This laboratory has been supported by the Canadian Institutes of Health Research, the Natural Sciences and Engineering Research Council of Canada, and the Multiple Sclerosis Society of Canada (MSSC). GSTS was the recipient of a Doctoral Studentship from the MSSC. We are grateful to Dr. Joan Boggs (Hospital for Sick Children, Toronto) for many helpful discussions and comments on this manuscript. We are grateful to Biotium for their generous gift of additional NucView 488 caspase-3.

1. Thawing and Culturing Cells
<ol>
 <li>Obtain stock of frozen immortalized N19-oligodendroglial cells from long term liquid nitrogen stores.</li>
 <li>Immerse vial containing cells at 37 &deg;C water bath until the cell suspension is completely thawed.</li>
 <li>Add 7 mL of DMEM (Dulbecco's Modified Eagle Medium) with high-glucose supplemented with 10% FBS (Fetal Bovine Serum) and 1% penicillin/streptomycin to a 10 cm culture plate.</li>
 <li>Add the cell-suspension drop-wise to the plate, and gently agitate and rock the Petri plate to disperse the cells evenly.</li>
 <li>Culture cells at 34 &deg;C/5% CO2 incubator.</li>
 <li>After 4 hours, aspirate media from plates to remove any remaining DMSO (dimethyl sulphoxide) that had been used as a cryoprotectant, and replace with fresh media (7 mL DMEM high-glucose media supplemented with 10% FBS and 1% penicillin/streptomycin).</li>
</ol>
2. Passaging Cells
<ol>
 <li>At 70-80% confluence (4-7 days growth), aspirate media from cells.</li>
 <li>Add 1 mL of 0.25% trypsin to plate. Pipette trypsin to detach cells (about 5 minutes).</li>
 <li>Make the appropriate dilutions for experimental conditions and passage an additional 10 cm plate for future experiments at a cell density no less that 0.1 x 106 cells / mL.</li>
</ol>
Note: You should passage your cells at least twice after thawing before using them for experiments.
3. Counting and Plating Cells
<ol>
 <li>To plate cells for live-cell imaging, trypsinize as previously described.</li>
 <li>Remove approximately 30 &mu;L of cells and count using a haemocytometer.</li>
 <li>Place an uncoated glass coverslip in a 6 well plate. Add cells to the well at a density of 0.1 x 106 cells/mL in 2 mL phenol-free DMEM high-glucose media, supplemented with 10% FBS and 1% penicillin/streptomycin, at 34 &deg;C/5% CO2.</li>
 <li>Allow cells to grow overnight (16-20 h) at 34 &deg;C/5% CO2 before transfection.</li>
</ol>
4. Transfection
<ol>
 <li>Combine 100 &mu;L serum-free media, 0.5-4 &mu;g purified plasmid DNA, and 4 &mu;L FuGENE HD (Roche). Vortex gently to mix.</li>
 <li>Vortex again briefly and allow the DNA to complex for 5 min at room temperature, and then add FuGENE HD DNA mixture directly to the cultured cells. Tilt the plate gently to mix.</li>
 <li>Culture the cells for an additional 48 h at 34 &deg;C/5% CO2 prior to treatment or experimentation.</li>
</ol>
5. Preparing Cells for Live-Cell Imaging (LCI)
<ol>
 <li>Turn on the LCI Chamlide live-cell instrument control box at least 1 h before you want to start your experiment. The control box also regulates the temperature and humidity within the LCI Chamlide, and regulates the flow of the premixed 5% CO2.</li>
</ol>
Note: It is advisable to turn on the control box up to 3 h before beginning experiments to ensure that the entire stage reaches 34 &deg;C. This step will reduce focal drift, which is caused by fluxulation and thermal expansion of the metal stage as it warms, during imaging.
<ol start="2">
 <li>Working in a flow-hood, spray the Chamlide magnetic-type culture chamber, and tweezers, with 70% ethanol and allow them to dry for 5 minutes.</li>
 <li>Remove cells from incubator and confirm that they are healthy. They should appear adherent and well-spread on the glass coverslip with numerous membrane processes. Cells that are stressed from transfection are not suitable for experimentation, and will have a reduced number of process extensions, and often have an irregularly-shaped nucleus.</li>
 <li>Tilt the 6-well plate and remove the coverslip with tweezers. Quickly place the coverslip cell side up into the bottom plate of the culture chamber. Do not allow the coverslip to dry.</li>
 <li>Attach the magnetic main body of the culture chamber and add 500 &mu;L of media from the original 6-well plate onto the top of the coverslip. Re-using this media will decrease the amount of stress placed on the cells caused by environmental changes, and can also be useful for assessing extracellular secreted factors.</li>
 <li>Place the glass cover on the culture chamber.</li>
 <li>Use a KimWipe sprayed with 70% ethanol to remove any residual material from the bottom of the coverslip, which would interfere with microscopy.</li>
 <li>Place the culture chamber in the 34 &deg;C/5% CO2 incubator for 30 min. This step will ensure that the chamber itself warms to 34 &deg;C to reduce shifting of the coverslip as the metal warms up.</li>
</ol>
6. Microscope Settings
Images were acquired here using a Leica DMIRE2 inverted microscope with a custom relay lens and emission filter wheel housing for cartridge loading of multiple wheels (Quorum Technologies Inc., Guelph, ON).
<ol>
 <li>Brightfield - Lamp set to 2.5 V and exposure time to 240 ms.</li>
 <li>Red Fluorescent Protein (RFP) - Gain set at 140 for 200 ms.</li>
 <li>Green Fluorescent Protein (GFP) - Gain set at 140 for 500 ms.</li>
 <li>Our microscope has a dimmable lamp rather than a spinning disk to control the amount of light available to cells. We run our experiments with the light at 90% of maximum lamp intensity.</li>
</ol>
7. Treatment of Cells with Apoptosis Inducers and NucView 488 Caspase-3 substrate
<ol>
 <li>It is important to prepare large stocks of apoptosis inducers ahead of time and freeze them in aliquots, so that the concentrations will be consistent between experiments.</li>
 <li>The NucView 488 substrate is light sensitive. Prepare aliquots of 15 &mu;L to reduce freeze/thawing, and store tubes at -20 &deg;C covered in aluminum foil. Work with the ambient room lighting as low as possible to reduce exposure of the NucView 488 substrate to light, prior to its use.</li>
 <li>Retrieve the culture chamber quickly from the incubator so that the cells are not exposed to a reduction in temperature. Place the culture chamber onto the environmental chamber of the microscope, and use clamps to keep the culture chamber from moving. At this point you should also dim the room lights.</li>
 <li>Turn on the 5% premixed CO2 tank with dual regulator.</li>
 <li>Using the 10x objective, focus to find cells on the computer monitor using bright-field microscopy. Note: you would want to use the largest numerical aperture on the objective with the desired magnification to reduce the exposure time.</li>
 <li>Deliver inducers of apoptosis to the cells (in our case 80 mM potassium, or 100 mM glutamate), one of the following methods can be used. We will use delivery of 80 mM potassium as an example, using KCl dissolved as a 10x concentrate in our typical culture media.
 <ol>
 <li>Media exchange by slow and local perfusion: 
 - Use a peristaltic pump to exchange media in the culture chamber with new media containing 80 mM potassium. 
 - One end of the tubing will deliver 10 mL of a stock of 80 mM potassium dissolved in phenol-free DMEM, and the other end of the tubing will remove media from the chamber. 
 - You want at least a 10x exchange of media to ensure that the media left in the chamber has the correct concentration of potassium. 
 - After media is exchanged, add 3 &mu;L of NucView 488 substrate to the media in the chamber. Pipette to mix.</li>
 <li>Direct addition of 80 mM K+ and NucView 488 substrate to media in culture chamber: 
 - Prepare a 10x stock solution of 80 mM potassium dissolved in phenol-free DMEM. 
 - Add 3 &mu;L of NucView 488 substrate to 50 &mu;L of 10x 80 mM potassium. Pipette to mix. Add to 500 &mu;L phenol-free DMEM in culture chamber. 
 Note: this method is preferred if you are interested in maintaining or assessing growth or other secreted factors that may be present in the original media. We have found that the NucView 488 substrate is stable in cell culture for experiments lasting as long as 36 h.</li>
 </ol>
 </li>
</ol>
8. Live-cell Imaging
<ol>
 <li>Click on the red channel to display transfected cells.</li>
 <li>Select and save around a dozen frames where there are several transfected cells, and save these "X-Y stage points". Depending on the amount of computer memory, you may be limited to the number of stage points that you will be able to acquire.</li>
 <li>Have the microscope capture images (in bright-field, red channel, and green channel) at these saved stage points every 6 minutes.</li>
 <li>Any green staining that is visible during the early stages of the experiments likely indicates cells that are already undergoing apoptosis, due usually to transfection and/or environmental stress. Apoptosis due to the experimental treatments will be detected at a later timepoint in the experiment.</li>
</ol>
9. Statistical Analysis
<ol>
 <li> For each experiment, we program the microscope to acquire multiple X-Y points to gather a large data set efficiently. Each experiment is performed in duplicate or triplicate, and data are compiled from separate experiments performed on different days. From each data set, we analyze up to 15 fields of view and compare the ratio of caspase-negative cells to caspase-positive cells (total number of cells in field of view/total number of caspase-positive cells). </li>
 <li>The recorded measurements from each data set are grouped into a larger sample set, and are then compared to one another using an ANOVA table (p = 0.05). We show the standard errors of the mean (SEM) of each experiment, and then compare the difference in means by performing a Tukey means comparison test (p = 0.05) to determine which treatments are significantly different from each other.</li>
</ol>
10. Representative Results
We have described an experiment to illustrate how the NucView 488 substrate can indicate an increased rate of apoptosis of N19-OLG cell cultures following a treatment with a high extracellular potassium concentration. The N19-cells were transfected with RFP, and were either treated with 3 &mu;L NucView 488 substrate (control), or 3 &mu;L NucView 488 substrate and 80 mM [K+] (treatment). Cells were monitored and images were acquired over a 12 h time course, which is adequate for studies involving neurological insults (Figure 1A). In control conditions, we did not observe significant amounts of apoptosis compared to the 80 mM [K+] treated cultures (hashed box), which showed approximately 45% cell death after 12 h (Figure 1B). The background green signal observed in the control conditions indicates the cells that are undergoing apoptosis without the addition of extracellular potassium. Virtually none of the cells in the control experiment exhibit apoptosis by the 12 h time point, although in other situations the experiments may be required to run longer. Images were acquired using a 10x objective. Bar = 100 &mu;m.
<img alt="Figure 1" src="/files/ftp_upload/3422/3422fig1.jpg" /> 
Figure 1. (A) A 12 h time course experiment of N19 OLG cultures 48 h post-transfection expressing RFP-MBP (red channel) along with 3 &mu;L of NucView 488 substrate (green channel). Cultures were either treated with a final concentration of 80 mM [K+] (left panels), or no treatment as a control (right panels). Images were acquired at 6 min intervals, and significant activation of cleaved caspase-3 (green signal) can be observed in cultures treated with 80 mM [K+] within the cell nuclei (hashed box) compared to the control experiment. (B) Percentage of cleaved caspase-3 cells (calculated by dividing the total number of cells in the field of view by the total number of caspase-positive cells). In comparison to control conditions, we observed around 45% cell death following K+-treatment by 12 h.
sVideo 1. A 12 h time course experiment of N19-OLG cultures 48 h post-transfection expressing RFP-MBP (red channel) with 3 &mu;L of NucView 488 substrate (green channel), along with bright-field images and a three-way merged image, following treatment with 80 mM [K+]. Please <a href="https://www.jove.com/files/ftp_upload/3422/sVideo1_N19_OLGs_80mM_KCl.mov">click here</a> to see/download this video file.
sVideo 2. A 12 h time course experiment of N19-OLG cultures 48 h post-transfection expressing RFP-MBP (red channel) with 3 &mu;L of NucView 488 substrate (green channel), along with bright-field images and a three-way merged image. No treatment was applied to the cultures and N19-OLGs can be seen migrating through the microscope field in contrast to cell cultures treated with 80 mM [K+] (compare with sVideo 1). Please <a href="https://www.jove.com/files/ftp_upload/3422/sVideo2_N19_OLGs_control_no_treatment.mov">click here</a> to see/download this video file.

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The authors have no conflicts of interest to disclose.

Although this is not the only fluorogenic product available for apoptosis detection, there are several significant advantages to using the NucView 488 substrate. One of the main benefits is the ability to follow apoptosis in live cells in real time, whereas most alternative products either require cell lysis or have poor cell permeability. Other benefits include high sensitivity for caspase-3 recognition, high cell permeability, low cytotoxicity, and no interference with the progression of apoptosis. The substrate also has low background fluorescence until it is cleaved and enters the nucleus, which eliminates background fluorescence. The caspase-3 recognition sequence contains 3 negative charges and the DNA-binding dye has one positive charge 2. The DEVD-NucView 488 molecule thus has a net negative charge, which prevents the activation and binding of the dye to DNA in cells where caspase is not active.
Maintaining consistency between experiments is required to be able to draw meaningful comparisons between replicates. One of the main parameters to keep consistent is cell density, as it affects the rate of transfection. Obtaining high transfection rates in immortalized N19-OLG cell cultures is more difficult than with other common cell lines such as HeLa or HEK293, and is highly dependent on density, in our experience 12, 13. Our best transfection efficiencies for these cells have been obtained with Fugene HD (Roche) and are normally about 15%, but can reach over 30%, depending on the construct. Careful cell counting will aid in obtaining consistent transfection rates. It is also important to expose cells from different treatments to the same degree of environmental stress, particularly when measuring rates of apoptosis. Specifically, the length of time over which the cells are exposed to transfection reagents, or the amount of light exposure, are important environmental variables to consider, and should remain constant across experiments. For our applications, we have found that collecting a large number of fields-of-view provides a sufficiently large sample size to make statistically-significant comparisons [ibid].

Table of specific reagents and equipment
<table border="1">
 <tr>
 <th>Name of Reagent</th>
 <th>Company</th>
 <th>Catalogue Number</th>
 </tr>
 <tr>
 <td>NucView 488 Caspase-3 Assay Kit for Live Cells</td>
 <td>Biotium</td>
 <td>30029</td>
 </tr>
 <tr>
 <td>FuGENE HD transfection reagent</td>
 <td>Roche</td>
 <td>04709705001</td>
 </tr>
 <tr>
 <td>Dulbecco&rsquo;s Modified Eagle Medium</td>
 <td>Gibco</td>
 <td>31053-028</td>
 </tr>
 <tr>
 <td>0.25% Trypsin</td>
 <td>Gibco</td>
 <td>15050-065</td>
 </tr>
 <tr>
 <td>Fetal Bovine Serum</td>
 <td>Gibco</td>
 <td>12483-020</td>
 </tr>
 <tr>
 <td>Penicillin/streptomycin</td>
 <td>Gibco</td>
 <td>15140122</td>
 </tr>
 <tr>
 <td>#1.5-25 mm glass coverslip </td>
 <td>Warner Instruments</td>
 <td>64-0715</td>
 </tr>
 <tr>
 <td>Chamlide CMB magnetic 
 culture chamber for 25 mm
 coverslip</td>
 <td>Quorum Technologies</td>
 <td>CM-B-40</td>
 </tr>
 <tr>
 <td>Cellstart tissue culture 10 cm dishes </td>
 <td>VWR</td>
 <td>82050-576</td>
 </tr>
 <tr>
 <td>BD Falcon 6-well tissue culture dishes</td>
 <td>VWR</td>
 <td>CA62406-161</td>
 </tr>
</table>

monitoring cleaved caspase-3 activity and apoptosis of immortalized oligodendroglial cells using live-cell imaging and cleaveable fluorogenic-dye substrates following potassium-induced membrane depolarization

The central nervous system can experience a number of stresses and neurological insults, which can have numerous adverse effects that ultimately lead to a reduction in neuronal population and function. Damaged axons can release excitatory molecules including potassium or glutamate into the extracellular matrix, which in turn, can produce further insult and injury to the supporting glial cells including astrocytes and oligodendrocytes 8, 16. If the insult persists, cells will undergo programmed cell death (apoptosis), which is regulated and activated by a number of well-established signal transduction cascades 14. Apoptosis and tissue necrosis can occur after traumatic brain injury, cerebral ischemia, and seizures. A classical example of apoptotic regulation is the family of cysteine-dependent aspartate-directed proteases, or caspases. Activated proteases including caspases have also been implicated in cell death in response to chronic neurodegenerative diseases including Alzheimer's, Huntington's, and Multiple Sclerosis 4, 14, 3, 11, 7.
In this protocol we describe the use of the NucView 488 caspase-3 substrate to measure the rate of caspase-3 mediated apoptosis in immortalized N19-oligodendrocyte (OLG) cell cultures 15, 5, following exposure to different extracellular stresses such as high concentrations of potassium or glutamate. The conditionally-immortalized N19-OLG cell line (representing the O2A progenitor) was obtained from Dr. Anthony Campagnoni (UCLA Semel Institute for Neuroscience) 15, 5, and has been previously used to study molecular mechanisms of myelin gene expression and signal transduction leading to OLG differentiation (e.g.6, 10). We have found this cell line to be robust with respect to transfection with exogenous myelin basic protein (MBP) constructs fused to either RFP or GFP (red or green fluorescent protein) 13, 12. Here, the N19-OLG cell cultures were treated with either 80 mM potassium chloride or 100 mM sodium glutamate to mimic axonal leakage into the extracellular matrix to induce apoptosis 9. We used a bi-functional caspase-3 substrate containing a DEVD (Asp-Glu-Val-Asp) caspase-3 recognition subunit and a DNA-binding dye 2. The substrate quickly enters the cytoplasm where it is cleaved by intracellular caspase-3. The dye, NucView 488 is released and enters the cell nucleus where it binds DNA and fluoresces green at 488 nm, signaling apoptosis. Use of the NucView 488 caspase-3 substrate allows for live-cell imaging in real-time 1, 10. In this video, we also describe the culturing and transfection of immortalized N19-OLG cells, as well as live-cell imaging techniques.

Watch this Scientific Journal Video about Monitoring Cleaved Caspase-3 Activity and Apoptosis of Immortalized Oligodendroglial Cells using Live-cell Imaging and Cleaveable Fluorogenic-dye Substrates F

Monitoring Cleaved Caspase-3 Activity and Apoptosis of Immortalized Oligodendroglial Cells using Live-cell Imaging and Cleaveable Fluorogenic-dye Substrates Following Potassium-induced Membrane Depola

Live-cell imaging of caspase-3 mediated apoptosis in immortalized N19-oligodendrocyte cell cultures using the NucView 488 caspase-3 substrate. This technique is applicable for programmed cell death assays in real-time in a variety of cell types and tissues.

Monitoring Cleaved Caspase-3 Activity and Apoptosis of Immortalized Oligodendroglial Cells using Live-cell Imaging and Cleaveable Fluorogenic-dye Substrates Following Potassium-induced Membrane Depolarization

The central nervous system can experience a number of stresses and neurological insults, which can have numerous adverse effects that ultimately lead to a ...

monitoring-cleaved-caspase-3-activity-apoptosis-immortalized

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Neuroscience

15.9K ビュー.  University of Guelph. 使用して不死化N19 -オリゴデンドロサイトの培養細胞におけるカスパーゼ-3を介するアポトーシスの生細胞イメージング NucView 488カスパーゼ3基質。この手法は、細胞の種類や組織の様々なリアルタイムでのプログラム細胞死のアッセイに適用可能です。

ビデオ: Monitoring Cleaved Caspase-3 Activity and Apoptosis of Immortalized Oligodendroglial Cells using Live-cell Imaging and Cleaveable Fluorogenic-dye Substrates Following Potassium-induced Membrane Depola

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding fundamental biological mechanisms. Striking progress has been particularly evident in Drosophila, whose extensive toolkit of mutants and transgenic lines provides a convenient model to study evolutionarily-conserved developmental and cell biological mechanisms. We are interested in understanding the mechanisms that control cell fate specification in the adult peripheral nervous system (PNS) in Drosophila. Bristles that cover the head, thorax, abdomen, legs and wings of the adult fly are individual mechanosensory organs, and have been studied as a model system for understanding mechanisms of Notch-dependent cell fate decisions. Sensory organ precursor (SOP) cells of the microchaetes (or small bristles), are distributed throughout the epithelium of the pupal thorax, and are specified during the first 12 hours after the onset of pupariation. After specification, the SOP cells begin to divide, segregating the cell fate determinant Numb to one daughter cell during mitosis. Numb functions as a cell-autonomous inhibitor of the Notch signaling pathway.
Here, we show a method to follow protein dynamics in SOP cell and its progeny within the intact pupal thorax using a combination of tissue-specific Gal4 drivers and GFP-tagged fusion proteins 1,2.This technique has the advantage over fixed tissue or cultured explants because it allows us to follow the entire development of an organ from specification of the neural precursor to growth and terminal differentiation of the organ. We can therefore directly correlate changes in cell behavior to changes in terminal differentiation. Moreover, we can combine the live imaging technique with mosaic analysis with a repressible cell marker (MARCM) system to assess the dynamics of tagged proteins in mitotic SOPs under mutant or wildtype conditions. Using this technique, we and others have revealed novel insights into regulation of asymmetric cell division and the control of Notch signaling activation in SOP cells (examples include references 1-6,7 ,8).

Live-cell Imaging of Sensory Organ Precursor Cells in Intact Drosophila Pupae

In this video, we describe a method for live cell imaging of asymmetrically dividing sensory organ progenitor cells and epidermal cells in intact Drosophila pupae

Since the discovery of Green Fluorescent Protein (GFP), there has been a revolutionary change in the use of live-cell imaging as a tool for understanding ...

Quantitative Analysis of Synaptic Vesicle Pool Replenishment in Cultured Cerebellar Granule Neurons using FM Dyes

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis. Retrieved SVs are then refilled with neurotransmitter and rejoin the recycling pool, defined as SVs that are available for exocytosis1,2. The recycling pool can generally be subdivided into two distinct pools - the readily releasable pool (RRP) and the reserve pool (RP). As their names imply, the RRP consists of SVs that are immediately available for fusion while RP SVs are released only during intense stimulation1,2. It is important to have a reliable assay that reports the differential replenishment of these SV pools in order to understand 1) how SVs traffic after different modes of endocytosis (such as clathrin-dependent endocytosis and activity-dependent bulk endocytosis) and 2) the mechanisms controlling the mobilisation of both the RRP and RP in response to different stimuli.
FM dyes are routinely employed to quantitatively report SV turnover in central nerve terminals3-8. They have a hydrophobic hydrocarbon tail that allows reversible partitioning in the lipid bilayer, and a hydrophilic head group that blocks passage across membranes. The dyes have little fluorescence in aqueous solution, but their quantum yield increases dramatically when partitioned in membrane9. Thus FM dyes are ideal fluorescent probes for tracking actively recycling SVs. The standard protocol for use of FM dye is as follows. First they are applied to neurons and are taken up during endocytosis (Figure 1). After non-internalised dye is washed away from the plasma membrane, recycled SVs redistribute within the recycling pool. These SVs are then depleted using unloading stimuli (Figure 1). Since FM dye labelling of SVs is quantal10, the resulting fluorescence drop is proportional to the amount of vesicles released. Thus, the recycling and fusion of SVs generated from the previous round of endocytosis can be reliably quantified.
Here, we present a protocol that has been modified to obtain two additional elements of information. Firstly, sequential unloading stimuli are used to differentially unload the RRP and the RP, to allow quantification of the replenishment of specific SV pools. Secondly, each nerve terminal undergoes the protocol twice. Thus, the response of the same nerve terminal at S1 can be compared against the presence of a test substance at phase S2 (Figure 2), providing an internal control. This is important, since the extent of SV recycling across different nerve terminals is highly variable11.
Any adherent primary neuronal cultures may be used for this protocol, however the plating density, solutions and stimulation conditions are optimised for cerebellar granule neurons (CGNs)12,13.

A live fluorescence imaging technique to quantify the replenishment and mobilisation of specific synaptic vesicle (SV) pools in central nerve terminals is described. Two rounds of SV recycling are monitored in the same nerve terminals providing an internal control.

After neurotransmitter release in central nerve terminals, SVs are rapidly retrieved by endocytosis.  Retrieved SVs are then refilled with ...

Caspase-3 Activity in the Rat Amygdala Measured by Spectrofluorometry After Myocardial Infarction

Myocardial infarction (MI) has dramatic mid- and long-term consequences at the physiological and behavioral levels, but the mechanisms involved are still unclear. Our laboratory has developed a rat model of post-MI syndrome that displays impaired cardiac functions, neuronal loss in the limbic system, cognitive deficits and behavioral signs of depression. At the neuronal level, caspase-3 activation mediates post-MI apoptosis in different limbic regions, such as the amygdala &#8211; peaking at 3 days post-MI. Cognitive and behavioral impairments appear 2-3 weeks post-MI and these correlate statistically with measures of caspase-3 activity. The protocol described here is used to induce MI, collect amygdala tissue and measure caspase-3 activity using spectrofluorometry. To induce MI, the descending coronary artery is occluded for 40 min. The protocol for evaluation of caspase-3 activation starts 3 days after MI: the rats are sacrificed and the amygdala isolated rapidly from the brain. Samples are quickly frozen in liquid nitrogen and kept at -80 &#176;C until actual analysis. The technique performed to assess caspase-3 activation is based on cleavage of a substrate (DEVD-AMC) by caspase-3, which releases a fluorogenic compound that can be measured by spectrofluorometry. The methodology is quantitative and reproducible but the equipment required is expensive and the procedure for quantifying the samples is time-consuming. This technique can be applied to other tissues, such as the heart and kidneys. DEVD-AMC can be replaced by other substrates to measure the activity of other caspases.

Myocardial infarction (MI) is not only followed by impaired cardiac function but also by apoptosis in the amygdala, a brain region involved in the behavioral consequences of MI. This protocol describes how to induce MI, collect amygdala tissue and measure caspase-3 activity, a marker of apoptosis, therein.

Myocardial infarction (MI) has dramatic mid- and long-term consequences at the physiological and behavioral levels, but the mechanisms involved are still ...

Live-cell Measurement of Odorant Receptor Activation Using a Real-time cAMP Assay

The enormous sizes of the mammalian odorant receptor (OR) families present difficulties to find their cognate ligands among numerous volatile chemicals. To efficiently and accurately deorphanize ORs, we combine the use of a heterologous cell line to express mammalian ORs and a genetically modified biosensor plasmid to measure cAMP production downstream of OR activation in real time. This assay can be used to screen odorants against ORs and vice versa. Positive odorant-receptor interactions from the screens can be subsequently confirmed by testing against various odor concentrations, generating concentration-response curves. Here we used this method to perform a high-throughput screening of an odorous compound against a human OR library expressed in Hana3A cells and confirmed that the positively-responding receptor is the cognate receptor for the compound of interest. We found this high-throughput detection method to be efficient and reliable in assessing OR activation and our data provide an example of its potential use in OR functional studies.

Characterizing the function of odorant receptors serves an indispensable part in the deorphanization process. We describe a method to measure the activation of odorant receptors in real time using a cAMP assay.

The enormous sizes of the mammalian odorant receptor (OR) families present difficulties to find their cognate ligands among numerous volatile chemicals. ...

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in adult mammalian animals remains elusive. In this study, we describe an experimental procedure for measuring calcium dynamics through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy. This method enables recordings and quantifications of neural activity in bilateral mouse brain regions one at a time in the same experiment for a prolonged period in vivo. Key aspects of this method, which can be completed within an hour, include minimally invasive surgery procedures for creating dual optical windows, and the use of 2P imaging. Although we only demonstrate the technique in the S1 area, the method can be applied to other regions of the living brain facilitating the elucidation of structural and functional complexities of brain neural networks.

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in ...

Analyzing Neural Activity and Connectivity Using Intracranial EEG Data with SPM Software

Measuring neural activity and connectivity associated with cognitive functions at high spatial and temporal resolutions is an important goal in cognitive neuroscience. Intracranial electroencephalography (EEG) can directly record electrical neural activity and has the unique potential to accomplish this goal. Traditionally, averaging analysis has been applied to analyze intracranial EEG data; however, several new techniques are available for depicting neural activity and intra- and inter-regional connectivity. Here, we introduce two analytical protocols we recently applied to analyze intracranial EEG data using the Statistical Parametric Mapping (SPM) software: time-frequency SPM analysis for neural activity and dynamic causal modeling of induced responses for intra- and inter-regional connectivity. We report our analysis of intracranial EEG data during the observation of faces as representative results. The results revealed that the inferior occipital gyrus (IOG) showed gamma-band activity at very early stages (110 ms) in response to faces, and both the IOG and amygdala showed rapid intra- and inter-regional connectivity using various types of oscillations. These analytical protocols have the potential to identify the neural mechanisms underlying cognitive functions with high spatial and temporal profiles.

We present two analytical protocols that can be used to analyze intracranial electroencephalography data using the Statistical Parametric Mapping (SPM) software: time-frequency statistical parametric mapping analysis for neural activity, and dynamic causal modeling of induced responses for intra- and inter-regional connectivity.

Measuring neural activity and connectivity associated with cognitive functions at high spatial and temporal resolutions is an important goal in cognitive ...

ROS Live Cell Imaging During Neuronal Development

Reactive oxygen species (ROS) are well-established signaling molecules, which are important in normal development, homeostasis, and physiology. Among the different ROS, hydrogen peroxide (H2O2) is best characterized with respect to roles in cellular signaling. H2O2 has been implicated during the development in several species. For example, a transient increase in H2O2 has been detected in zebrafish embryos during the first days following fertilization. Furthermore, depleting an important cellular H2O2 source, NADPH oxidase (NOX), impairs nervous system development such as the differentiation, axonal growth, and guidance of retinal ganglion cells (RGCs) both in&#160;vivo and in vitro. Here, we describe a method for imaging intracellular H2O2 levels in cultured zebrafish neurons and whole larvae during development using the genetically encoded H2O2-specific biosensor, roGFP2-Orp1. This probe can be transiently or stably expressed in zebrafish larvae. Furthermore, the ratiometric readout diminishes the probability of detecting artifacts due to differential gene expression or volume effects. First, we demonstrate how to isolate and culture RGCs derived from zebrafish embryos that transiently express roGFP2-Orp1. Then, we use whole larvae to monitor H2O2 levels at the tissue level. The sensor has been validated by the addition of H2O2. Additionally, this methodology could be used to measure H2O2 levels in specific cell types and tissues by generating transgenic animals with tissue-specific biosensor expression. As zebrafish facilitate genetic and developmental manipulations, the approach demonstrated here could serve as a pipeline to test the role of H2O2 during neuronal and general embryonic development in vertebrates.

This protocol describes the use of a genetically encoded hydrogen peroxide (H2O2)-biosensor in cultured zebrafish neurons and larvae for assessing the physiological signaling roles of H2O2 during nervous system development. It can be applied to different cell types and modified with experimental treatments to study reactive oxygen species (ROS) in general development.

Reactive oxygen species (ROS) are well-established signaling molecules, which are important in normal development, homeostasis, and physiology. Among the ...

Microfluidics-Assisted Selective Depolarization of Axonal Mitochondria

Mitochondria are the primary suppliers of ATP (adenosine triphosphate) in neurons. Mitochondrial dysfunction is a common phenotype in many neurodegenerative diseases. Given some axons' elaborate architecture and extreme length, it is not surprising that mitochondria in axons can experience different environments compared to their cell body counterparts. Interestingly, dysfunction of axonal mitochondria often precedes effects on the cell body. To model axonal mitochondrial dysfunction in vitro, microfluidic devices allow treatment of axonal mitochondria without affecting the somal mitochondria. The fluidic pressure gradient in these chambers prevents diffusion of molecules against the gradient, thus allowing for analysis of mitochondrial properties in response to local pharmacological challenges within axons. The current protocol describes the seeding of dissociated hippocampal neurons in microfluidic devices, staining with a membrane-potential sensitive dye, treatment with a mitochondrial toxin, and the subsequent microscopic analysis. This versatile method to study axonal biology can be applied to many pharmacological perturbations and imaging readouts, and is suitable for several neuronal subtypes.

The present protocol describes the seeding and staining of neuronal mitochondria in microfluidic chambers. The fluidic pressure gradient in these chambers allows for the selective treatment of mitochondria in axons to analyze their properties in response to pharmacological challenges without affecting the cell body compartment.

Mitochondria are the primary suppliers of ATP (adenosine triphosphate) in neurons. Mitochondrial dysfunction is a common phenotype in many ...

Human Microglia-like Cells: Differentiation from Induced Pluripotent Stem Cells and In Vitro Live-cell Phagocytosis Assay using Human Synaptosomes

Microglia are the resident immune cells of myeloid origin that maintain homeostasis in the brain microenvironment and have become a key player in multiple neurological diseases. Studying human microglia in health and disease represents a challenge due to the extremely limited supply of human cells. Induced pluripotent stem cells (iPSCs) derived from human individuals can be used to circumvent this barrier. Here, it is demonstrated how to differentiate human iPSCs into microglia-like cells (iMGs) for in vitro experimentation. These iMGs exhibit the expected and physiological properties of microglia, including microglia-like morphology, expression of proper markers, and active phagocytosis. Additionally, documentation for isolating and labeling synaptosome substrates derived from human iPSC-derived lower motor neurons (i3LMNs) is provided. A live-cell, longitudinal imaging assay is used to monitor engulfment of human synaptosomes labeled with a pH-sensitive dye, allowing for investigations of iMG's phagocytic capacity. The protocols described herein are broadly applicable to different fields that are investigating human microglia biology and the contribution of microglia to disease.

Human Microglia-like Cells: Differentiation from Induced Pluripotent Stem Cells and In Vitro Live-cell Phagocytosis Assay using Human Synaptosomes

This protocol describes the differentiation process of human induced pluripotent stem cells (iPSCs) into microglia-like cells for in vitro experimentation. We also include a detailed procedure for generating human synaptosomes from iPSC-derived lower motor neurons that can be used as a substrate for in vitro phagocytosis assays using live-cell imaging systems.

Microglia are the resident immune cells of myeloid origin that maintain homeostasis in the brain microenvironment and have become a key player in multiple ...

Unveiling Mitochondrial Ultrastructure in Neurons Using STED Microscopy

Using Live Cell STED Imaging to Visualize Mitochondrial Inner Membrane Ultrastructure in Neuronal Cell Models

Mitochondria play many essential roles in the cell, including energy production, regulation of Ca2+ homeostasis, lipid biosynthesis, and production of reactive oxygen species (ROS). These mitochondria-mediated processes take on specialized roles in neurons, coordinating aerobic metabolism to meet the high energy demands of these cells, modulating Ca2+ signaling, providing lipids for axon growth and regeneration, and tuning ROS production for neuronal development and function. Mitochondrial dysfunction is therefore a central driver in neurodegenerative diseases. Mitochondrial structure and function are inextricably linked. The morphologically complex inner membrane with structural infolds called cristae harbors many molecular systems that perform the signature processes of the mitochondrion. The architectural features of the inner membrane are ultrastructural and therefore, too small to be visualized by traditional diffraction-limited resolved microscopy. Thus, most insights on mitochondrial ultrastructure have come from electron microscopy on fixed samples. However, emerging technologies in super-resolution fluorescence microscopy now provide resolution down to tens of nanometers, allowing visualization of ultrastructural features in live cells. Super-resolution imaging therefore offers an unprecedented ability to directly image fine details of mitochondrial structure, nanoscale protein distributions, and cristae dynamics, providing fundamental new insights that link mitochondria to human health and disease. This protocol presents the use of stimulated emission depletion (STED) super-resolution microscopy to visualize the mitochondrial ultrastructure of live human neuroblastoma cells and primary rat neurons. This procedure is organized into five sections: (1) growth and differentiation of the SH-SY5Y cell line, (2) isolation, plating, and growth of primary rat hippocampal neurons, (3) procedures for staining cells for live STED imaging, (4) procedures for live cell STED experiments using a STED microscope&#160;for reference, and (5) guidance for segmentation and image processing using examples to measure and quantify morphological features of the inner membrane.

Author Spotlight: Decoding Mitochondrial Aging 

This protocol presents a workflow for the propagation, differentiation, and staining of cultured SH-SY5Y cells and primary rat hippocampal neurons for mitochondrial ultrastructure visualization and analysis using stimulated emission depletion (STED) microscopy.

Mitochondria play many essential roles in the cell, including energy production, regulation of Ca2+ homeostasis, lipid biosynthesis, and production of ...