of their sensing performances. In particular dendrimers exhibit reduced cell leakage, improved intracellular targeting and allow ratiometric measurements. These novel sensors were successfully employed to measure pH in living HeLa cells and in vivo
in mouse brain.
Chemistry, Issue 79, Investigative Techniques, Chemistry and Materials (General), dendrimer, fluorescence, sensors, pH, delivery, confocal
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
Test Samples for Optimizing STORM Super-Resolution Microscopy
Institutions: National Physical Laboratory.
STORM is a recently developed super-resolution microscopy technique with up to 10 times better resolution than standard fluorescence microscopy techniques. However, as the image is acquired in a very different way than normal, by building up an image molecule-by-molecule, there are some significant challenges for users in trying to optimize their image acquisition. In order to aid this process and gain more insight into how STORM works we present the preparation of 3 test samples and the methodology of acquiring and processing STORM super-resolution images with typical resolutions of between 30-50 nm. By combining the test samples with the use of the freely available rainSTORM processing software it is possible to obtain a great deal of information about image quality and resolution. Using these metrics it is then possible to optimize the imaging procedure from the optics, to sample preparation, dye choice, buffer conditions, and image acquisition settings. We also show examples of some common problems that result in poor image quality, such as lateral drift, where the sample moves during image acquisition and density related problems resulting in the 'mislocalization' phenomenon.
Molecular Biology, Issue 79, Genetics, Bioengineering, Biomedical Engineering, Biophysics, Basic Protocols, HeLa Cells, Actin Cytoskeleton, Coated Vesicles, Receptor, Epidermal Growth Factor, Actins, Fluorescence, Endocytosis, Microscopy, STORM, super-resolution microscopy, nanoscopy, cell biology, fluorescence microscopy, test samples, resolution, actin filaments, fiducial markers, epidermal growth factor, cell, imaging
Preparation of Segmented Microtubules to Study Motions Driven by the Disassembling Microtubule Ends
Institutions: Russian Academy of Sciences, Federal Research Center of Pediatric Hematology, Oncology and Immunology, Moscow, Russia, University of Pennsylvania.
Microtubule depolymerization can provide force to transport different protein complexes and protein-coated beads in vitro
. The underlying mechanisms are thought to play a vital role in the microtubule-dependent chromosome motions during cell division, but the relevant proteins and their exact roles are ill-defined. Thus, there is a growing need to develop assays with which to study such motility in vitro
using purified components and defined biochemical milieu. Microtubules, however, are inherently unstable polymers; their switching between growth and shortening is stochastic and difficult to control. The protocols we describe here take advantage of the segmented microtubules that are made with the photoablatable stabilizing caps. Depolymerization of such segmented microtubules can be triggered with high temporal and spatial resolution, thereby assisting studies of motility at the disassembling microtubule ends. This technique can be used to carry out a quantitative analysis of the number of molecules in the fluorescently-labeled protein complexes, which move processively with dynamic microtubule ends. To optimize a signal-to-noise ratio in this and other quantitative fluorescent assays, coverslips should be treated to reduce nonspecific absorption of soluble fluorescently-labeled proteins. Detailed protocols are provided to take into account the unevenness of fluorescent illumination, and determine the intensity of a single fluorophore using equidistant Gaussian fit. Finally, we describe the use of segmented microtubules to study microtubule-dependent motions of the protein-coated microbeads, providing insights into the ability of different motor and nonmotor proteins to couple microtubule depolymerization to processive cargo motion.
Basic Protocol, Issue 85, microscopy flow chamber, single-molecule fluorescence, laser trap, microtubule-binding protein, microtubule-dependent motor, microtubule tip-tracking
Optimized Staining and Proliferation Modeling Methods for Cell Division Monitoring using Cell Tracking Dyes
Institutions: Roswell Park Cancer Institute, University of Pennsylvania , SciGro, Inc., University of Pennsylvania .
Fluorescent cell tracking dyes, in combination with flow and image cytometry, are powerful tools with which to study the interactions and fates of different cell types in vitro
and in vivo
Although there are literally thousands of publications using such dyes, some of the most commonly encountered cell tracking applications include monitoring of:
stem and progenitor cell quiescence, proliferation and/or differentiation6-8
antigen-driven membrane transfer9
and/or precursor cell proliferation3,4,10-18
immune regulatory and effector cell function1,18-21
Commercially available cell tracking dyes vary widely in their chemistries and fluorescence properties but the great majority fall into one of two classes based on their mechanism of cell labeling. "Membrane dyes", typified by PKH26, are highly lipophilic dyes that partition stably but non-covalently into cell membranes1,2,11
. "Protein dyes", typified by CFSE, are amino-reactive dyes that form stable covalent bonds with cell proteins4,16,18
. Each class has its own advantages and limitations. The key to their successful use, particularly in multicolor studies where multiple dyes are used to track different cell types, is therefore to understand the critical issues enabling optimal use of each class2-4,16,18,24
The protocols included here highlight three common causes of poor or variable results when using cell-tracking dyes. These are:
Failure to achieve bright, uniform, reproducible labeling
. This is a necessary starting point for any cell tracking study but requires attention to different variables when using membrane dyes than when using protein dyes or equilibrium binding reagents such as antibodies.
Suboptimal fluorochrome combinations and/or failure to include critical compensation controls
. Tracking dye fluorescence is typically 102
times brighter than antibody fluorescence. It is therefore essential to verify that the presence of tracking dye does not compromise the ability to detect other probes being used.
Failure to obtain a good fit with peak modeling software
. Such software allows quantitative comparison of proliferative responses across different populations or stimuli based on precursor frequency or other metrics. Obtaining a good fit, however, requires exclusion of dead/dying cells that can distort dye dilution profiles and matching of the assumptions underlying the model with characteristics of the observed dye dilution profile.
Examples given here illustrate how these variables can affect results when using membrane and/or protein dyes to monitor cell proliferation.
Cellular Biology, Issue 70, Molecular Biology, Cell tracking, PKH26, CFSE, membrane dyes, dye dilution, proliferation modeling, lymphocytes
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
Real-time Imaging of Single Engineered RNA Transcripts in Living Cells Using Ratiometric Bimolecular Beacons
Institutions: University of Pennsylvania, Integrated DNA Technologies, Inc..
The growing realization that both the temporal and spatial regulation of gene expression can have important consequences on cell function has led to the development of diverse techniques to visualize individual RNA transcripts in single living cells. One promising technique that has recently been described utilizes an oligonucleotide-based optical probe, ratiometric bimolecular beacon (RBMB), to detect RNA transcripts that were engineered to contain at least four tandem repeats of the RBMB target sequence in the 3’-untranslated region. RBMBs are specifically designed to emit a bright fluorescent signal upon hybridization to complementary RNA, but otherwise remain quenched. The use of a synthetic probe in this approach allows photostable, red-shifted, and highly emissive organic dyes to be used for imaging. Binding of multiple RBMBs to the engineered RNA transcripts results in discrete fluorescence spots when viewed under a wide-field fluorescent microscope. Consequently, the movement of individual RNA transcripts can be readily visualized in real-time by taking a time series of fluorescent images. Here we describe the preparation and purification of RBMBs, delivery into cells by microporation and live-cell imaging of single RNA transcripts.
Genetics, Issue 90, RNA, imaging, single molecule, fluorescence, living cell
A Step Beyond BRET: Fluorescence by Unbound Excitation from Luminescence (FUEL)
Institutions: Institut Pasteur, Stanford School of Medicine, Institut d'Imagerie Biomédicale, Vanderbilt School of Medicine, The Walter & Eliza Hall Institute of Medical Research, Institut Pasteur, Institut Pasteur.
Fluorescence by Unbound Excitation from Luminescence (FUEL) is a radiative excitation-emission process that produces increased signal and contrast enhancement in vitro
and in vivo
. FUEL shares many of the same underlying principles as Bioluminescence Resonance Energy Transfer (BRET), yet greatly differs in the acceptable working distances between the luminescent source and the fluorescent entity. While BRET is effectively limited to a maximum of 2 times the Förster radius, commonly less than 14 nm, FUEL can occur at distances up to µm or even cm in the absence of an optical absorber. Here we expand upon the foundation and applicability of FUEL by reviewing the relevant principles behind the phenomenon and demonstrate its compatibility with a wide variety of fluorophores and fluorescent nanoparticles. Further, the utility of antibody-targeted FUEL is explored. The examples shown here provide evidence that FUEL can be utilized for applications where BRET is not possible, filling the spatial void that exists between BRET and traditional whole animal imaging.
Bioengineering, Issue 87, Biochemical Phenomena, Biochemical Processes, Energy Transfer, Fluorescence Resonance Energy Transfer (FRET), FUEL, BRET, CRET, Förster, bioluminescence, In vivo
A Microplate Assay to Assess Chemical Effects on RBL-2H3 Mast Cell Degranulation: Effects of Triclosan without Use of an Organic Solvent
Institutions: University of Maine, Orono, University of Maine, Orono.
Mast cells play important roles in allergic disease and immune defense against parasites. Once activated (e.g.
by an allergen), they degranulate, a process that results in the exocytosis of allergic mediators. Modulation of mast cell degranulation by drugs and toxicants may have positive or adverse effects on human health. Mast cell function has been dissected in detail with the use of rat basophilic leukemia mast cells (RBL-2H3), a widely accepted model of human mucosal mast cells3-5
. Mast cell granule component and the allergic mediator β-hexosaminidase, which is released linearly in tandem with histamine from mast cells6
, can easily and reliably be measured through reaction with a fluorogenic substrate, yielding measurable fluorescence intensity in a microplate assay that is amenable to high-throughput studies1
. Originally published by Naal et al.1
, we have adapted this degranulation assay for the screening of drugs and toxicants and demonstrate its use here.
Triclosan is a broad-spectrum antibacterial agent that is present in many consumer products and has been found to be a therapeutic aid in human allergic skin disease7-11
, although the mechanism for this effect is unknown. Here we demonstrate an assay for the effect of triclosan on mast cell degranulation. We recently showed that triclosan strongly affects mast cell function2
. In an effort to avoid use of an organic solvent, triclosan is dissolved directly into aqueous buffer with heat and stirring, and resultant concentration is confirmed using UV-Vis spectrophotometry (using ε280
= 4,200 L/M/cm)12
. This protocol has the potential to be used with a variety of chemicals to determine their effects on mast cell degranulation, and more broadly, their allergic potential.
Immunology, Issue 81, mast cell, basophil, degranulation, RBL-2H3, triclosan, irgasan, antibacterial, β-hexosaminidase, allergy, Asthma, toxicants, ionophore, antigen, fluorescence, microplate, UV-Vis
Visualizing Protein-DNA Interactions in Live Bacterial Cells Using Photoactivated Single-molecule Tracking
Institutions: University of Oxford, University of Oxford.
Protein-DNA interactions are at the heart of many fundamental cellular processes. For example, DNA replication, transcription, repair, and chromosome organization are governed by DNA-binding proteins that recognize specific DNA structures or sequences. In vitro
experiments have helped to generate detailed models for the function of many types of DNA-binding proteins, yet, the exact mechanisms of these processes and their organization in the complex environment of the living cell remain far less understood. We recently introduced a method for quantifying DNA-repair activities in live Escherichia coli
cells using Photoactivated Localization Microscopy (PALM) combined with single-molecule tracking. Our general approach identifies individual DNA-binding events by the change in the mobility of a single protein upon association with the chromosome. The fraction of bound molecules provides a direct quantitative measure for the protein activity and abundance of substrates or binding sites at the single-cell level. Here, we describe the concept of the method and demonstrate sample preparation, data acquisition, and data analysis procedures.
Immunology, Issue 85, Super-resolution microscopy, single-particle tracking, Live-cell imaging, DNA-binding proteins, DNA repair, molecular diffusion