This work was supported by the National Institutes of Health (K22NS050137 to J.C.B.).

1. Cell culture
<ol>
 <li>Cortical neurons are isolated and grown using previously described techniques and plated on culture dishes with a glass bottom (MatTek Corporation, Ashland, MA ) coated with poly-D-lysine and laminin1.</li>
</ol>
2. Preparing the stock solutions for the fluorescent probes TMRM and H2DCF-DA
<ol>
 <li>Prepare a 10-mM stock solution of TMRM by dissolving 5.0 mg of TMRM in 1 ml of anhydrous dimethylsulfoxide. Vortex it for 1 min. Then, make aliquots and store them at -20&deg;C, protect from light, and use within one month.</li>
 <li>Next, prepare a 10-mM stock solution of H2DCF-DA by dissolving 4.87 mg of H2DCF-DA in 1 ml of anhydrous DMSO. Similarly, vortex it for 1 min. Then, make aliquots and store them at -20&deg;C, protect from light, and use within one week.</li>
</ol>
3. Loading rat cortical neurons with TMRM and H2DCF-DA
TMRM is a potentiometric, cell-permeable fluorescent indicator that accumulates in the highly negatively charged interior of mitochondria. It is important to use the low concentrations (10-50 nM range) of TMRM to avoid auto-quenching of mitochondrial TMRM. Then, the fluorescence signal of TMRM can be directly co-related to &Delta;&Psi;m across the inner mitochondrial membrane. A loss of &Delta;&Psi;m causes TMRM to leak from mitochondria resulting in a loss of fluorescence intensity. H2DCF-DA is cell-permeable probe converted into DCF-DA by intracellular esterases, and its oxidation results in fluorescent DCF. The final concentration of H2DCF-DA ranges between 2-10 &mu;M and it should be tested empirically in neurons derived from different brain regions since high loading concentrations might result in the saturation of DCF fluorescence even in the absence of H2O2. The presence of any endogenous or exogenous oxidant (e.g., nitric oxide, hydrogen peroxide) will increase DFC fluorescence intensity. Below, we provide a protocol for loading rat cortical neurons with TMRM and H2DCF-DA. 
<ol>
 <li>To load the rat cortical neurons with TMRM, first, wash the cultured neurons 3 times with Tyrode's buffer (Text Overlay: TB: 145 mM NaCl, 5 mM KCl, 10 mM glucose, 1.5 mM CaCl2, 1 mM MgCl2, and 10 mM HEPES; adjust pH to 7.4 with NaOH). Then, prepare 20 nM of TMRM by diluting the 10 mM TMRM stock 1/1000 times in TB and then add 2 &mu;l of diluted TMRM per 1 ml of TB. Incubate the neurons with TMRM for 45 min in the dark at room temperature. After 45 min, mount the culture dish on the stage of the microscope and start imaging.</li>
 <li>To load the rat cortical neurons with H2DCF-DA, wash the cultured neurons 3 times with TB. Next, prepare 2 &mu;M of H2DCF-DA by diluting the 10 mM H2DCF-DA stock 1/10 times in TB and then add 2 &mu;l of diluted H2DCF-DA per 1 ml of TB. Then, incubate the neurons with H2DCF-DA for 45 min in dark at room temperature. After 45 min, wash the neurons 4 times with TB to remove excess fluorescent indicator before obtaining images.</li>
</ol>
4. Live imaging of neurons incubated with TMRM to determine &Delta;&Psi;m
<ol>
 <li>To perform live imaging of neurons incubated with TMRM, confocal laser scanning microscopy (Text Overlay: LSM 510, Carl Zeiss Inc.), with the application of live time-series program, is used. Apply low-resolution and attenuated laser power (Text Overlay: low resolution: 256 x 256; laser power: 1%) to minimize the time needed to obtain images and to avoid photobleaching. </li>
 <li>. Next, adjust the focus of the mounted neurons loaded with TMRM using reflected light. Examine the TMRM fluorescence by illumination at 514 nm and detection at 570 nm. Set the detection gain of a camera just below the saturation level.</li>
 <li>Once all parameters which include resolution, laser power, detection gain of a camera, and time-lapse interval to obtain images are set; do not change these settings between experiments. Next, change the field. Start collecting images.</li>
 <li>To test changes in &Delta;&Psi;m, stimuli such as 1 &mu;M of FCCP or 2 &mu;g/ml of oligomycin, can be applied, which will significantly depolarize or hyperpolarize the mitochondrial membrane potential, respectively. These changes will be reflected by a decrease in TMRM fluorescence intensity compared with the baseline fluorescence intensity in the case of FCCP, or an increase in TMRM fluorescence intensity in the case of oligomycin. </li>
</ol>
5. Live imaging of neurons incubated with H2DCF-DA to determine ROS 
<ol>
 <li>To perform live imaging of neurons incubated with H2DCF-DA, first, mount the culture dish on the stage of a microscope. Adjust the focus of the cells using reflected light. Examine DCF fluorescence by excitation at 488 nm and emission at 515 nm.</li>
 <li>Next, adjust the laser power to 5-7%, detector gain, and resolution of 256 x 256. Do not change these settings between experiments. Then, set the frequency for obtaining live images using the time series program.</li>
 <li>Select a new field and start acquiring images. To detect changes in ROS levels, treat cells with 100-200 &mu;M of H2O2. This will be reflected by an increase in DCF fluorescence intensity compared with baseline level.</li>
</ol>
6. Data analysis
<ol>
 <li>Use region of interest (Text Overlay: ROI) tool from the LSM program to select the areas. Then, measure the TMRM or ROS fluorescence intensities. Select ROIs from mitochondrial regions or ROIs from the entire cell body in imaged cells to measure the fluorescence intensities of TMRM or ROS, respectively.</li>
 <li>Calculate the average fluorescence intensities from all ROIs of each cell for TMRM or from whole cell bodies for all imaged cells for ROS for each time point. Select regions next to the cells to calculate the background fluorescence intensity. Take several measurements and calculate the average background intensity.</li>
 <li>Subtract the average background fluorescence intensity from average fluorescence intensities of ROIs in each cell for each time point using Microsoft Excel. After subtracting background intensity, normalize the TMRM or DCF fluorescence intensity to the baseline fluorescence using this formula (Text Overlay: &Delta;F = F-Fo/Fo x 100, where F= fluorescence intensity at any time point, Fo= baseline fluorescence). Then, use the Sigma Plot program to generate the plot showing the changes in fluorescence intensity over time.</li>
</ol>
7. Representative Results
Figure 1A shows a fluorescence image of rat cortical neurons incubated with TMRM. Addition of FCCP, a mitochondrial uncoupler, leads to mitochondrial depolarization and a loss of TMRM fluorescence intensity (Fig. 1B). The baseline TMRM fluorescence level remains stable before addition of FCCP (the first 350 sec; Fig. 1C). Quantitative analysis of TMRM fluorescence changes over time shows a significant decrease in TMRM fluorescence after addition of FCCP (Fig. 1C).
Figure 1D shows the fluorescence image of rat cortical neurons loaded with DCF. Addition of H2O2 results in increased DCF fluorescence intensity in cell bodies (Fig. 1E). The baseline DCF fluorescence level is unchanged (the first 120 sec) before application of H2O2. Time-lapse measurements of DCF fluorescence show its steady levels, which increase after H2O2 treatment (Fig. 1F).

<img src="/files/ftp_upload/2704/2704fig1.jpg" alt="Figure 1"/> 
Figure 1. Assessment of mitochondrial membrane potential and ROS levels in live rat cortical neurons. (A) Representative fluorescence image of cortical neurons loaded with TMRM. After scanning the baseline TMRM fluorescence, neurons were treated with the protonophore FCCP (1 &mu;M). To the right is a pseudocolor intensity bar of TMRM fluorescence with bright yellow and black representing maximum and minimum intensity, respectively. The loss of TMRM fluorescence from the mitochondrial regions indicates the collapse of &Delta;&Psi;m upon FCCP treatment (panel B). The quantitative representation of change in TMRM fluorescence intensity at different time points before and after FCCP treatment is shown in panel C. (D) Fluorescence image of rat cortical neurons loaded with H2DCF-DA. After determining the baseline DCF fluorescence, cells were treated with 200 &mu;M H2O2, and the change in DCF fluorescence was assessed. An increase in the DCF fluorescence reflects the increase in ROS levels upon H2O2 treatment (E). Quantitative analysis of change in DCF fluorescence, before and after H2O2 treatment, is shown in panel F. Scale bar = 10 &mu;m
Video.7.1 &#x2013; labmedia 2704_Joshi.avi 
Live cell imaging of TMRM in cortical neurons before and after FCCP addition using 40X objective. The pseudocolor intensity shows a maximum (bright yellow, before FCCP addition) and decreased (red color, after FCCP addition) TMRM fluorescence intensity after FCCP addition. <a href="https://www.jove.com/files/ftp_upload/2704/supplemental_video1.avi">Click here to view video</a>
Video. 7.5 - labmedia 2704_Joshi.avi 
Live cell imaging of DCF in cortical neurons before and after H2O2 addition using 40X objective. The baseline DCF fluorescence has light green color in cell bodies and H2O2 addition increases the DCF fluorescence intensity to bright green color. <a href="https://www.jove.com/files/ftp_upload/2704/supplemental_video2.avi">Click here to view video</a>

<ol>
	<li>Hilgenberg, L. G., Smith, M. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+dissociated+mouse+cortical+neuron+cultures.">Preparation of dissociated mouse cortical neuron cultures.</a> J. Vis. Exp. , (2007).</li><li>Scaduto, R. C., Grotyohann, L. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+mitochondrial+membrane+potential+using+fluorescent+rhodamine+derivatives.">Measurement of mitochondrial membrane potential using fluorescent rhodamine derivatives.</a> Biophys. J. 76, 469-477 (1999).</li><li>Ward, M. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+amyloid+precursor+protein+intracellular+domain+(AICD)+disrupts+actin+dynamics+and+mitochondrial+bioenergetics.">The amyloid precursor protein intracellular domain (AICD) disrupts actin dynamics and mitochondrial bioenergetics.</a> J. Neurochem. 113, 275-284 (2010).</li><li>Gunasekar, P. G., Kanthasamy, A. G., Borowitz, J. L., Isom, G. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NMDA+receptor+activation+produces+concurrent+generation+of+nitric+oxide+and+reactive+oxygen+species:+implication+for+cell+death.">NMDA receptor activation produces concurrent generation of nitric oxide and reactive oxygen species: implication for cell death.</a> J. Neurochem. 65, 2016-2021 (1995).</li></ol>

No conflicts of interest declared.

We have presented a step-by-step procedure describing how to determine &Delta;&Psi;m and ROS in rat cortical neurons using the fluorescent indicators TMRM and H2DCF-DA, respectively. For other cell types, it is important to empirically determine the final concentration and loading time for TMRM or H2DCF-DA. In general, TMRM concentrations range from 20-200 nM, and the cell incubation time with TMRM varies from 20 to 60 min. The final concentration of H2DCF-DA ranges from 2-10 &mu;M, and incubation of cells in a loading solution containing this indicator varies from 30-45 min.
It is important to optimize the laser power and scan speed of taking the images to avoid both photo-toxicity to the cells and changes in the fluorescence intensity (for example flickering of TMRM fluorescence) in the absence of any stimulus. The optimized optical settings should result in a fluorescence signal that is not over or under saturated (threshold) in the absence of stimulus. The optimal conditions to collect the images from a selected field at a particular laser power and a scan speed are achieved when there are no changes in the fluorescence intensity of the probe in the absence of any stimulus for 10-15 min of live imaging.

Other fluorescence probes to determine &Delta;&Psi;m include rhodamine 123 and tetra methyl rhodamine ethyl ester (TMRE). However, they were found to inhibit the respiratory processes in isolated mitochondria2. Importantly, TMRM has no effect on mitochondrial respiration at low concentrations2 and has low phototoxicity and photobleaching3 compared with other probes. H2DCF-DA is a good indicator for ROS as it is well retained in the cells and recognizes several oxidant species, such as peroxides, super oxides, and nitric oxide4.

<table border="1">
 <tbody>
 <tr>
 <th>Name of reagent</th>
 <th>Company</th>
 <th>Catalogue number</th>
 </tr>
 <tr>
 <td>Glass bottom culture dish</td>
 <td>MatTek Corporation</td>
 <td>P35G-1.5-14-C</td>
 </tr>
 <tr>
 <td>NbActive4</td>
 <td>BrainBits</td>
 <td>NbActive4</td>
 </tr>
 <tr>
 <td>TMRM</td>
 <td>Invitrogen</td>
 <td>T668</td>
 </tr>
 <tr>
 <td>H2DCF-DA</td>
 <td>Invitrogen</td>
 <td>C400</td>
 </tr>
 <tr>
 <td>NaCl</td>
 <td>Sigma</td>
 <td>S6191</td>
 </tr>
 <tr>
 <td>KCl</td>
 <td>Sigma</td>
 <td>P3911</td>
 </tr>
 <tr>
 <td>CaCl2&bull;2H2O</td>
 <td>Sigma</td>
 <td>C3306</td>
 </tr>
 <tr>
 <td>MgCl2&bull;6H2O</td>
 <td>Sigma</td>
 <td>M2670</td>
 </tr>
 <tr>
 <td>D-Glucose</td>
 <td>Sigma</td>
 <td>G6152</td>
 </tr>
 <tr>
 <td>HEPES</td>
 <td>Invitrogen</td>
 <td>15630</td>
 </tr>
 </tbody>
</table>

determination of mitochondrial membrane potential and reactive oxygen species in live rat cortical neurons

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP. A significant loss of &Delta;&Psi;m renders cells depleted of energy with subsequent death. Reactive oxygen species (ROS) are important signaling molecules, but their accumulation in pathological conditions leads to oxidative stress. The two major sources of ROS in cells are environmental toxins and the process of oxidative phosphorylation. Mitochondrial dysfunction and oxidative stress have been implicated in the pathophysiology of many diseases; therefore, the ability to determine &Delta;&Psi;m and ROS can provide important clues about the physiological status of the cell and the function of the mitochondria. 
Several fluorescent probes (Rhodamine 123, TMRM, TMRE, JC-1) can be used to determine &Delta;&psi;m in a variety of cell types, and many fluorescence indicators (Dihydroethidium, Dihydrorhodamine 123, H2DCF-DA) can be used to determine ROS. Nearly all of the available fluorescence probes used to assess &Delta;&Psi;m or ROS are single-wavelength indicators, which increase or decrease their fluorescence intensity proportional to a stimulus that increases or decreases the levels of &Delta;&Psi;m or ROS. Thus, it is imperative to measure the fluorescence intensity of these probes at the baseline level and after the application of a specific stimulus. This allows one to determine the percentage of change in fluorescence intensity between the baseline level and a stimulus. This change in fluorescence intensity reflects the change in relative levels of &Delta;&Psi;m or ROS. In this video, we demonstrate how to apply the fluorescence indicator, TMRM, in rat cortical neurons to determine the percentage change in TMRM fluorescence intensity between the baseline level and after applying FCCP, a mitochondrial uncoupler. The lower levels of TMRM fluorescence resulting from FCCP treatment reflect the depolarization of mitochondrial membrane potential. We also show how to apply the fluorescence probe H2DCF-DA to assess the level of ROS in cortical neurons, first at baseline and then after application of H2O2. This protocol (with minor modifications) can be also used to determine changes in &#x2206;&#x03A8;m and ROS in different cell types and in neurons isolated from other brain regions.

Watch this Scientific Journal Video about Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons at JoVE.com

Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

We demonstrate application of the fluorescence indicator, TMRM, in cortical neurons to determine the relative changes in TMRM fluorescence intensity before and after application of a specific stimulus. We also show application of the fluorescence probe H2DCF-DA to assess the relative level of reactive oxygen species in cortical neurons.

Mitochondrial membrane potential (&Delta;&Psi;m) is critical for maintaining the physiological function of the respiratory chain to generate ATP.  A ...

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64.9K Views.  Loyola University Chicago. Dimostriamo applicazione del indicatore di fluorescenza, TMRM, nei neuroni corticali per determinare le variazioni relative a TMRM intensità di fluorescenza, prima e dopo l'applicazione di uno stimolo specifico. Mostriamo anche l'applicazione della sonda di fluorescenza H 2 DCF-DA per valutare il livello relativo di specie reattive dell'ossigeno nei neuroni corticali.

Video: Determination of Mitochondrial Membrane Potential and Reactive Oxygen Species in Live Rat Cortical Neurons

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured neurons for extended durations1-3. This is now possible through the use of vital dyes and fluorescent proteins with which cytoskeletal components, organelles, and other structures in living cells can be labeled and then visualized via dynamic fluorescence microscopy. For example, in embryonic chicken sympathetic neurons, mitochondrial movement was characterized using the vital dye rhodamine 1234. In another study, mitochondria were visualized in rat forebrain neurons by transfection of mitochondrially targeted eYFP5. However, imaging of primary neurons over minutes, hours, or even days presents a number of issues. Foremost among these are: 1) maintenance of culture conditions such as temperature, humidity, and pH during long imaging sessions; 2) a strong, stable fluorescent signal to assure both the quality of acquired images and accurate measurement of signal intensity during image analysis; and 3) limiting exposure times during image acquisition to minimize photobleaching and avoid phototoxicity.
Here, we describe a protocol that permits the observation, visualization, and analysis of mitochondrial movement in cultured hippocampal neurons with high temporal resolution and under optimal life support conditions. We have constructed an affordable stage-top incubator that provides good temperature regulation and atmospheric gas flow, and also limits the degree of media evaporation, assuring stable pH and osmolarity. This incubator is connected, via inlet and outlet hoses, to a standard tissue culture incubator, which provides constant humidity levels and an atmosphere of 5-10% CO2/air. This design offers a cost-effective alternative to significantly more expensive microscope incubators that don't necessarily assure the viability of cells over many hours or even days. To visualize mitochondria, we infect cells with a lentivirus encoding a red fluorescent protein that is targeted to the mitochondrion. This assures a strong and persistent signal, which, in conjunction with the use of a stable xenon light source, allows us to limit exposure times during image acquisition and all but precludes photobleaching and phototoxicity. Two injection ports on the top of the stage-top incubator allow the acute administration of neurotransmitters and other reagents intended to modulate mitochondrial movement. In sum, lentivirus-mediated expression of an organelle-targeted red fluorescent protein and the combination of our stage-top incubator, a conventional inverted fluorescence microscope, CCD camera, and xenon light source allow us to acquire time-lapse images of mitochondrial transport in living neurons over longer durations than those possible in studies deploying conventional vital dyes and off-the-shelf life support systems.

We describe a protocol that allows imaging of mitochondria in living neurons via fluorescence microscopy over long durations. Imaging over extended periods is accomplished through lentivirus-mediated expression of a mitochondrially targeted fluorescent protein and use of an inexpensive stage-top incubator that was designed and built in our laboratory.

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured ...

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 ...

Bilaminar Co-culture of Primary Rat Cortical Neurons and Glia

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or co-culture model. This system is suitable for a variety of experimental needs requiring either a glass or plastic growth substrate and can also be used for culture of other types of neurons.
Rat cortical neurons obtained from the late embryonic stage (E17) are plated on glass coverslips or tissue culture dishes facing a feeder layer of glia grown on dishes or plastic coverslips (known as Thermanox), respectively. The choice between the two configurations depends on the specific experimental technique used, which may require, or not, that neurons are grown on glass (e.g. calcium imaging versus Western blot). The glial feeder layer, an astroglia-enriched secondary culture of mixed glia, is separately prepared from the cortices of newborn rat pups (P2-4) prior to the neuronal dissection.
A major advantage of this culture system as compared to a culture of neurons only is the support of neuronal growth, survival, and differentiation provided by trophic factors secreted from the glial feeder layer, which more accurately resembles the brain environment in vivo. Furthermore, the co-culture can be used to study neuronal-glial interactions1.
At the same time, glia contamination in the neuronal layer is prevented by different means (low density culture, addition of mitotic inhibitors, lack of serum and use of optimized culture medium) leading to a virtually pure neuronal layer, comparable to other established methods1-3. Neurons can be easily separated from the glial layer at any time during culture and used for different experimental applications ranging from electrophysiology4, cellular and molecular biology5-8, biochemistry5, imaging and microscopy4,6,7,9,10. The primary neurons extend axons and dendrites to form functional synapses11, a process which is not observed in neuronal cell lines, although some cell lines do extend processes.
A detailed protocol of culturing rat hippocampal neurons using this co-culture system has been described previously4,12,13. Here we detail a modified protocol suited for cortical neurons. As approximately 20x106 cells are recovered from each rat embryo, this method is particularly useful for experiments requiring large numbers of neurons (but not concerned about a highly homogenous neuronal population). The preparation of neurons and glia needs to be planned in a time-specific manner. We will provide the step-by-step protocol for culturing rat cortical neurons as well as culturing glial cells to support the neurons.

Here we provide a protocol for culturing rat cortical neurons in the presence of a glial feeder layer. The cultured neurons establish polarity and create synapses, and can be separated from the glia for use in various applications, such as electrophysiology, calcium imaging, cell survival assays, immunocytochemistry, and RNA/DNA/protein isolation.

This video will guide you through the process of culturing rat cortical neurons in the presence of a glial feeder layer, a system known as a bilaminar or ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Membrane Potential Dye Imaging of Ventromedial Hypothalamus Neurons From Adult Mice to Study Glucose Sensing

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with increasing connective tissue and decreased neuronal viability that occur with age. Here, we describe a methodology for the dissociation of healthy hypothalamic neurons from adult-aged mice. The ability to study neurons from adult-aged mice allows the use of disease models that manifest at a later age and might be more developmentally accurate for certain studies. Fluorescence imaging of dissociated neurons can be used to study the activity of a population of neurons, as opposed to using electrophysiology to study a single neuron. This is particularly useful when studying a heterogeneous neuronal population in which the desired neuronal type is rare such as for hypothalamic glucose sensing neurons. We utilized membrane potential dye imaging of adult ventromedial hypothalamic neurons to study their responses to changes in extracellular glucose. Glucose sensing neurons are believed to play a role in central regulation of energy balance. The ability to study glucose sensing in adult rodents is particularly useful since the predominance of diseases related to dysfunctional energy balance (e.g.&#160;obesity) increase with age.

The activity of single neurons from adult-aged mice can be studied by dissociating neurons from specific brain regions and using fluorescent membrane potential dye imaging. By testing responses to changes in glucose, this technique can be used to study the glucose sensitivity of adult ventromedial hypothalamic neurons.

Studies of neuronal activity are often performed using neurons from rodents less than 2 months of age due to the technical difficulties associated with ...

Quantification of Filamentous Actin (F-actin) Puncta in Rat Cortical Neurons

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich structures suggest alterations in synaptic integrity and connectivity. Here we provide a detailed protocol for culturing primary rat cortical neurons, Phalloidin staining for F-actin puncta, and subsequent quantification techniques. First, the frontal cortex of E18 rat embryos are dissociated into low-density cell culture, then the neurons grown in vitro for at least 12-14 days. Following experimental treatment, the cortical neurons are stained with AlexaFluor 488 Phalloidin (to label the dendritic F-actin puncta) and microtubule-associated protein 2 (MAP2; to validate the neuronal cells and dendritic integrity). Finally, specialized software is used to analyze and quantify randomly selected neuronal dendrites. F-actin rich structures are identified on second order dendritic branches (length range 25-75 &#181;m) with continuous MAP2 immunofluorescence. The protocol presented here will be a useful method for investigating changes in dendritic synapse structures subsequent to experimental treatments.

Quantification of Filamentous Actin F-actin Puncta in Rat Cortical Neurons

Filamentous actin (F-actin) plays an important role in spinogenesis, synaptic plasticity, and synaptic stability. Quantification of F-actin puncta is therefore a useful tool to study the integrity of synaptic structures. This protocol describes the procedures of quantifying F-actin puncta labeled with Phalloidin in low-density primary cortical neuronal cultures.

Filamentous actin protein (F-actin) plays a major role in spinogenesis, synaptic plasticity, and synaptic stability. Changes in dendritic F-actin rich ...

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging methods exist for examining the brain tissue of live newborn mammals. Herein we summarize a protocol for imaging individual cortical neurons in living neonatal mice. This protocol includes the following two methodologies: (1) the Supernova system for sparse and bright labeling of cortical neurons in the developing brain, and (2) a surgical procedure for the fragile neonatal skull. This protocol allows the observation of temporal changes of individual cortical neurites during neonatal stages with a high signal-to-noise ratio. Labeled cell-specific gene silencing and knockout can also be achieved by combining the Supernova with RNA interference and CRISPR/Cas9 gene editing systems. This protocol can, thus, be used for analyzing the developmental dynamics of cortical neurons, molecular mechanisms that control the neuronal dynamics, and changes in neuronal dynamics in disease models.

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

We present an in vivo two-photon imaging protocol for imaging the cerebral cortex of neonatal mice. This method is suitable for analyzing the developmental dynamics of cortical neurons, the molecular mechanisms that control the neuronal dynamics, and the changes in neuronal dynamics in disease models.

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging ...

Stimulus-specific Cortical Visual Evoked Potential Morphological Patterns

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using 128-channel high-density electroencephalography (EEG). The specific aim of the described stimuli and analyses is to examine whether it is feasible to replicate previously reported CVEP morphological patterns elicited by an apparent motion stimulus, designed to simultaneously stimulate both ventral and dorsal central visual networks, using object and motion stimuli designed to separately stimulate ventral and dorsal visual cortical networks.&#160; Four visual paradigms are presented: 1. Randomized visual objects with consistent temporal presentation. 2. Randomized visual objects with inconsistent temporal presentation (or jitter).&#160; 3. Visual motion via a radial field of coherent central dot motion without jitter.&#160; 4. Visual motion via a radial field of coherent central dot motion with jitter.&#160; These four paradigms are presented in a pseudo-randomized order for each participant.&#160; Jitter is introduced in order to view how possible anticipatory-related effects may affect the morphology of the object-onset and motion-onset CVEP response.&#160; EEG data analyses are described in detail, including steps of data exportation from and importation to signal processing platforms, bad channel identification&#160;and removal, artifact rejection, averaging, and categorization of average CVEP morphological pattern type based upon latency ranges of component peaks. Representative data show that the methodological approach is indeed sensitive in eliciting differential object-onset and motion-onset CVEP morphological patterns and may, therefore, be useful in addressing the larger research aim. Given the high temporal resolution of EEG and the possible application of high-density EEG in source localization analyses, this protocol is ideal for the investigation of distinct CVEP morphological patterns and the underlying neural mechanisms which generate these differential responses.&#160; &#160; &#160;&#160;

In this paper, we present a protocol to investigate differential cortical visual evoked potential morphological patterns through stimulation of ventral and dorsal networks using high-density EEG. Visual object and motion stimulus paradigms, with and without temporal jitter, are described. Visual evoked potential morphological analyses are also outlined.&#160;&#160;

This paper presents a methodology for the recording and analysis of cortical visual evoked potentials (CVEPs) in response to various visual stimuli using ...

Generation of Neurospheres from Mixed Primary Hippocampal and Cortical Neurons Isolated from E14-E16 Sprague Dawley Rat Embryo

Primary neuron culture is an essential technique in the field of neuroscience. To gain deeper mechanistic insights into the brain, it is essential to have a robust in vitro model that can be exploited for various neurobiology studies. Though primary neuron cultures (i.e., long-term hippocampal cultures) have provided scientists with models, it does not yet represent the complexity of brain network completely. In the wake of these limitations, a new model has emerged using neurospheres, which bears a closer resemblance to the brain tissue. The present protocol describes the plating of high and low densities of mixed cortical and hippocampal neurons isolated from the embryo of embryonic day 14-16 Sprague Dawley rats. This allows for the generation of neurospheres and long-term primary neuron culture as two independent platforms to conduct further studies. This process is extremely simple and cost-effective, as it minimizes several steps and reagents previously deemed essential for neuron culture. This is a robust protocol with minimal requirements that can be performed with achievable results and further used for a diversity of studies related to neuroscience.

Presented here is a protocol for the spontaneous generation of neurospheres enriched in neural progenitor cells from high density plated neurons. During the same experiment, when neurons are plated at a lower density, the protocol also results in prolonged primary rat neuron cultures.

Primary neuron culture is an essential technique in the field of neuroscience. To gain deeper mechanistic insights into the brain, it is essential to have ...

Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Mitochondrial respiration is a critical energy-generating pathway in all cells, especially retinal photoreceptors that possess a highly active metabolism. In addition, photoreceptors also exhibit high aerobic glycolysis like cancer cells. Precise measurements of these metabolic activities can provide valuable insights into cellular homeostasis under physiological conditions and in disease states. High throughput microplate-based assays have been developed to measure mitochondrial respiration and various metabolic activities in live cells. However, a vast majority of these are developed for cultured cells and have not been optimized for intact tissue samples and for application ex vivo. Described here is a detailed step-by-step protocol, using microplate-based fluorescence technology, to directly measure oxygen consumption rate (OCR) as an indicator of mitochondrial respiration, as well as extracellular acidification rate (ECAR) as an indicator of glycolysis, in intact ex vivo retinal tissue. This method has been used to successfully assess metabolic activities in adult mouse retina and demonstrate its application in investigating cellular mechanisms of aging and disease.

Determination of Mitochondrial Respiration and Glycolysis in Ex Vivo Retinal Tissue Samples

Described here is a detailed protocol for performing mitochondrial stress assay and glycolytic rate assay in ex vivo retinal tissue samples using a commercial bioanalyzer.

Mitochondrial respiration is a critical energy-generating pathway in all cells, especially retinal photoreceptors that possess a highly active metabolism. ...