We thank Harald Hutter and Helena Decker for their careful reading of this manuscript. We also thank Reg Sidhu from Leica Microsystems for his technical expertise. This research was supported by the National Sciences and Engineering Council of Canada, Award #327100-06, and the Simon Fraser University Faculty of Science.

Part 1: Transfection of neurons using Lipofectamine 2000 (Invitrogen)
Equipment set-up:
Rat or mouse hippocampal neurons are cultured according to Kaech and Banker, 2006 [4]. The typical cell density used for transfections is 250,000 cells/6-cm dish. Required reagents and equipment include 50 mM kynurenic acid, regular benchtop tube holder, a benchtop cooler (best to keep Lipofectamine reagent cold), 
Lipofectamine reagent, MEM, microfuge tubes, micropipetters, and ...

<ol>
	<li>Kulic, I. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+microtubule+movement+in+bidirectional+organelle+transport.">The role of microtubule movement in bidirectional organelle transport.</a> Proc Natl Acad Sci U S A. 105 (29), 10011-10016 (2008).</li><li>Jacobson, C., Schnapp, B., Banker, G. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+change+in+the+selective+translocation+of+the+Kinesin-1+motor+domain+marks+the+initial+specification+of+the+axon.">A change in the selective translocation of the Kinesin-1 motor domain marks the initial specification of the axon.</a> Neuron. 49 (6), 797-804 (2006).</li><li>Barkus, R. V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+an+Axonal+Kinesin-3+Motor+for+Fast+Anterograde+Vesicle+Transport+that+Facilitates+Retrograde+Transport+of+Neuropeptides.">Identification of an Axonal Kinesin-3 Motor for Fast Anterograde Vesicle Transport that Facilitates Retrograde Transport of Neuropeptides.</a> Mol Biol Cell. 19 (1), 274-283 (2008).</li><li>Kaech, S., Banker, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Culturing+hippocampal+neurons.">Culturing hippocampal neurons.</a> Nat Protoc. 1 (5), 2406-2415 (2006).</li></ol>

Live cell imaging is a challenging, but powerful technique for the direct observation of organelle transport in cultured neurons. Difficulties can arise upstream of the procedure with poor neuron health due to culture complications. Therefore, cell health (for examples see ref. 4) should be assessed before and after transfection by observing the coverslips while in growth medium using a tissue culture light microscope. The process of transferring neuron-containing coverslips to the imaging chamber can be stressful to...

A correction was made to <a href="http://www.jove.com/details.php?ID=1144">Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons</a>. There was an error in the author's name. The author name was corrected to include a middle initial as follows:

David M. Kwinter, Michael A. Silverman

instead of:

David Kwinter, Michael Silverman.

A correction to author names has been made for article Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons.

Erratum: Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons.

<!DOCTYPE html PUBLIC "-//W3C//DTD XHTML 1.0 Transitional//EN" "http://www.w3.org/TR/xhtml1/DTD/xhtml1-transitional.dtd">
<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>MEM-Eagle with Earle salts and L-glutamine</td>
<td>Reagent</td>
<td>Mediatech</td>
<td>10-010-CV</td>
<td>&nbsp;</td>
<tr>
<td>Lipofectamine 2000</td>
<td>Reagent</td>
<td>Invitrogen</td>
<td>11668-027</td>
<td>Transfection reagent</td>
</tr>
<tr>
<td>10X Hanks with Ca2+ and Mg2+</td>
<td>Reagent</td>
<td>Gibco/Invitrogen</td>
<td>14185-052</td>
<td>Live-imaging medium</td>
</tr>
<tr>
<td>1M HEPES</td>
<td>Reagent</td>
<td>Gibco/Invitrogen</td>
<td>15630-130</td>
<td>Live-imaging medium</td>
</tr>		</tbody>
		</table>
		</div>

live imaging of dense-core vesicles in primary cultured hippocampal neurons

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.

Watch this Scientific Journal Video about Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons at JoVE.com

Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

Live cell imaging is of particular utility when studying the dynamics of organelle trafficking. Here we describe a protocol for live imaging of dense-core vesicles in cultured neurons using wide-field fluorescence microscopy. This protocol is flexible and can be adapted to image other organelles such as mitochondria, endosomes, and peroxisomes.

Observing and characterizing dynamic cellular processes can yield important information about cellular activity that cannot be gained from static images.  ...

live-imaging-dense-core-vesicles-primary-cultured-hippocampal

Research

JoVE Journal

Biology

12.5K ビュー.  Simon Fraser University. オルガネラの人身売買のダイナミクスを研究するときに、ライブセルイメージングには特に有用です。ここでは、広視野蛍光顕微鏡を用いた培養神経細胞における高密度コア小胞のライブイメージングのためのプロトコルについて説明します。このプロトコルは柔軟性があり、画像その他のようなミトコンドリアなどのオルガネラ、エンドソーム、およ?...

ビデオ: Live Imaging of Dense-core Vesicles in Primary Cultured Hippocampal Neurons

Gene-gun Transfection of Hippocampal Neurons

Primary Culture of Hippocampal Neurons from P0 Newborn Rats

The physiological properties of hippocampal neurons are commonly investigated, especially because of the involvement of the hippocampus in learning and memory. Primary hippocampal cell culturing allows neuroscientists to examine the activity and properties of neurons at the individual cell and single synapse level. In this video, we will demonstrate how to isolate and grow primary hippocampal cells from newborn rats. The hippocampus may be isolated from each newborn animal in as short as 2 to 3 minutes, and the cultures can be maintained for up to two weeks. We will also briefly demonstrate how to use these hippocampal neurons for ratiometric calcium imaging. While this protocol describes the process for the hippocampus, with little to no modification, it can be applied to other regions of the brain.

The dissection and growth of cells from an individual brain area facilitates investigation of cellular and physiological parameters. We describe a method for primary cell culturing that produces neuron-enriched cultures in a serum-free environment.

The physiological properties of hippocampal neurons are commonly investigated, especially because of the involvement of the hippocampus in learning and ...

Live Imaging of Cell Motility and Actin Cytoskeleton of Individual Neurons and Neural Crest Cells in Zebrafish Embryos

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily accessible at all stages of development. Moreover, their optical clarity allows high resolution imaging of cell and molecular dynamics in the natural environment of the intact embryo. We are using a live imaging approach to analyze cell behaviors during neural crest cell migration and the outgrowth and guidance of neuronal axons.
Live imaging is particularly useful for understanding mechanisms that regulate cell motility processes. To visualize details of cell motility, such as protrusive activity and molecular dynamics, it is advantageous to label individual cells. In zebrafish, plasmid DNA injection yields a transient mosaic expression pattern and offers distinct benefits over other cell labeling methods. For example, transgenic lines often label entire cell populations and thus may obscure visualization of the fine protrusions (or changes in molecular distribution) in a single cell. In addition, injection of DNA at the one-cell stage is less invasive and more precise than dye injections at later stages.
Here we describe a method for labeling individual developing neurons or neural crest cells and imaging their behavior in vivo. We inject plasmid DNA into 1-cell stage embryos, which results in mosaic transgene expression. The vectors contain cell-specific promoters that drive expression of a gene of interest in a subset of sensory neurons or neural crest cells. We provide examples of cells labeled with membrane targeted GFP or with a biosensor probe that allows visualization of F-actin in living cells1.
Erica Andersen, Namrata Asuri, and Matthew Clay contributed equally to this work.

This protocol describes imaging of individual neurons or neural crest cells in living zebrafish embryos. This method is used to examine cellular behaviors and actin localization using fluorescence confocal time-lapse microscopy.

The zebrafish is an ideal model for imaging cell behaviors during development in vivo. Zebrafish embryos are externally fertilized and thus easily ...

Adenovirus-mediated Genetic Removal of Signaling Molecules in Cultured Primary Mouse Embryonic Fibroblasts

The ability to genetically remove specific components of various cell signalling cascades has been an integral tool in modern signal transduction analysis. One particular method to achieve this conditional deletion is via the use of the Cre-loxP system. This method involves flanking the gene of interest with loxP sites, which are specific recognition sequences for the Cre recombinase protein. Exposure of the so-called floxed (flanked by loxP site) DNA to this enzyme results in a Cre-mediated recombination event at the loxP sites, and subsequent excision of the intervening gene3. Several different methods exist to administer Cre recombinase to the site of interest. In this video, we demonstrate the use of an adenovirus containing the Cre recombinase gene to infect primary mouse embryonic fibroblasts (MEFs) obtained from embryos containing a floxed Rac1 allele1. Our rationale for selecting Rac1 MEFs for our experiments is that clear morphological changes can be seen upon deletion of Rac1, due to alterations in the actin cytoskeleton2,5. 72 hours following viral transduction and Cre expression, cells were stained using the actin dye phalloidin and imaged using confocal laser scanning microscopy. It was observed that MEFs which had been exposed to the adeno-Cre virus appeared contracted and elongated in morphology compared to uninfected cells, consistent with previous reports2,5. The adenovirus method of Cre recombinase delivery is advantageous as the adeno-Cre virus is easily available, and gene deletion via Cre in nearly 100% of the cells can be achieved with optimized adenoviral infection.

In this video we use an adenovirus carrying the Cre recombinase gene to infect primary mouse embryonic fibroblasts carrying a floxed Rac1 allele.

The ability to genetically remove specific components of various cell signalling cascades has been an integral tool in modern signal transduction ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila 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.

Organelle Transport in Cultured Drosophila Cells: S2 Cell Line and Primary Neurons.

Drosophila S2 cells and cultured neurons are great systems for imaging of motor-driven organelle transport in vivo. Here we describe detailed protocols for culturing both cell types, their imaging and analysis of transport.

Drosophila S2 cells plated on a coverslip in the presence of any actin-depolymerizing drug form long unbranched processes filled with uniformly polarized ...

Visualization of G3BP Stress Granules Dynamics in Live Primary Cells

SGs can be visualized in cells by immunostaining of specific protein components or polyA+ mRNAs. SGs are highly dynamic and the study of their assembly and fate is important to understand the cellular response to stress. The deficiency in key factors of SGs like G3BP (RasGAP SH3 domain Binding Protein) leads to developmental defects in mice&#160;and alterations of the Central Nervous System. To study the dynamics of SGs in cells from an organism, one can culture primary cells and follow the localization of a transfected tagged component of SGs. We describe time-lapse experiment to observe G3BP1-containing SGs in Mouse Embryonic Fibroblasts (MEFs). This technique can also be used to study G3BP-containing SGs in live neurons, which is crucial as it was recently shown that these SGs are formed at the onset of neurodegenerative diseases like Alzheimer&#39;s disease. This approach can be adapted to any other cellular body and granule protein component, and performed with transgenic animals, allowing the live study of granules dynamics for example in the absence of a specific factor of these granules.

Stress granules (SGs) are cytoplasmic RNA granules containing stalled ribonucleoprotein particles (RNPs), and important in cellular response to various stresses. Dynamics of SGs can be followed in live cells by visualizing the localization of a tagged component of SGs in transfected primary cells after stress.

SGs can be visualized in cells by immunostaining of specific protein components or polyA+ mRNAs. SGs are highly dynamic and the study of their assembly ...

Live Cell Imaging of Primary Rat Neonatal Cardiomyocytes Following Adenoviral and Lentiviral Transduction Using Confocal Spinning Disk Microscopy

Primary rat neonatal cardiomyocytes are useful in basic in vitro cardiovascular research because they can be easily isolated in large numbers in a single procedure. Due to advances in microscope technology it is relatively easy to capture live cell images for the purpose of investigating cellular events in real time with minimal concern regarding phototoxicity to the cells. This protocol describes how to take live cell timelapse images of primary rat neonatal cardiomyocytes using a confocal spinning disk microscope following lentiviral and adenoviral transduction to modulate properties of the cell. The application of two different types of viruses makes it easier to achieve an appropriate transduction rate and expression levels for two different genes. Well focused live cell images can be obtained using the microscope&#8217;s autofocus system, which maintains stable focus for long time periods. Applying this method, the functions of exogenously engineered proteins expressed in cultured primary cells can be analyzed. Additionally, this system can be used to examine the functions of genes through the use of siRNAs as well as of chemical modulators.

This protocol describes a method of live cell imaging using primary rat neonatal cardiomyocytes following lentiviral and adenoviral transduction using confocal spinning disk microscopy. This enables detailed observations of cellular processes in living cardiomyocytes.

Primary rat neonatal cardiomyocytes are useful in basic in vitro cardiovascular research because they can be easily isolated in large numbers in a single ...