Understanding the mechanisms by which molecular motors coordinate their activities to transport vesicular cargoes within neurons requires the quantitative analysis of motor/cargo associations at the single vesicle level. The goal of this protocol is to use quantitative fluorescence microscopy to correlate (“map”) the position and directionality of movement of live cargo to the composition and relative amounts of motors associated with the same cargo. “Cargo mapping” consists of live imaging of fluorescently labeled cargoes moving in axons cultured on microfluidic devices, followed by chemical fixation during recording of live movement, and subsequent immunofluorescence (IF) staining of the exact same axonal regions with antibodies against motors. Colocalization between cargoes and their associated motors is assessed by assigning sub-pixel position coordinates to motor and cargo channels, by fitting Gaussian functions to the diffraction-limited point spread functions representing individual fluorescent point sources. Fixed cargo and motor images are subsequently superimposed to plots of cargo movement, to “map” them to their tracked trajectories. The strength of this protocol is the combination of live and IF data to record both the transport of vesicular cargoes in live cells and to determine the motors associated to these exact same vesicles. This technique overcomes previous challenges that use biochemical methods to determine the average motor composition of purified heterogeneous bulk vesicle populations, as these methods do not reveal compositions on single moving cargoes. Furthermore, this protocol can be adapted for the analysis of other transport and/or trafficking pathways in other cell types to correlate the movement of individual intracellular structures with their protein composition. Limitations of this protocol are the relatively low throughput due to low transfection efficiencies of cultured primary neurons and a limited field of view available for high-resolution imaging. Future applications could include methods to increase the number of neurons expressing fluorescently labeled cargoes.
14 Related JoVE Articles!
Use of Stopped-Flow Fluorescence and Labeled Nucleotides to Analyze the ATP Turnover Cycle of Kinesins
Institutions: University of Nottingham.
The kinesin superfamily of microtubule associated motor proteins share a characteristic motor domain which both hydrolyses ATP and binds microtubules. Kinesins display differences across the superfamily both in ATP turnover and in microtubule interaction. These differences tailor specific kinesins to various functions such as cargo transport, microtubule sliding, microtubule depolymerization and microtubule stabilization. To understand the mechanism of action of a kinesin it is important to understand how the chemical cycle of ATP turnover is coupled to the mechanical cycle of microtubule interaction. To dissect the ATP turnover cycle, one approach is to utilize fluorescently labeled nucleotides to visualize individual steps in the cycle. Determining the kinetics of each nucleotide transition in the ATP turnover cycle allows the rate-limiting step or steps for the complete cycle to be identified. For a kinesin, it is important to know the rate-limiting step, in the absence of microtubules, as this step is generally accelerated several thousand fold when the kinesin interacts with microtubules. The cycle in the absence of microtubules is then compared to that in the presence of microtubules to fully understand a kinesin’s ATP turnover cycle. The kinetics of individual nucleotide transitions are generally too fast to observe by manually mixing reactants, particularly in the presence of microtubules. A rapid mixing device, such as a stopped-flow fluorimeter, which allows kinetics to be observed on timescales of as little as a few milliseconds, can be used to monitor such transitions. Here, we describe protocols in which rapid mixing of reagents by stopped-flow is used in conjunction with fluorescently labeled nucleotides to dissect the ATP turnover cycle of a kinesin.
Chemistry, Issue 92, Kinesin, ATP turnover, mantATP, mantADP, stopped-flow fluorescence, microtubules, enzyme kinetics, nucleotide
Flexural Rigidity Measurements of Biopolymers Using Gliding Assays
Institutions: Lawrence University.
Microtubules are cytoskeletal polymers which play a role in cell division, cell mechanics, and intracellular transport. Each of these functions requires microtubules that are stiff and straight enough to span a significant fraction of the cell diameter. As a result, the microtubule persistence length, a measure of stiffness, has been actively studied for the past two decades1
. Nonetheless, open questions remain: short microtubules are 10-50 times less stiff than long microtubules2-4
, and even long microtubules have measured persistence lengths which vary by an order of magnitude5-9
Here, we present a method to measure microtubule persistence length. The method is based on a kinesin-driven microtubule gliding assay10
. By combining sparse fluorescent labeling of individual microtubules with single particle tracking of individual fluorophores attached to the microtubule, the gliding trajectories of single microtubules are tracked with nanometer-level precision. The persistence length of the trajectories is the same as the persistence length of the microtubule under the conditions used11
. An automated tracking routine is used to create microtubule trajectories from fluorophores attached to individual microtubules, and the persistence length of this trajectory is calculated using routines written in IDL.
This technique is rapidly implementable, and capable of measuring the persistence length of 100 microtubules in one day of experimentation. The method can be extended to measure persistence length under a variety of conditions, including persistence length as a function of length along microtubules. Moreover, the analysis routines used can be extended to myosin-based acting gliding assays, to measure the persistence length of actin filaments as well.
Biophysics, Issue 69, Bioengineering, Physics, Molecular Biology, Cellular Biology, microtubule, persistence length, flexural rigidity, gliding assay, mechanics, cytoskeleton, actin
Isolation and Purification of Kinesin from Drosophila Embryos
Institutions: University of California, Irvine.
Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a variety of neurodegenerative diseases, understanding microtubule based motor transport and its regulation will likely ultimately lead to improved therapeutic approaches. Kinesin-1 is a eukaryotic motor protein which moves in an anterograde (plus-end) direction along microtubules (MTs), powered by ATP hydrolysis. Here we report a detailed purification protocol to isolate active full length kinesin from Drosophila
embryos, thus allowing the combination of Drosophila
genetics with single-molecule biophysical studies. Starting with approximately 50 laying cups, with approximately 1000 females per cup, we carried out overnight collections. This provided approximately 10 ml of packed embryos. The embryos were bleach dechorionated (yielding approximately 9 grams of embryos), and then homogenized. After disruption, the homogenate was clarified using a low speed spin followed by a high speed centrifugation. The clarified supernatant was treated with GTP and taxol to polymerize MTs. Kinesin was immobilized on polymerized MTs by adding the ATP analog, 5'-adenylyl imidodiphosphate at room temperature. After kinesin binding, microtubules were sedimented via high speed centrifugation through a sucrose cushion. The microtubule pellet was then re-suspended, and this process was repeated. Finally, ATP was added to release the kinesin from the MTs. High speed centrifugation then spun down the MTs, leaving the kinesin in the supernatant. This kinesin was subjected to a centrifugal filtration using a 100 KD cut off filter for further purification, aliquoted, snap frozen in liquid nitrogen, and stored at -80 °C. SDS gel electrophoresis and western blotting was performed using the purified sample. The motor activity of purified samples before and after the final centrifugal filtration step was evaluated using an in vitro
single molecule microtubule assay. The kinesin fractions before and after the centrifugal filtration showed processivity as previously reported in literature. Further experiments are underway to evaluate the interaction between kinesin and other transport related proteins.
Developmental Biology, Issue 62, Drosophila, Kinesin, clarification, polymerization, sedimentation, microtubule
Cargo Loading onto Kinesin Powered Molecular Shuttles
Institutions: University of Florida, Columbia University.
Cells have evolved sophisticated molecular machinery, such as kinesin motor proteins and microtubule filaments, to support active intracellular transport of cargo. While kinesins tail domain binds to a variety of cargoes, kinesins head domains utilize the chemical energy stored in ATP molecules to step along the microtubule lattice. The long, stiff microtubules serve as tracks for long-distance intracellular transport.
These motors and filaments can also be employed in microfabricated synthetic environments as components of molecular shuttles 1
. In a frequently used design, kinesin motors are anchored to the track surface through their tails, and functionalized microtubules serve as cargo carrying elements, which are propelled by these motors. These shuttles can be loaded with cargo by utilizing the strong and selective binding between biotin and streptavidin. The key components (biotinylated tubulin, streptavidin, and biotinylated cargo) are commercially available.
Building on the classic inverted motility assay 2
, the construction of molecular shuttles is detailed here. Kinesin motor proteins are adsorbed to a surface precoated with casein; microtubules are polymerized from biotinylated tubulin, adhered to the kinesin and subsequently coated with rhodamine-labeled streptavidin. The ATP concentration is maintained at subsaturating concentration to achieve a microtubule gliding velocity optimal for loading cargo 3
. Finally, biotinylated fluorescein-labeled nanospheres are added as cargo. Nanospheres attach to microtubules as a result of collisions between gliding microtubules and nanospheres adhering to the surface.
The protocol can be readily modified to load a variety of cargoes such as biotinylated DNA4
, quantum dots 5
or a wide variety of antigens via biotinylated antibodies 4-6
Cellular Biology, Issue 45, motility assay, microtubules, kinesin, motor protein, molecular shuttle, nanobiotechnology
Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.
Institutions: Feinberg School of Medicine, Northwestern University, Basque Foundation for Science.
S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized microtubules. Organelles move along these processes by microtubule motors. Easy maintenance, high sensitivity to RNAi-mediated protein knock-down and efficient procedure for creating stable cell lines make Drosophila
S2 cells an ideal model system to study cargo transport by live imaging. The results obtained with S2 cells can be further applied to a more physiologically relevant system: axonal transport in primary neurons cultured from dissociated Drosophila
embryos. Cultured neurons grow long neurites filled with bundled microtubules, very similar to S2 processes. Like in S2 cells, organelles in cultured neurons can be visualized by either organelle-specific fluorescent dyes or by using fluorescent organelle markers encoded by DNA injected into early embryos or expressed in transgenic flies. Therefore, organelle transport can be easily recorded in neurons cultured on glass coverslips using living imaging. Here we describe procedures for culturing and visualizing cargo transport in Drosophila
S2 cells and primary neurons. We believe that these protocols make both systems accessible for labs studying cargo transport.
Cellular Biology, Issue 81, Drosophila melanogaster, cytoskeleton, S2 cells, primary neuron culture, microtubules, kinesin, dynein, fluorescence microscopy, live imaging
Preparation and Maintenance of Dorsal Root Ganglia Neurons in Compartmented Cultures
Institutions: Dana Farber Cancer Institute, Harvard Medical School.
Neurons extend axonal processes that are far removed from the cell body to innervate target tissues, where target-derived growth factors are required for neuronal survival and function. Neurotrophins are specifically required to maintain the survival and differentiation of innervating sensory neurons but the question of how these target-derived neurotrophins communicate to the cell body of innervating neurons has been an area of active research for over 30 years. The most commonly accepted model of how neurotrophin signals reach the cell body proposes that signaling endosomes carry this signal retrogradely along the axon. In order to study retrograde transport, a culture system was originally devised by Robert Campenot, in which cell bodies are isolated from their axons. The technique of preparing these compartmented chambers for culturing sensory neurons recapitulates the selective stimulation of neuron terminals that occurs in vivo following release of target-derived neurotrophins. Retrograde signaling events that require long-range microtubule dependent retrograde transport have important implications for the treatment of neurodegenerative disorders.
Neuroscience, Issue 20, campenot cultures, cell culture, dorsal root ganglia (DRG) neurons, neuronal culture
A Novel Stretching Platform for Applications in Cell and Tissue Mechanobiology
Institutions: University of Ottawa, University of Ottawa, University of Calgary, University of Ottawa, University of Ottawa.
Tools that allow the application of mechanical forces to cells and tissues or that can quantify the mechanical properties of biological tissues have contributed dramatically to the understanding of basic mechanobiology. These techniques have been extensively used to demonstrate how the onset and progression of various diseases are heavily influenced by mechanical cues. This article presents a multi-functional biaxial stretching (BAXS) platform that can either mechanically stimulate single cells or quantify the mechanical stiffness of tissues. The BAXS platform consists of four voice coil motors that can be controlled independently. Single cells can be cultured on a flexible substrate that can be attached to the motors allowing one to expose the cells to complex, dynamic, and spatially varying strain fields. Conversely, by incorporating a force load cell, one can also quantify the mechanical properties of primary tissues as they are exposed to deformation cycles. In both cases, a proper set of clamps must be designed and mounted to the BAXS platform motors in order to firmly hold the flexible substrate or the tissue of interest. The BAXS platform can be mounted on an inverted microscope to perform simultaneous transmitted light and/or fluorescence imaging to examine the structural or biochemical response of the sample during stretching experiments. This article provides experimental details of the design and usage of the BAXS platform and presents results for single cell and whole tissue studies. The BAXS platform was used to measure the deformation of nuclei in single mouse myoblast cells in response to substrate strain and to measure the stiffness of isolated mouse aortas. The BAXS platform is a versatile tool that can be combined with various optical microscopies in order to provide novel mechanobiological insights at the sub-cellular, cellular and whole tissue levels.
Bioengineering, Issue 88, cell stretching, tissue mechanics, nuclear mechanics, uniaxial, biaxial, anisotropic, mechanobiology
Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae
Institutions: University of Michigan, University of Michigan, University of Michigan, University of Michigan, University of Michigan.
Live imaging is an important technique for studying cell biological processes, however this can be challenging in live animals. The translucent cuticle of the Drosophila
larva makes it an attractive model organism for live imaging studies. However, an important challenge for live imaging techniques is to noninvasively immobilize and position an animal on the microscope. This protocol presents a simple and easy to use method for immobilizing and imaging Drosophila
larvae on a polydimethylsiloxane (PDMS) microfluidic device, which we call the 'larva chip'. The larva chip is comprised of a snug-fitting PDMS microchamber that is attached to a thin glass coverslip, which, upon application of a vacuum via a syringe, immobilizes the animal and brings ventral structures such as the nerve cord, segmental nerves, and body wall muscles, within close proximity to the coverslip. This allows for high-resolution imaging, and importantly, avoids the use of anesthetics and chemicals, which facilitates the study of a broad range of physiological processes. Since larvae recover easily from the immobilization, they can be readily subjected to multiple imaging sessions. This allows for longitudinal studies over time courses ranging from hours to days. This protocol describes step-by-step how to prepare the chip and how to utilize the chip for live imaging of neuronal events in 3rd
instar larvae. These events include the rapid transport of organelles in axons, calcium responses to injury, and time-lapse studies of the trafficking of photo-convertible proteins over long distances and time scales. Another application of the chip is to study regenerative and degenerative responses to axonal injury, so the second part of this protocol describes a new and simple procedure for injuring axons within peripheral nerves by a segmental nerve crush.
Bioengineering, Issue 84, Drosophila melanogaster, Live Imaging, Microfluidics, axonal injury, axonal degeneration, calcium imaging, photoconversion, laser microsurgery
Simple Microfluidic Devices for in vivo Imaging of C. elegans, Drosophila and Zebrafish
Institutions: NCBS-TIFR, TIFR.
Micro fabricated fluidic devices provide an accessible micro-environment for in vivo
studies on small organisms. Simple fabrication processes are available for microfluidic devices using soft lithography techniques 1-3
. Microfluidic devices have been used for sub-cellular imaging 4,5
, in vivo
laser microsurgery 2,6
and cellular imaging 4,7
. In vivo
imaging requires immobilization of organisms. This has been achieved using suction 5,8
, tapered channels 6,7,9
, deformable membranes 2-4,10
, suction with additional cooling 5
, anesthetic gas 11
, temperature sensitive gels 12
, cyanoacrylate glue 13
and anesthetics such as levamisole 14,15
. Commonly used anesthetics influence synaptic transmission 16,17
and are known to have detrimental effects on sub-cellular neuronal transport 4
. In this study we demonstrate a membrane based poly-dimethyl-siloxane (PDMS) device that allows anesthetic free immobilization of intact genetic model organisms such as Caenorhabditis elegans
larvae and zebrafish larvae. These model organisms are suitable for in vivo
studies in microfluidic devices because of their small diameters and optically transparent or translucent bodies. Body diameters range from ~10 μm to ~800 μm for early larval stages of C. elegans
and zebrafish larvae and require microfluidic devices of different sizes to achieve complete immobilization for high resolution time-lapse imaging. These organisms are immobilized using pressure applied by compressed nitrogen gas through a liquid column and imaged using an inverted microscope. Animals released from the trap return to normal locomotion within 10 min.
We demonstrate four applications of time-lapse imaging in C. elegans
namely, imaging mitochondrial transport in neurons, pre-synaptic vesicle transport in a transport-defective mutant, glutamate receptor transport and Q neuroblast cell division. Data obtained from such movies show that microfluidic immobilization is a useful and accurate means of acquiring in vivo
data of cellular and sub-cellular events when compared to anesthetized animals (Figure 1J
and 3C-F 4
Device dimensions were altered to allow time-lapse imaging of different stages of C. elegans
, first instar Drosophila
larvae and zebrafish larvae. Transport of vesicles marked with synaptotagmin tagged with GFP (syt.eGFP) in sensory neurons shows directed motion of synaptic vesicle markers expressed in cholinergic sensory neurons in intact first instar Drosophila
larvae. A similar device has been used to carry out time-lapse imaging of heartbeat in ~30 hr post fertilization (hpf) zebrafish larvae. These data show that the simple devices we have developed can be applied to a variety of model systems to study several cell biological and developmental phenomena in vivo
Bioengineering, Issue 67, Molecular Biology, Neuroscience, Microfluidics, C. elegans, Drosophila larvae, zebrafish larvae, anesthetic, pre-synaptic vesicle transport, dendritic transport of glutamate receptors, mitochondrial transport, synaptotagmin transport, heartbeat
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
The Cell-based L-Glutathione Protection Assays to Study Endocytosis and Recycling of Plasma Membrane Proteins
Institutions: Children's Hospital of Pittsburgh of UPMC, University of Pittsburgh School of Medicine.
Membrane trafficking involves transport of proteins from the plasma membrane to the cell interior (i.e.
endocytosis) followed by trafficking to lysosomes for degradation or to the plasma membrane for recycling. The cell based L-glutathione protection assays can be used to study endocytosis and recycling of protein receptors, channels, transporters, and adhesion molecules localized at the cell surface. The endocytic assay requires labeling of cell surface proteins with a cell membrane impermeable biotin containing a disulfide bond and the N-hydroxysuccinimide (NHS) ester at 4 ºC - a temperature at which membrane trafficking does not occur. Endocytosis of biotinylated plasma membrane proteins is induced by incubation at 37 ºC. Next, the temperature is decreased again to 4 ºC to stop endocytic trafficking and the disulfide bond in biotin covalently attached to proteins that have remained at the plasma membrane is reduced with L-glutathione. At this point, only proteins that were endocytosed remain protected from L-glutathione and thus remain biotinylated. After cell lysis, biotinylated proteins are isolated with streptavidin agarose, eluted from agarose, and the biotinylated protein of interest is detected by western blotting. During the recycling assay, after biotinylation cells are incubated at 37 °C to load endocytic vesicles with biotinylated proteins and the disulfide bond in biotin covalently attached to proteins remaining at the plasma membrane is reduced with L-glutathione at 4 ºC as in the endocytic assay. Next, cells are incubated again at 37 °C to allow biotinylated proteins from endocytic vesicles to recycle to the plasma membrane. Cells are then incubated at 4 ºC, and the disulfide bond in biotin attached to proteins that recycled to the plasma membranes is reduced with L-glutathione. The biotinylated proteins protected from L-glutathione are those that did not recycle to the plasma membrane.
Basic Protocol, Issue 82, Endocytosis, recycling, plasma membrane, cell surface, EZLink, Sulfo-NHS-SS-Biotin, L-Glutathione, GSH, thiol group, disulfide bond, epithelial cells, cell polarization
Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo
Institutions: University of California, Davis.
This protocol describes the use of the Drosophila melanogaster
syncytial embryo for studying mitosis1
has useful genetics with a sequenced genome, and it can be easily maintained and manipulated. Many mitotic mutants exist, and transgenic flies expressing functional fluorescently (e.g. GFP) - tagged mitotic proteins have been and are being generated. Targeted gene expression is possible using the GAL4/UAS system2
early embryo carries out multiple mitoses very rapidly (cell cycle duration, ≈10 min). It is well suited for imaging mitosis, because during cycles 10-13, nuclei divide rapidly and synchronously without intervening cytokinesis at the surface of the embryo in a single monolayer just underneath the cortex. These rapidly dividing nuclei probably use the same mitotic machinery as other cells, but they are optimized for speed; the checkpoint is generally believed to not be stringent, allowing the study of mitotic proteins whose absence would cause cell cycle arrest in cells with a strong checkpoint. Embryos expressing GFP labeled proteins or microinjected with fluorescently labeled proteins can be easily imaged to follow live dynamics (Fig. 1). In addition, embryos can be microinjected with function-blocking antibodies or inhibitors of specific proteins to study the effect of the loss or perturbation of their function3
. These reagents can diffuse throughout the embryo, reaching many spindles to produce a gradient of concentration of inhibitor, which in turn results in a gradient of defects comparable to an allelic series of mutants. Ideally, if the target protein is fluorescently labeled, the gradient of inhibition can be directly visualized4
. It is assumed that the strongest phenotype is comparable to the null phenotype, although it is hard to formally exclude the possibility that the antibodies may have dominant effects in rare instances, so rigorous controls and cautious interpretation must be applied. Further away from the injection site, protein function is only partially lost allowing other functions of the target protein to become evident.
Developmental Biology, Issue 31, mitosis, Drosophila melanogaster syncytial embryo, microinjection, protein inhibition
In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons
Institutions: SUNY-University at Buffalo.
Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different neurological diseases. Defects in this essential process can have detrimental effects on neuronal functioning and development. We have developed a dissection protocol that is designed to expose the Drosophila
larval segmental nerves to view axonal transport in real time. We have adapted this protocol for live imaging from the one published by Hurd and Saxton (1996) used for immunolocalizatin of larval segmental nerves. Careful dissection and proper buffer conditions are critical for maximizing the lifespan of the dissected larvae. When properly done, dissected larvae have shown robust vesicle transport for 2-3 hours under physiological conditions. We use the UAS-GAL4 method 1
to express GFP-tagged APP or synaptotagmin vesicles within a single axon or many axons in larval segmental nerves by using different neuronal GAL4 drivers. Other fluorescently tagged markers, for example mitochrondria (MitoTracker) or lysosomes (LysoTracker), can be also applied to the larvae before viewing. GFP-vesicle movement and particle movement can be viewed simultaneously using separate wavelengths.
Neuroscience, Issue 44, Live imaging, Axonal transport, GFP-tagged vesicles
Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons
Institutions: Simon Fraser University.
Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images. Vital fluorescent probes, particularly green fluorescent protein (GFP) have revolutionized cell biology stemming from the ability to label specific intracellular compartments and cellular structures. For example, the live imaging of GFP (and its spectral variants) chimeras have allowed for a dynamic
analysis of the cytoskeleton, organelle transport, and membrane dynamics in a multitude of organisms and cell types [1-3]. Although live imaging has become prevalent, this approach still poses many technical challenges, particularly in primary cultured neurons. One challenge is the expression of GFP-tagged proteins in post-mitotic neurons; the other is the ability to capture fluorescent images while minimizing phototoxicity, photobleaching,
and maintaining general cell health. Here we provide a protocol that describes a lipid-based transfection method that yields a relatively low transfection rate (~0.5%), however is ideal for the imaging of fully polarized neurons. A low transfection rate is essential so that single axons and dendrites can be characterized as to their orientation to the cell body to confirm directionality of transport, i.e., anterograde v. retrograde. Our approach to imaging
GFP expressing neurons relies on a standard wide-field fluorescent microscope outfitted with a CCD camera, image capture software, and a heated imaging chamber. We have imaged a wide variety of organelles or structures, for example, dense-core vesicles, mitochondria, growth cones, and actin without any special optics or excitation requirements other than a fluorescent light source. Additionally, spectrally-distinct, fluorescently
labeled proteins, e.g., GFP and dsRed-tagged proteins, can be visualized near simultaneously to characterize co-transport or other coordinated cellular events. The imaging approach described here is flexible for a variety of imaging applications and can be adopted by a laboratory for relatively little cost provided a microscope is available.
Neuroscience, Issue 27, Live cell imaging, intracellular transport, membrane-bound organelles, green fluorescent protein, hippocampal neurons, transfection, fluorescence microscopy