This work was supported by Ministerio de Econom&#236;a y competitividad (BFU2012-37146) and Junta de Castilla y Le&#242;n (BIO103/VA45/11, VA145U13 and BIO/VA33/13), Spain. MCR was supported by Junta de Castilla y Le&#242;n (Spain) and the European Social Fund. We thank the late Dr. Philippe Br&#251;let (1947-2013) for the mitochondrial GFP Aequorin plasmid.

Ethics Statement: Procedures involving animal subjects have been handled under protocols approved by the Valladolid University animal housing facility in agreement with the European Convention 123/Council of Europe and Directive 86/609/EEC.
1. Short and Long-term Culture of Rat Hippocampal Neurons
<ol>
	<li>Preparation of poly-D-lysine coated, 12 mm glass coverslips.
	<ol>
		<li>Sterilize 12 mm diameter glass coverslips in ethanol for at least 24 hr. Allow them to dry under sterile conditions.</li>
		<li>Distribute the coverslips on a strip of parafilm in a Petri dish. Cover the surface of each coverslip with 200 &#181;l of 1 mg/ml poly-D-lysine. Allow treating O/N.</li>
		<li>The next day wash the coverslips with double distilled sterile water every 15 min for 90 min under sterile conditions.</li>
		<li>Fill a four-well multidish plate with 500 &#181;l of Neurobasal Culture Medium per well. Neurobasal Culture Medium should be supplemented with 10% fetal bovine serum, 2 % B27, 1 &#181;g/ml gentamicin and 2 mM L-glutamine.</li>
		<li>Place the coverslips in the multidish plate. Maintain them in a humidified 37 &#186;C and 5% CO2 incubator until use.</li>
	</ol>
	</li>
	<li>Isolation of hippocampal neurons from neonatal rats.
	<ol>
		<li>Prepare 1.8 ml papain solution (directly purchased) in a vial (20 &#181;l/ml) under sterile conditions: To obtain a final concentration of 20 &#181;l/ml add around 40 &#181;l papain in 1.8 ml Hank&#39;s plus 0.6% BSA (&#181;l should be calculated depending on the supplier conditions). Hank&#39;s 0.6% BSA is prepared by mixing 85 ml of HBSS medium with 15 ml of 4% bovine serum albumin (BSA, prepared solving 4 g of BSA in 100 ml HBSS). Prepare it in a sterile hood.</li>
		<li>Incubate the papain in Hank&#39;s plus 0.6% BSA for 30 min at 37 &#186;C before use. Filter the solution using a 0.22 &#181;m filter prior to use.</li>
		<li>Prepare Ham&#39;s F-12 medium by adding Dulbecco&#39;s Modified Eagle&#39;s Medium powder (13.5 g per L) in 900 ml of double distilled sterile water. Then add 6 g HEPES and 336 mg NaHCO3 and adjust the pH to 7.42 using NaOH 4 N. Add sterile water in order to obtain a 1 L solution and filter it using a 0.20 &#181;m polyethersulfone (PES) bottle top filter. Carry this process in a sterile hood. Finally saturate the solution with CO2 under sterile conditions before use. Store solutions at 4 &#186;C and use them chilled.</li>
		<li>Euthanize newborn (P0) rat pups by decapitation and wash the head in sterile HBS medium. Open the skull with the help of sterile scissors. Extract the brain with a spatula and wash it quickly in Ham&#39;s F-12 medium.</li>
		<li>Make a diagonal cut with the aid of a scalpel in each hemisphere (Figure 1B), and carry it to a Petri dish containing Ham&#39;s F-12 medium.</li>
		<li>Discard meninges carefully with the help of dissection forceps and under a magnifying glass.</li>
		<li>Identify the hippocampus in a characteristic concave location over the cortex using magnifying glass. Then, separate the hippocampus from the cortex by pulling carefully from one border and removing it from its position using eye scissors.</li>
		<li>Transfer the hippocampal tissue to a 4-well plate filled with sterile Hank&#39;s medium lacking Ca2+ and Mg2+ plus 0.6% BSA and wash hippocampal tissue. Without removing the media cut the tissue in small pieces (around 2 x 2 mm) using the same eye scissors.</li>
		<li>Transfer hippocampal pieces to a vial containing 1.8 ml pre-filtered papain solution. Then incubate them at 37 &#186;C with occasional, gentle shaking. 15 min later, add 90 &#181;l of DNase I solution (50 &#181;g/ml final) and incubate for 15 additional min.</li>
		<li>Transfer the tissue to a 10 ml centrifuge tube and wash the fragments with fresh Neurobasal Culture Medium. Obtain cell suspension by passing tissue fragments through a 5 ml plastic pipette.</li>
		<li>Centrifuge cell suspension at 160 x g for 5 min. Remove the supernatant using a plastic sterile Pasteur pipette and suspend pellet carefully with a 1 ml automated pipette in 1 ml of Neurobasal Culture Medium.</li>
		<li>Measure cell density using a Neubauer counting chamber. Put the glass coverslip over the Neubauer chamber. Add 10 &#181;l of cell suspension. Make sure the cell suspension enters the chamber uniformly and making no bubbles. Count the number of cells under the microscope and perform corresponding calculations to obtain a suitable drop of 40-80 &#181;l containing 30 x 103 cells. The total number of cells will depend on the number of animals used.</li>
	</ol>
	</li>
	<li>Short-term and long-term culture of rat hippocampal neurons.
	<ol>
		<li>Over the four-well multidish plate containing 500 &#181;l of Neurobasal Culture Medium prepared before and kept in the incubator, plate around 30 x 103 cells in a drop of about 50 &#181;l to each well of the multidish plate containing one poly-D-lysine-coated, 12 mm diameter glass coverslip.</li>
		<li>Maintain primary hippocampal cells in a humidified 37 &#186;C and 5% CO2 incubator for 2-5 days in vitro (DIV, short-term, young cultures) or &#62;15 DIV (Long-term, aged cultures) before experiments without changing the culture media.</li>
	</ol>
	</li>
</ol>
2. Fluorescence Imaging of Cytosolic Ca 2+ Concentration
<ol>
	<li>Preparation of test solutions for fluorescence imaging.
	<ol>
		<li>Prepare an external HEPES-buffered saline (HBS) solution containing, in mM: NaCl 145, KCl 5, MgCl2 1, CaCl2 1, glucose 10, sodium-HEPES 10 (pH 7.42).</li>
		<li>Prepare an external, Mg2+-free, HEPES-buffered saline (HBS) solution containing, in mM: NaCl 146, KCl 5, CaCl2 1, glucose 10 and HEPES 10 (pH 7.42).</li>
		<li>Solve NMDA in Mg2+-free, HEPES-buffered saline (HBS) at a final concentration of 100 &#181;M and supplement it with 10 &#181;M glycine.</li>
		<li>Prepare a HBS solution containing KCl instead of NaCl by solving (in mM): KCl 145, MgCl2 1, CaCl2 1, glucose 10 and HEPES 10 (pH 7.42).</li>
	</ol>
	</li>
	<li>Loading of hippocampal cells with fluorescent calcium probe fura2/AM.
	<ol>
		<li>Take the cell culture containing coverslips off the incubator and wash them with HBS medium at RT by transferring them to a new four-well multidish plate containing 500 &#181;l of HBS per well.</li>
		<li>Incubate cells with fura2/AM 4 &#181;M (prepared in the same HBS medium) for 60 min at RT (25 &#186;C) in a dark place.</li>
		<li>After 60 min, wash coverslips with fresh HBS medium.</li>
	</ol>
	</li>
	<li>Recording fluorescence images of cytosolic [Ca 2+ ] in cultured cells.
	<ol>
		<li>Turn on the lamp, microscope, perfusion system, fluorescence camera and computer.</li>
		<li>Place the coverslips in a thermostated platform for open 12 mm glass coverslips on the stage of the inverted microscope and select a microscopic field using a 40X objective (oil, NA:1.3). Perfuse cells continuously with pre-warmed (37 &#186;C) HBS medium in absence or presence of test substances. Maintain the flow at a rate of about 5-10 ml/min. 
		Note: Cell perfusion system for living cells is mounted in a thermostated platform for open 12 mm glass coverslips. 8-lines gravity-driven perfusion system equipped with a valve controller is used to perfuse the solutions. A vacuum pump is responsible for removing any excess medium. Solutions are heated using an in-tube heating system.</li>
		<li>Capture a background image with the shutter closed at both excitation wavelengths.</li>
		<li>Epi-illuminate cells alternately at 340 and 380 nm. Record light emitted at 520 nm every 5-10 sec with a fluorescence camera, which is filtered by a fura-2 dichroic mirror.</li>
		<li>When the recording period is finished, store the complete sequence of images emitted at 520 nm in the computer for further analysis.</li>
	</ol>
	</li>
	<li>Analysis of recorded fluorescent images.
	<ol>
		<li>Open the experiment file. Using the aquacosmos software, click on &#39;Ratio&#39; and select the desired ratio range. Calculate the pixel by pixel ratio in the resulting images in order to obtain a sequence of ratio images.</li>
		<li>Subtract background by adjusting the &#39;background elimination&#39;. Press &#39;start calculation&#39;.</li>
		<li>Press &#39;all times sequence&#39; button and erase the ancient regions of interest (ROIs). For quantitative analysis of individual cells, establish new regions of interest or ROIs corresponding to individual neurons. Average all ratio values in each pixel corresponding to each ROI and each image to obtain a recording of ratio fluorescence values for individual ROIs (cells).</li>
		<li>To graph the individual recordings export the ratio fluorescence values corresponding to each ROI to a graphing program.</li>
		<li>Make the corresponding calculations for estimating the size of the rises in ratio fluorescences in response to each stimulation using a suitable data analysis and graphing software.</li>
	</ol>
	</li>
</ol>
3. Bioluminescence Imaging of Mitochondrial Ca 2+ Concentration
<ol>
	<li>Transfection of cultured hippocampal neurons with the mitGAmut plasmid. 
	NOTE: Cells are first transfected with the mitGAmut plasmid containing a mitochondria-targeted sequence and a mutated aequorin lacking a Ca2+ binding site fused to the green fluorescent protein (GFP). This construction detects large rises in mitochondrial Ca2+ uptake and allows the identification of transfected cells by the GFP fluorescence. The plasmid was developed and kindly provided by P. Br&#251;let and co-workers 18 (CNRS, Paris).
	<ol>
		<li>Prepare Neurobasal TF, containing Neurobasal Culture Medium, lacking fetal bovine serum, B27, gentamicin and 2 mM L-glutamine.</li>
		<li>Prepare 50 &#181;l of Neurobasal TF plus 2.5 &#181;l transfection reagent in a vial (Solution A). Prepare 50 &#181;l of Neurobasal TF plus 4 &#181;g mitGAmut plasmid in a vial (Solution B). Add gently solution B over solution A. Allow mixing during 20 min, without shaking.</li>
		<li>Transfer the coverslips containing hippocampal neurons to new multidish plates containing 200 &#181;l of fresh Neurobasal TF.</li>
		<li>Add drop by drop 100 &#181;l of solution A + B over every coverslip. Incubate for 30 min. Remove the medium. Wash once cells with fresh Neurobasal TF. Return the coverslips to the original Neurobasal Culture Medium.</li>
		<li>Culture neurons for an additional 24 hr at 37 &#186;C and 5% CO2 after transfection to allow efficient expression and targeting of the probe.</li>
	</ol>
	</li>
	<li>Preparation of test solutions for bioluminescence imaging.
	<ol>
		<li>Prepare HBS solutions containing NMDA 100 &#181;M or 145 mM KCl as above (Step 2.1). Prepare HBS solution containing 10 mM Ca2+ plus digitonin 100 &#181;M.</li>
	</ol>
	</li>
	<li>Reconstitution of mitochondria-targeted aequorin with coelenterazine n.
	<ol>
		<li>Transfer coverslips containing transfected hippocampal neurons to a four-well plate containing 200 &#181;l of HBS.</li>
		<li>Add 4 &#181;l of coelenterazine n (200 &#181;M) to 200 &#181;l of HBS for a final concentration of 4 &#181;M and mix gently. Incubate for 2 hr at RT (25 &#186;C) and in the dark.</li>
	</ol>
	</li>
	<li>Recording bioluminescence images of mitochondrial [Ca 2+ ].
	<ol>
		<li>Turn on the microscope, the lamp, the perfusion system, bioluminescence camera and the computer.</li>
		<li>Transfer reconstituted coverslips to a thermostated (37 &#186;C) perfusion chamber and perfuse continuously with pre-warmed HBS solution at a rate of 5-10 ml/min.</li>
		<li>Using a 40X objective (oil, NA: 1.3), select a microscopic field containing cells expressing the apoaequorins as shown by fluorescence emission associated with expression of green fluorescent protein (GFP) using the FITC filters. A typical field may contain 1-2 transfected cells. Capture the GFP fluorescence image, using a CCD camera.</li>
		<li>Turn off the microscope light. Remove the dichroic-containing box located in the light pathway. Turn off the excitation light and close the dark-box for complete darkness.</li>
		<li>Perfuse cells at 5-10 ml/min with HBS medium with or without test solutions pre-warmed at 37 &#186;C.</li>
		<li>Capture photonic emission images every 10 sec with a photon-counting camera handled with an image processor.</li>
		<li>At the end of each experiment, permeabilize cells with 0.1 mM digitonin in 10 mM CaCl2 in HBS in order to release all remaining photonic emissions. 
		Note: These photonic emissions must be added up in order to estimate the total photonic emissions, a value required for calibration in Ca2+ concentrations.</li>
		<li>Store bioluminescence images in the computer with the GFP associated fluorescence image captured before the photon counting imaging.</li>
	</ol>
	</li>
	<li>Analysis of bioluminescence images reflecting mitochondrial [Ca 2+ ].
	<ol>
		<li>Quantify photonic emissions using specific software. Photonic emissions can be converted to mitochondrial free Ca2+ concentration ([Ca2+]mit) using the algorithm described by Brini et al. 15.</li>
		<li>Open the experiment and the GFP fluorescence image. Select regions of interest (ROIs) with the help of specific software by drawing the shape of the transfected cells according to the fluorescence images captured at the beginning of the experiment.</li>
		<li>Paste the same ROIs on every image. Total photonic emissions in each ROI are computed with specific software to obtain the luminescence emission value (L) for each cell at each point in time.</li>
		<li>Add up all the photonic emissions in the bioluminescence images, including the ones obtained after digitonin permeabilization, using the specific software, to obtain a bioluminescence image containing all the photonic emissions.</li>
		<li>Export values of photonic emissions for every region of interest to a specific software for calculations. To do this, click on &#39;Graph&#39; in order to compute the bioluminescence values for every ROI at each time. Select &#39;total value&#39; and &#39;use current ROI to all images. Press &#39;calculate&#39;. Save the &#39;txt.file&#39; and export data to a worksheet. For every ROI, [Ca2+]mit is calculated using the following algorithm: 
		 
		[Ca2+](M) = [R + (R &#183; KTR) - 1] / [KR - (R &#183; KR)]; 
		 
		where, 
		R = [L/(Ltotal&#955;)]1/n 
		L is the luminescence emitted at the time of measurement in each region of interest. 
		Ltotal is the total luminescence remaining after addition of the counts present in the tissue at that time at the time of measurement for each region of interest. &#955;, KTR, KR and n are constants whose values depend on combination of aequorin (wild type or mutated) and coelenterazin used (wild type or n type) as well as the recording temperature 16. For the combination used here (mutated AEQ and coelenterzine n), at 37 &#186;C, we used the following values: KR = 8.47 x 107, KTR = 165.6 x 103, n = 1,204 and &#955; = 0,138.</li>
	</ol>
	</li>
	<li>Make the corresponding calculations for estimating the size of the rises in bioluminescence in response to each stimulus using a suitable data analysis and graphing software.</li>
</ol>

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None of the authors have competing interests or conflicting interests.

The remodeling of intracellular Ca2+ homeostasis in the aging brain has been related to cognitive loss, increased susceptibility to ischemic damage, excitotoxicity and neurodegeneration. To investigate this hypothesis in vitro, Ca2+ imaging procedures are available. Unfortunately, viable cultures of old hippocampal neurons are not reliable. Recently, it has been observed that long-term cultures of rat hippocampal neurons present many of the typical hallmarks of aging in vivo includ...

Excitotoxicity is one of the most important mechanisms contributing to neuronal damage and cell death in neurological insults such as ischemia, and in some neurodegenerative diseases such as Alzheimer&#39;s disease 1. This type of neurotoxicity is mainly mediated by glutamate acting on Ca2+-permeable, ionotropic NMDA receptors (NMDAR) 2. Exposure of cultured neurons to glutamate can lead to excitotoxicity 3, which causes neuronal apoptosis 4. We and others have previously reported that neuronal vulnerability to NMDA-induced apoptosis may change with development in vitro and aging 5-8.
It is widely accepted that an increase in the cytosolic-free Ca2+ concentration ([Ca2+]cyt) leads to cells activation. However, if this rise is too high and/or sustained enough, it can trigger cell death 9. Moreover, it has been proposed that excitotoxicity requires mitochondrial Ca2+ uptake 10, since treating neurons with a mitochondrial uncoupler protected neurons against glutamate-induced cell death 11. If mitochondria take up too much Ca2+, the opening of the mitochondrial permeability transition pore may occur, leading to release of cytochrome c and other pro-apoptotic factors, and inducing apoptosis. We have recently shown that this mitochondrial Ca2+ uptake is directly related to the age-dependant susceptibility to excitotoxicity, by directly measuring NMDA-induced mitochondrial Ca2+ uptake in single hippocampal neurons 5, a method which is reported in this article. The hippocampus, involved in physiological processes such as learning, memory and other cognitive processes 12, is highly vulnerable to aging and neurodegenerative disorders 13. It has been proposed that, after several weeks in vitro, cultured hippocampal neurons show a number of typical characteristics of aged neurons 14. Accordingly, long-term cultured hippocampal neurons may provide a comprehensive model to investigate Ca2+-mediated mechanisms of enhanced excitotoxicity in aging.
The overall goal of the method presented is, therefore, to investigate substantial changes in intracellular Ca2+ homeostasis or Ca2+ remodeling in the aging brain including the differential Ca2+ responses elicited by NMDA receptor agonists in a long-term cultured hippocampal neurons. The method includes a detailed description of the culture of rat hippocampal neurons and the monitoring of cytosolic and mitochondrial Ca2+ concentrations by fluorescence and bioluminescence imaging in individual neurons, respectively. Fluorescence imaging of cytosolic Ca2+ in cultured neurons is a standard procedure. However, this method is less reliable for subcellular Ca2+ measurements including mitochondrial Ca2+. Reasons for this include lack of proper targeting of synthetic probes and inappropriate affinity for Ca2+ concentrations that may change in mitochondria from the low &#181;M level even to the mM level. The use of Ca2+ probes based on proteins as for instance aequorin, has allowed the targeting to subcellular organelles and the use of derivatives different Ca2+ affinities using different coelenterazines or mutated probes lacking specific Ca2+ binding sites 15. In this way, bioluminescence imaging of cells expressing mitochondria-targeted aequorin may allow the monitoring of mitochondrial Ca2+ concentrations in individual neurons. Yet, this procedure may require the use of photon counting cameras or ultrasensitive CCD cameras for bioluminescence imaging 16-18. This method may yield novel results that should be confirmed in more established brain aging models as, for instance, brain slices from old animals.

<table>
	<tbody>
		<tr>
			<td>Neurobasal Culture Medium</td>
			<td>Gibco</td>
			<td>21103-049</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS medium</td>
			<td>Gibco</td>
			<td>14170-088</td>
			<td></td>
		</tr>
		<tr>
			<td>Ham&#39;s F-12 medium</td>
			<td>Gibco</td>
			<td>31330-038</td>
			<td></td>
		</tr>
		<tr>
			<td>DNase I (from bovine pancreas)</td>
			<td>Sigma</td>
			<td>D5025-15KU</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal Bobine Serum</td>
			<td>Lonza</td>
			<td>DE14-801E</td>
			<td></td>
		</tr>
		<tr>
			<td>B27</td>
			<td>Gibco</td>
			<td>17504-044</td>
			<td></td>
		</tr>
		<tr>
			<td>Gentamicin</td>
			<td>Gibco</td>
			<td>15750</td>
			<td></td>
		</tr>
		<tr>
			<td>L-glutamine</td>
			<td>Gibco</td>
			<td>25030-032</td>
			<td></td>
		</tr>
		<tr>
			<td>12 mm glass coverslips</td>
			<td>Labolan</td>
			<td>0111520/20012</td>
			<td></td>
		</tr>
		<tr>
			<td>Papain</td>
			<td>Worthington</td>
			<td>LS003127</td>
			<td></td>
		</tr>
		<tr>
			<td>4-well multidish plaques</td>
			<td>Nunc</td>
			<td>176740</td>
			<td></td>
		</tr>
		<tr>
			<td>Petry dishes</td>
			<td>JD Catalan s.l.</td>
			<td>2120044T</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile pipettes</td>
			<td>Fisher Scientific</td>
			<td>431030</td>
			<td></td>
		</tr>
		<tr>
			<td>Fura2-AM</td>
			<td>Life Technologies</td>
			<td>F1201</td>
			<td></td>
		</tr>
		<tr>
			<td>Lipofectamine2000</td>
			<td>Invitrogen</td>
			<td>11668-027</td>
			<td></td>
		</tr>
		<tr>
			<td>Coelenterazine n</td>
			<td>Biotium</td>
			<td>BT-10115-2250 uG</td>
			<td></td>
		</tr>
		<tr>
			<td>Digitonin</td>
			<td>Sigma</td>
			<td>D5628</td>
			<td></td>
		</tr>
		<tr>
			<td>NMDA</td>
			<td>Sigma&#160;</td>
			<td>M3262</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycine</td>
			<td>Sigma</td>
			<td>50046</td>
			<td></td>
		</tr>
		<tr>
			<td>Zeiss Axiovert S100 TV microscope</td>
			<td>Carl Zeiss Inc.</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Xcite ilumination system</td>
			<td>EXFO</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ORCA ER fluorescence camera</td>
			<td>Hamamatsu</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>VIM photon counting CCD camera</td>
			<td>Hamamatsu</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>VC-8 valve controller</td>
			<td>Warner Instruments</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>SH-27B heating system&#160;</td>
			<td>Warner Instruments</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Aquacosmos Software</td>
			<td>Hamamatsu Photonics</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

fluorescence and bioluminescence imaging of subcellular ca2+ in aged hippocampal neurons

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible are badly known. Excitotoxicity, a process believed to contribute to neuron damage induced by both insults, is mediated by activation of glutamate receptors that promotes Ca2+ influx and mitochondrial Ca2+ overload. A substantial change in intracellular Ca2+ homeostasis or remodeling of intracellular Ca2+ homeostasis may favor neuron damage in old neurons. For investigating Ca2+ remodeling in aging we have used live cell imaging in long-term cultures of rat hippocampal neurons that resemble in some aspects aged neurons in vivo. For this end, hippocampal cells are, in first place, freshly dispersed from new born rat hippocampi and plated on poli-D-lysine coated, glass coverslips. Then cultures are kept in controlled media for several days or several weeks for investigating young and old neurons, respectively. Second, cultured neurons are loaded with fura2 and subjected to measurements of cytosolic Ca2+ concentration using digital fluorescence ratio imaging. Third, cultured neurons are transfected with plasmids expressing a tandem of low-affinity aequorin and GFP targeted to mitochondria. After 24 hr, aequorin inside cells is reconstituted with coelenterazine and neurons are subjected to bioluminescence imaging for monitoring of mitochondrial Ca2+ concentration. This three-step procedure allows the monitoring of cytosolic and mitochondrial Ca2+ responses to relevant stimuli as for example the glutamate receptor agonist NMDA and compare whether these and other responses are influenced by aging. This procedure may yield new insights as to how aging influence cytosolic and mitochondrial Ca2+ responses to selected stimuli as well as the testing of selected drugs aimed at preventing neuron cell death in age-related diseases.

Here we describe a simple method to assess Ca2+ remodeling and the effects of NMDA on cytosolic and mitochondrial [Ca2+] in aged neurons. Figure 1 shows the schematic of the procedure for isolating and culturing hippocampal neurons from neonatal rats. Before starting, we need to prepare sterile, poly-D-lysine coated, glass coverslips and locate them in a 4-well dish. Then, neonatal rats are killed and the brain removed. After isolating the hippocampus, tissue is carefully dispersed ...

Watch this Scientific Journal Video about Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons at JoVE.com

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Intracellular Ca2+ remodeling in aging may contribute to excitotoxicity and neuron damage, processes mediated by Ca2+ overload. We aimed at investigating Ca2+ remodeling in the aging brain using fluorescence and bioluminescence imaging of cytosolic and mitochondrial Ca2+ in long-term cultures of rat hippocampal neurons, a model of neuronal aging.

Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Susceptibility to neuron cell death associated to neurodegeneration and ischemia are exceedingly increased in the aged brain but mechanisms responsible ...

fluorescence-bioluminescence-imaging-subcellular-ca2-aged-hippocampal

Research

JoVE Journal

Neuroscience

8.3K 조회수.  Consejo Superior de Investigaciones Cientìficas. Intracellular Ca2+ remodeling in aging may contribute to excitotoxicity and neuron damage, processes mediated by Ca2+ overload. We aimed at investigating Ca2+ remodeling in the aging brain using fluorescence and bioluminescence imaging of cytosolic and mitochondrial Ca2+ in long-term cultures of rat hippocampal neurons, a model of neuronal aging. 

비디오: Fluorescence and Bioluminescence Imaging of Subcellular Ca2+ in Aged Hippocampal Neurons

Nucleofection and Primary Culture of Embryonic Mouse Hippocampal and Cortical Neurons

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and synapse formation and function. An advantage of culturing these neurons is that they readily polarize, forming distinctive axons and dendrites, on a two dimensional substrate at very low densities. This property has made them extremely useful for determining many aspects of neuronal development. Furthermore, by providing glial conditioning for these neurons they will continue to develop, forming functional synaptic connections and surviving for several months in culture. In this protocol we outline a technique to dissect, culture and transfect embryonic mouse hippocampal and cortical neurons. Transfection is accomplished by electroporating DNA into the neurons before plating via nucleofection. This protocol has the advantage of expressing fluorescently-tagged fusion proteins early in development (~4-8hrs after plating) to study the dynamics and function of proteins during polarization, axon outgrowth and branching. We have also discovered that this single transfection before plating maintains fluorescently-tagged fusion protein expression at levels appropriate for imaging throughout the lifetime of the neuron (&gt; 2 months in culture). Thus, this methodology is useful for studying protein localization and function throughout CNS development with little or no disruption of neuronal function.

This protocol outlines the steps required to dissect, transfect via electroporation and culture mouse hippocampal and cortical neurons. Short-term cultures may be used for studies of axon outgrowth and guidance, while long-term cultures can be used for studies of synaptogenesis and dendritic spine analysis.

Hippocampal and cortical neurons have been used extensively to study central nervous system (CNS) neuronal polarization, axon/dendrite outgrowth, and ...

Fluorescence Recovery After Photobleaching (FRAP) of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached in the region of interest. The fluorescence of the region recovers when the unbleached GFP-tagged protein from outside of the region diffuses into the region of interest. The mobility of the protein is then analyzed by measuring the fluorescence recovery rate. This technique could be used to characterize protein mobility and turnover rate.
In this study, we express the (enhanced green fluorescent protein) EGFP vector in cultured hippocampal neurons. Using the Zeiss 710 confocal microscope, we photobleach the fluorescence signal of the GFP protein in a single spine, and then take time lapse images to record the fluorescence recovery after photobleaching. Finally, we estimate the percentage of mobile and immobile fractions of the GFP in spines, by analyzing the imaging data using ImageJ and Graphpad softwares.
This FRAP protocol shows how to perform a basic FRAP experiment as well as how to analyze the data.

Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of Green Fluorescence Protein (GFP)-tagged proteins in cultured cells. We examined the mobile/immobile fractions of the GFP by analyzing the fluorescence recovery percentage after photobleaching. In this study, FRAP was performed at spines of hippocampal neurons.

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached ...

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 ...

Isolation and Culture of Hippocampal Neurons from Prenatal Mice

Primary cultures of rat and murine hippocampal neurons are widely used to reveal cellular mechanisms in neurobiology. By isolating and growing individual neurons, researchers are able to analyze properties related to cellular trafficking, cellular structure and individual protein localization using a variety of biochemical techniques. Results from such experiments are critical for testing theories addressing the neural basis of memory and learning. However, unambiguous results from these forms of experiments are predicated on the ability to grow neuronal cultures with minimum contamination by other brain cell types. In this protocol, we use specific media designed for neuron growth and careful dissection of embryonic hippocampal tissue to optimize growth of healthy neurons while minimizing contaminating cell types (i.e. astrocytes). Embryonic mouse hippocampal tissue can be more difficult to isolate than similar rodent tissue due to the size of the sample for dissection. We show detailed dissection techniques of hippocampus from embryonic day 19 (E19) mouse pups. Once hippocampal tissue is isolated, gentle dissociation of neuronal cells is achieved with a dilute concentration of trypsin and mechanical disruption designed to separate cells from connective tissue while providing minimum damage to individual cells. A detailed description of how to prepare pipettes to be used in the disruption is included. Optimal plating densities are provided for immuno-fluorescence protocols to maximize successful cell culture. The protocol provides a fast (approximately 2 hr) and efficient technique for the culture of neuronal cells from mouse hippocampal tissue.

We provide a protocol for the culture of highly purified hippocampal neurons from prenatal mouse brains without the use of a feeder glial cell layer.

Primary cultures of rat and murine hippocampal neurons are widely used to reveal cellular mechanisms in neurobiology. By isolating and growing individual ...

Assaying DNA Damage in Hippocampal Neurons Using the Comet Assay

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been extensively evaluated. Traditional methods are time consuming, expensive, resource intensive and require replicating cells. In contrast, the comet assay, a single cell gel electrophoresis assay, is a faster, non-invasive, inexpensive, direct and sensitive measure of DNA damage and repair. All forms of DNA damage as well as DNA repair can be visualized at the single cell level using this powerful technique.
	The principle underlying the comet assay is that intact DNA is highly ordered whereas DNA damage disrupts this organization. The damaged DNA seeps into the agarose matrix and when subjected to an electric field, the negatively charged DNA migrates towards the cathode which is positively charged. The large undamaged DNA strands are not able to migrate far from the nucleus. DNA damage creates smaller DNA fragments which travel farther than the intact DNA. Comet Assay, an image analysis software, measures and compares the overall fluorescent intensity of the DNA in the nucleus with DNA that has migrated out of the nucleus. Fluorescent signal from the migrated DNA is proportional to DNA damage. Longer brighter DNA tail signifies increased DNA damage. Some of the parameters that are measured are tail moment which is a measure of both the amount of DNA and distribution of DNA in the tail, tail length and percentage of DNA in the tail. This assay allows to measure DNA repair as well since resolution of DNA damage signifies repair has taken place. The limit of sensitivity is approximately 50 strand breaks per diploid mammalian cell 1,2. Cells treated with any DNA damaging agents, such as etoposide, may be used as a positive control. Thus the comet assay is a quick and effective procedure to measure DNA damage.

The comet assay is an efficient way of detecting single- and double-strand breaks, including alkali-labile sites and DNA-DNA/DNA-protein cross-links on the DNA in all cells including hippocampal neurons. The method takes advantage of the differential migration of DNA in an electric field due to differences in amount of DNA damage.

A number of drugs target the DNA repair pathways and induce cell kill by creating DNA damage. Thus, processes to directly measure DNA damage have been ...

Calcium Phosphate Transfection of Primary Hippocampal Neurons

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this method on hard-to-transfect cells like primary neurons. Here we describe our detailed protocol for calcium phosphate transfection of hippocampal neurons cocultured with astroglial cells.

Calcium phosphate precipitation is a convenient and economical method for transfection of cultured cells. With optimization, it is possible to use this ...

Imaging Dendritic Spines of Rat Primary Hippocampal Neurons using Structured Illumination Microscopy

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. Technological advances have identified these structures as key elements in neuron connectivity and synaptic plasticity. The quantitative analysis of spine morphology using light microscopy remains an essential problem due to technical limitations associated with light's intrinsic refraction limit. Dendritic spines can be readily identified by confocal laser-scanning fluorescence microscopy. However, measuring subtle changes in the shape and size of spines is difficult because spine dimensions other than length are usually smaller than conventional optical resolution fixed by light microscopy's theoretical resolution limit of 200 nm.
Several recently developed super resolution techniques have been used to image cellular structures smaller than the 200 nm, including dendritic spines. These techniques are based on classical far-field operations and therefore allow the use of existing sample preparation methods and to image beyond the surface of a specimen. Described here is a working protocol to apply super resolution structured illumination microscopy (SIM) to the imaging of dendritic spines in primary hippocampal neuron cultures. Possible applications of SIM overlap with those of confocal microscopy. However, the two techniques present different applicability. SIM offers higher effective lateral resolution, while confocal microscopy, due to the usage of a physical pinhole, achieves resolution improvement at the expense of removal of out of focus light. In this protocol, primary neurons are cultured on glass coverslips using a standard protocol, transfected with DNA plasmids encoding fluorescent proteins and imaged using SIM. The whole protocol described herein takes approximately 2 weeks, because dendritic spines are imaged after 16-17 days in vitro, when dendritic development is optimal. After completion of the protocol, dendritic spines can be reconstructed in 3D from series of SIM image stacks using specialized software.

This article describes a working protocol to image dendritic spines from hippocampal neurons in vitro using Structured Illumination Microscopy (SIM). Super-resolution microscopy using SIM provides image resolution significantly beyond the light diffraction limit in all three spatial dimensions, allowing the imaging of individual dendritic spines with improved detail.

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. ...

In Situ Ca2+ Imaging of the Enteric Nervous System

Reflex behaviors of the intestine are controlled by the enteric nervous system (ENS). The ENS is an integrative network of neurons and glia in two ganglionated plexuses housed in the gut wall. Enteric neurons and enteric glia are the only cell types within the enteric ganglia. The activity of enteric neurons and glia is responsible for coordinating intestinal functions. This protocol describes methods for observing the activity of neurons and glia within the intact ENS by imaging intracellular calcium (Ca2+) transients with fluorescent indicator dyes. Our technical discussion focuses on methods for Ca2+ imaging in whole-mount preparations of the myenteric plexus from the rodent bowel. Bulk loading of ENS whole-mounts with a high-affinity Ca2+ indicator such as Fluo-4 permits measurements of Ca2+ responses in individual neurons or glial cells. These responses can be evoked repeatedly and reliably, which permits quantitative studies using pharmacological tools. Ca2+ responses in cells of the ENS are recorded using a fluorescence microscope equipped with a cooled charge-coupled device (CCD) camera. Fluorescence measurements obtained using Ca2+ imaging in whole-mount preparations offer a straightforward means of characterizing the mechanisms and potential functional consequences of Ca2+ responses in enteric neurons and glial cells.

In Situ Ca2+ Imaging of the Enteric Nervous System

The enteric nervous system (ENS) is a network of neurons and glia located in the gut wall that controls intestinal reflexes. This protocol describes methods for recording the activity of enteric neurons and glia in live preparations of ENS using Ca2+ imaging.

Reflex behaviors of the intestine are controlled by the enteric nervous system (ENS). The ENS is an integrative network of neurons and glia in two ...

Assessment of Hippocampal Dendritic Complexity in Aged Mice Using the Golgi-Cox Method

Dendritic spines are the protuberances from the neuronal dendritic shafts that contain&#160; excitatory synapses. The morphological and branching variations of the neuronal dendrites within the hippocampus are implicated in cognition and memory formation. There are several approaches to Golgi staining, all of which have been useful for determining the morphological characteristics of dendritic arbors and produce a clear background. The present Golgi-Cox method, (a slight variation of the protocol that is provided with a commercial Golgi staining kit), was designed to assess how a relatively low dose of the chemotherapeutic drug 5-flurouracil (5-Fu) would affect dendritic morphology, the number of spines, and the complexity of arborization within the hippocampus. The 5-Fu significantly modulated the dendritic complexity and decreased the spine density throughout the hippocampus in a region-specific manner. The data presented show that the Golgi staining method effectively stained the mature neurons in the CA1, the CA3, and the dentate gyrus (DG) of the hippocampus. This protocol reports the details for each step so that other researchers can reliably stain tissue throughout the brain with high quality results and minimal troubleshooting.

Here we present a Golgi-Cox protocol in extensive detail. This reliable tissue stain method allows for a high-quality assessment of the cytoarchitecture in the hippocampus, and throughout the entire brain, with minimal troubleshooting.

Dendritic spines are the protuberances from the neuronal dendritic shafts that contain&#160; excitatory synapses. The morphological and branching ...

Measuring Synaptic Vesicle Endocytosis in Cultured Hippocampal Neurons

During endocytosis, fused synaptic vesicles are retrieved at nerve terminals, allowing for vesicle recycling and thus the maintenance of synaptic transmission during repetitive nerve firing. Impaired endocytosis in pathological conditions leads to decreases in synaptic strength and brain functions. Here, we describe methods used to measure synaptic vesicle endocytosis at the mammalian hippocampal synapse in neuronal culture. We monitored synaptic vesicle protein endocytosis by fusing a synaptic vesicular membrane protein, including synaptophysin and VAMP2/synaptobrevin, at the vesicular lumenal side, with pHluorin, a pH-sensitive green fluorescent protein that increases its fluorescence intensity as the pH increases. During exocytosis, vesicular lumen pH increases, whereas during endocytosis vesicular lumen pH is re-acidified. Thus, an increase of pHluorin fluorescence intensity indicates fusion, whereas a decrease indicates endocytosis of the labelled synaptic vesicle protein. In addition to using the pHluorin imaging method to record endocytosis, we monitored vesicular membrane endocytosis by electron microscopy (EM) measurements of Horseradish peroxidase (HRP) uptake by vesicles. Finally, we monitored the formation of nerve terminal membrane pits at various times after high potassium-induced depolarization. The time course of HRP uptake and membrane pit formation indicates the time course of endocytosis.

Synaptic vesicle endocytosis is detected by light microscopy of pHluorin fused with synaptic vesicle protein and by electron microscopy of vesicle uptake.