Work in the lab is supported by Nederlandse Organisatie voor Wetenschappelijk Onderzoek 	(NWO) (917.10.372) to RMM. JD is affiliated to the EU FP7 BrainTrain programme (<a href="http://www.brain-train.nl">www.brain-train.nl</a>). We thank Pieter Laurens-Baljon and Sabine Schmitz (both CNCR, VU University Amsterdam) for images of FP multielectrode array waveforms and neuronal cultures used in the video sequence.

1. Making horizontal entorhinal-hippocampal brain slices
Entorhinal-hippocampal brain slices are made using anatomical guides, detailed in a 3D overview of the region according to Canto, Wouterlood &amp; Witter (2008) Neural Plasticity ID 3812439.
<ol>
 <li>Make 500ml of slice solution, oxygenate with carbogen gas (95% oxygen, 5% carbon dioxide) for at least 20 minutes and then freeze 250ml until iced. Minutes before slicing, crush and blend the slice ice with the remaining 250ml bubbling slice solution to be used for dissection and for cutting in the slicing chamber.</li>
 <li>Make 200ml of r-ACSF (recovery ACSF) and at least 500ml of e-ACSF (experimental ACSF) solution. Continuously oxygenate both solutions with carbogen gas (95% oxygen, 5% carbon dioxide) for at least 20 minutes before tissue preparation.</li>
 <li>This protocol is valid for mice from postnatal day 0 up to a maximum of 3 weeks old. Decapitate the animal, according to local ethical committee procedures, and dissect out the brain quickly in a Petri dish containing ice-cold slice solution prepared in Step 1, above (see Table 1 for solutions &amp; reagents).</li>
 <li>Transfer the brain onto a filter paper soaked with slice solution and separate the two hemispheres with a sharp razor blade. Transfer one hemisphere back into ice-cold slice solution.</li>
 <li>Remove the cerebellum of the other hemisphere. Flip the brain onto its midline and cut off the top of the brain with a slight angle towards the rostral end.</li>
 <li>Flip the brain onto the cut (dorsal) surface and glue it onto the tissue holder of the slicer upside down by gently pushing with a cotton bud.</li>
 <li>Slice 300 &mu;m thick brain slices using a low frequency and speed in the remaining ice-cold slice solution made in Step 1. Under our conditions, we use a Thermo Fisher Scientific vibratome (Microm, HM 650V) at settings 0.05 mm/s and 36Hz.</li>
 <li>As soon as a slice is cut, transfer it into a slice holder (submerged) containing oxygenated Artificial Cerebro Spinal Fluid (ACSF) (2.5mM Mg2+, 1.6mM Ca2+; see Table 2) at room temperature. A higher magnesium concentration in r-ACSF reduces the influx of calcium into neurons via NMDA receptors following slicing and in our experience, leads to a better quality of slices for experiments. If needed, cut the second cerebral hemisphere afterwards.</li>
 <li>Leave the slices for one hour to recover.</li>
</ol>
<table border="1">
 <tbody>
 <tr>
 <td colspan="2">Slice solution (in mM) &#x2013; 10</td>
 </tr>
 <tr>
 <td>110</td>
 <td>choline chloride</td>
 </tr>
 <tr>
 <td>25</td>
 <td>NaHCO3</td>
 </tr>
 <tr>
 <td>11.6</td>
 <td>11.6</td>
 </tr>
 <tr>
 <td>10</td>
 <td>D-glucose</td>
 </tr>
 <tr>
 <td>7</td>
 <td>MgCl2</td>
 </tr>
 <tr>
 <td>3.1</td>
 <td>sodiumpyruvate</td>
 </tr>
 <tr>
 <td>2.5</td>
 <td>KCl</td>
 </tr>
 <tr>
 <td>1.25</td>
 <td>NaH2PO4</td>
 </tr>
 <tr>
 <td>0.5</td>
 <td>CaCl2</td>
 </tr>
 </tbody>
</table>
Table 1. Recipe for slice solution.
<table border="1">
 <tbody>
 <tr>
 <td colspan="2">ACSF (in mM)</td>
 </tr>
 <tr>
 <td>125</td>
 <td>NaCl</td>
 </tr>
 <tr>
 <td>26</td>
 <td>NaHCO3</td>
 </tr>
 <tr>
 <td>10</td>
 <td>D-glucose</td>
 </tr>
 <tr>
 <td>3</td>
 <td>KCl</td>
 </tr>
 <tr>
 <td>2.5/1.5</td>
 <td>MgCl2 (recovery/experiment)</td>
 </tr>
 <tr>
 <td>1.6</td>
 <td>CaCl2</td>
 </tr>
 <tr>
 <td>1.25</td>
 <td>NaH2PO4</td>
 </tr>
 </tbody>
</table>
Table 2. Recipe for both recovery (r-ACSF) and experimental (e-ACSF) solution.
2. Preparation of the staining chamber
To load the cells with the calcium-dependent indicator or cell-specific marker, slices need to be transferred to a chamber for the staining procedure. Although commercial chambers may be available, one can easily be assembled from standard lab equipment for very little cost. The key features of such a chamber are that the slices are warmed to between 30 and 35&deg;C, incubated in a continuously oxygenated medium and that the chamber is shielded from light.
<ol>
 <li>Using a heated rod, make a small hole in the side wall of two disposable polystyrene petri dishes (35 and 100mm diameter) that is just big enough to allow a section of silicon tubing (diameter approx 1.5mm) to pass through.</li>
 <li>Thread the silicon tubing through the hole in the small petri dish and form a loop inside the inner wall of the small dish.</li>
 <li>Use superglue to seal the open end of the tubing and stick the rest of the tubing to the inner wall of the petri dish. </li>
 <li>Using a needle, make fine holes at evenly-spaced intervals in the tubing within the inner petri dish.</li>
 <li>Glue the smaller petri dish inside the larger one, with both holes aligned. Thread the other end of the silicon tubing through the hole in the wall of the larger petri dish.</li>
 <li>For the interface dish, take a cell culture insert well with semi-permeable membrane and using a heated scalpel, cut the top 1cm away to leave a shallow dish to hold the slices during incubation.</li>
 <li>On the lid of the larger petri dish, make a hole approx. 0.5-1cm diameter to allow for carbogen (95% O2, 5% CO2) to flow into the chamber. </li>
 <li>Take a 5 or 10ml plastic syringe. Using a heated scalpel, make an angled cut to remove the end of the syringe and keep a few cm length of syringe tube with the tip. Using superglue, attach the cut surface of the syringe tube to the petri dish lid, over the 0.5-1cm diameter hole.</li>
 <li>Attach silicon tubing and a tubing connector to the syringe tip. Attach another tubing connector to the silicon tubing entering the base of the petri dishes.</li>
 <li>Connect up both tubes to a regulator for carbogen (95% O2, 5% CO2) supply. Place the staining chamber on a hot plate for incubation between 30-35&deg;C. Ensure the chamber can be shielded from light during the entire incubation process. The staining chamber is now ready to assemble and use for slice incubation. </li>
</ol>
<img src="/files/ftp_upload/3550/3550methodologyfig1.jpg" alt="Methodology Figure 1" /> 
Methodology Figure 1. Cross-section of the staining chamber showing slice incubation (above) and application of calcium-sensitive indicator (pipetted green dye, below).
3. Staining of slices
Throughout any handling involving fluorescent dyes, avoid photobleaching by working with little light and keep the dye and the stained tissue in the dark between handling.
<ol>
 <li>Put the staining chamber on a heat plate (35&deg;C), connect it to a carbogen gas supply and fill it with approximately 1.5ml r-ACSF from the slice holder.</li>
 <li>Place the interface dish in the staining chamber and fill it with 1ml ACSF.</li>
 <li>Ensure a good carbogen supply in the staining chamber to keep the ACSF bubbling at a gentle, steady rate.</li>
 <li>Add 9 &mu;l DMSO and 1 &mu;l pluronic acid (20% in DMSO stock solution, Invitrogen) into a vial of 50 &mu;g Fura 2-AM. Vortex the dye mix for 15 minutes.</li>
 <li>Transfer the slices into the interface dish and pipette the dye into the r-ACSF directly over the hippocampal and entorhinal cortex region of interest, as indicated in Methodology Figure 1 (above), achieving a final dye concentration of 50mM.</li>
 <li>Close the lid of the staining chamber and incubate the dye for 20 - 40 minutes depending on the age of the animal (see Table 3).</li>
 <li>Transfer the slices back into the slice holder.</li>
</ol>
<table border="1">
 <tbody>
 <tr>
 <td>Age (postnatal days)</td>
 <td>Incubation time (min)</td>
 </tr>
 <tr>
 <td>&lt;p8</td>
 <td>20</td>
 </tr>
 <tr>
 <td>p8-p9</td>
 <td>25</td>
 </tr>
 <tr>
 <td>p10-p11</td>
 <td>30</td>
 </tr>
 <tr>
 <td>p12-13</td>
 <td>35</td>
 </tr>
 <tr>
 <td>&gt;p13</td>
 <td>40</td>
 </tr>
 </tbody>
</table>
Table 3. Incubation times for different ages, empirically-determined in the lab.
4. Other dyes and older tissue
Perhaps due to increased myelination in the brain tissue, slices from older rodents do not take up Fura 2-AM ester that easily. To facilitate uptake of this dye a preincubation step using Cremophor EL (Sigma) is used for brains of mice at P13 and older11. Cremophor is a non-ionic surfactant used as an excipient in many pharmaceutical applications. Without this step, we find that cell-specific labelling is extremely poor and inconsistent throughout the cortical slice.
<ol>
 <li>Transfer old slices into a shallow preincubation dish filled with 3ml r-ACSF and 8 &mu;l 0.5% cremophor (Sigma) heated to 35&deg;C for 3 minutes.</li>
 <li>Transfer the slices into the interface dish and follow the normal staining procedure (Steps 4-7, see above).</li>
</ol>
Calcium dyes load both neurons and non-neurons in the slice preparation. To identify and distinguish between these cell types within the network, sulforhodamine 101 (SR101) can be used to label astrocytes within the slice.
Staining of slices for sulforhodamine 101
Follow steps 1-3 as before (section 3). 
Take 1 &mu;l of 10mM stock solution of sulforhodamine (Sigma) from its storage at -20&deg;C and dissolve in 999 &mu;l r-ACSF to give a 10 &mu;M solution. Pipette the purple dye over the slices as before (steps 5-7), leaving the slices incubating for a period of 15 minutes. 
Microglia and endothelial cells are labelled using FITC tomato lectin dye
Follow steps 1-3 as before. 
Take 25 &mu;l of 2mg/ml Lycopersicon esculentum (tomato) lectin FITC conjugate (L0401, Sigma) stock solution into 2.5ml r-ACSF to give 20 &mu;g/ml concentration. Pipette the dye over the slices as before (steps 5-7), leaving the slices incubating for a period of 45 minutes.
Note, it is not possible to combine this dye with a calcium-sensitive indicator like Fura 2-AM or Oregon Green BAPTA-1 (OGB-1) that fluoresce in the green range of the spectrum since the photons from both dyes will be emitted at overlapping wavelengths. However, there are other calcium-indicator dyes such as Calcium orange, Fura Red that may be used in combination with appropriate filters or Texas-Red lectin conjugates to combine with calcium-indicator dyes in the green spectrum.
5. Attaching slices to recording chamber
During imaging slices need to be stable under the microscope. Usually a metal harp is placed to hold down the tissue but it can unevenly distort the surface of the slice, giving only part of the field of view for imaging in focus. To avoid this, slices are stuck to the recording chamber using Polyethylenimine (PEI).
<ol>
 <li>Prepare the PEI solution (1ml Polyethyleneimine solution in 250ml boric buffer (table 4)). Make sure the PEI dissolves completely (stir overnight).</li>
 <li>Fill the recording chambers with PEI solution so that the bottom is covered with fluid at least one hour before you want to apply the slices</li>
 <li>Wash the recording chambers with distilled water and then with r-ACSF from the holding chamber.</li>
 <li>Transfer one slice into a recording chamber.</li>
 <li>Remove the r-ACSF with a pipette and position the slice in the middle using a brush.</li>
 <li>Remove the r-ACSF around the slice using pieces of filterpaper. For stability and adherence, it is important that there is no ACSF around the slice anymore.</li>
 <li>Pipette approximately 0.5-1ml ACSF, depending on the size of the recording chamber, onto the slice. ACSF should just cover the slice surface.</li>
 <li>Put the recording chamber with the slice into a large humidified interface container, perfused with carbogen, and let the slices recover for at least one hour.</li>
</ol>
<table border="1">
<tbody>
 <tr>
 <td colspan="2">PEI solution</td>
 </tr>
 <tr>
 <td>1ml</td>
 <td>Poly(ethyleneimine) solution</td>
 </tr>
 <tr>
 <td>in 250ml boric buffer</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>40mM</td>
 <td>boric acid</td>
 </tr>
 <tr>
 <td>10mM</td>
 <td>sodium tetraborate decahydrate</td>
 </tr>
</tbody>
</table>
Table 4. Recipe PEI solution.
6. Imaging
Calcium-dependent indicator dyes can be imaged using either one- or two-photon microscopy. Use of two-photon imaging only activates indicator dye within the focal volume of the region of interest, thus reducing the amount of light scatter in the tissue. Furthermore, it enables better depth penetration of the light into the slice.
For functional calcium imaging we use a Titanium sapphire laser supplied by Coherent coupled to an Olympus microscope with a 20x objective (NA 0.95) and a Trimscope system by LaVision Biotec. The Trimscope system enables frame scanning with 64 beamlets simultaneously and is coupled with a Hamamatsu C9100 EM-CCD camera for fast frame-scanning rates.
<ol>
 <li>Position the recording chamber under the microscope and establish a stable perfusion with imaging e-ACSF (1.6 Ca2+/1.5 Mg2+ ratio (table 2)) heated to a more physiological temperature of minimal 30&deg;C. </li>
 <li>Focus into the region of interest (ROI) using white light illumination. Avoid exposing the slice to light longer than necessary.</li>
 <li>Choose the wavelength, imaging field of view, the scanning frequency, pixel density and additional software options applicable for the tissue and biological signals you wish to measure. For calcium network imaging with Fura 2-AM we typically choose a wavelength of 820nm, a 250x250 &mu;m imaging field of view, a linescanning frequency of 1200Hz and 2-by-2 binning. This results in a frametime of about 100ms and thus an imaging frequency of about 10Hz. </li>
 <li>Check if your ROI is in focus using a continuous scanning mode and selected wavelength of laserlight. Adjust focus and laser intensity to avoid pixel saturation and bleaching of your image.</li>
 <li>Make a timelapse movie of your ROI. We typically acquire two timelapse movies of 2000 frames each.</li>
 <li>For cell detection and identification, take a z-stack using a stepsize of 1 &mu;m and image +/ 20 &mu;m around your imaged plane of focus.</li>
</ol>
7. Representative results:
Successful loading of calcium indicators, Fura 2-AM are shown in Figure 1 in developing neocortical and entorhinal cortex networks using multiphoton imaging. Some dye is still present as background staining in the tissue but cell soma and in some cases, proximal dendrites are clearly visible and separate from surrounding neuropil. If loading has not been successful, very little cell-specific staining is observed and small clusters of dye spots are often visible on the slice surface in dead membrane debris.
In these screenshots, the network of cells is clearly visible in a single plane of focus for simultaneous cell imaging. Use of a metal harp or incomplete sticking of the slice to the recording chamber can result in an uneven slice surface to be imaged.
<img src="/files/ftp_upload/3550/3550fig1.jpg" alt="Figure 1" /> 
Figure 1. Fura 2-AM ester-loaded developing neocortical (A, left) and entorhinal (B, right) networks. Scale bars 100 &mu;m. 
To separate neurons from astrocytes, co-application of sulforhodamine 101 with Fura 2-AM ester enables separation of cell types within the network.
<img src="/files/ftp_upload/3550/3550fig2.jpg" alt="Figure 2" /> 
Figure 2. Astrocyte labeling with sulforhodamine 101. A Co-labelling of Fura 2-AM ester and sulforhodamine 101. Excitation wavelength: 820nm. Image collection on PMTs with dichroic mirror at 560/70nm for wavelength separation. B Representative fluorescence traces from a neuron (above) and an astrocyte (below). Scale bars 60 sec, &Delta;F 10 (au fluo).
<img src="/files/ftp_upload/3550/3550fig3.jpg" alt="Figure 3" /> 
Figure 3. FITC-conjugated Lycopersicon esculentum (tomato) lectin staining. Labelling of microglia and endothelial cells in developing mouse hippocampus (A) and superficial entorhinal cortex (B).
Calcium indicator dyes are used to read-out activity from multiple cells simultaneously in developing cortical and hippocampal networks. 
Figure 4. Dynamic calcium transients in hippocampal and cortical networks. 
Movie 1: Fura 2-AM ester loading of mouse entorhinal cortex during second postnatal week. 
<a href="https://www.jove.com/files/ftp_upload/3550/3550movie1.avi">Click here to view movie 1.</a>
Movie 2: Fura 2-AM ester loading of mouse hippocampus during the first postnatal week. 
<a href="https://www.jove.com/files/ftp_upload/3550/3550movie2.avi">Click here to view movie 2.</a>
Movie 3: Fluo-4 loading of mouse cortex, during the first postnatal week. 
<a href="https://www.jove.com/files/ftp_upload/3550/3550movie3.avi">Click here to watch movie 3.</a>
In the case of Fura 2-AM ester, cell activation involving a depolarization-induced influx of calcium, decreases dye fluorescence. For dyes such as Fluo-4, the opposite is true and cell depolarization is observed as an increase in photon emissions. Somatic calcium transients are mainly measured but activity in larger proximal dendrites can also be seen in some preparations, as shown in Movie 2.
Read-out of network activity can be quantified using commercial or in-house software scripts. In our lab, cell identification and network activity is measured and analyzed in a semi-automated manner using in-house code for Matlab (Mathworks). 
<img src="/files/ftp_upload/3550/3550fig5.jpg" alt="Figure" /> 
Figure 5. Representative analysis of synchronized calcium transients from a developing cortical network. 3D representation of one neuron (A) used to automatically create a neuron mask (B) from a z-stack for automated neuron detection. Measurements of calcium transients include amplitude, frequency and # of active cells that can be read out from single neuron data (C). Synchrony between different traces can be visualized using a raster plot (D).

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LaVision Biotec GmBH (Bielefeld, Germany) sponsored the submission fees of this manuscript article.

The methods we demonstrate here show suitable protocols for calcium imaging of network dynamics from identifiable cells within developing cortical and hippocampal networks in the mouse and also in the rat brain. These methods provide optimal spatial resolution to visualize a local network of cell somas simultaneously for measuring suprathreshold activity. Temporal resolution of network activity can be varied, depending on frame acquisition CCD camera settings, to optimize signal:noise measurements for long duration suprathreshold events, typically found in the developing nervous system. These protocols are not restricted to hippocampal and cortical networks but also label cells throughout the developing nervous system. The limitation of this method is that only suprathreshold activity can be recorded.

<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Solutions</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>Sigma Aldrich</td>
			<td>22,350-6</td>
			<td></td>
		</tr>
		<tr>
			<td>Choline chloride</td>
			<td>Sigma</td>
			<td>C7527</td>
			<td></td>
		</tr>
		<tr>
			<td>cremaphor</td>
			<td>Fluka</td>
			<td>27963</td>
			<td>Cremophor EL&#174;</td>
		</tr>
		<tr>
			<td>DMSO (Dimethyl sulfoxide)</td>
			<td>Sigma Aldrich</td>
			<td>472301</td>
			<td></td>
		</tr>
		<tr>
			<td>D-glucose</td>
			<td>Sigma Aldrich</td>
			<td>16325</td>
			<td></td>
		</tr>
		<tr>
			<td>Fura-2 AM</td>
			<td>Invitrogen</td>
			<td>F-1221</td>
			<td>special packaging</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma Aldrich</td>
			<td>60130</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>Fluka</td>
			<td>63072</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma Aldrich</td>
			<td>31434</td>
			<td></td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>Sigma Aldrich</td>
			<td>31437</td>
			<td></td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>Merck</td>
			<td>106346</td>
			<td></td>
		</tr>
		<tr>
			<td>Pluronic&#174; F-127</td>
			<td>Invitrogen</td>
			<td>P-3000MP</td>
			<td>20% solution in DMSO</td>
		</tr>
		<tr>
			<td>sodium ascorbate</td>
			<td>Sigma</td>
			<td>A7631</td>
			<td></td>
		</tr>
		<tr>
			<td>sodium pyruvate</td>
			<td>Sigma Aldrich</td>
			<td>P2256</td>
			<td></td>
		</tr>
		<tr>
			<td>Lectin from Lycopersicon esculentum (tomato)</td>
			<td>Sigma</td>
			<td>L0401</td>
			<td></td>
		</tr>
		<tr>
			<td>Sulforhodamine 101 acid chloride</td>
			<td>Sigma</td>
			<td>S3388</td>
			<td></td>
		</tr>
		<tr>
			<td>Poly(ethyleneimine) solution</td>
			<td>Fluka</td>
			<td>P3143</td>
			<td></td>
		</tr>
		<tr>
			<td>boric acid</td>
			<td>Sigma</td>
			<td>B6768</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium tetraborate decahydrate</td>
			<td>Sigma</td>
			<td>B3545</td>
			<td></td>
		</tr>
		<tr>
			<td>PEI</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Staining chamber</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe (5 or 10 ml)</td>
			<td>Terumo</td>
			<td></td>
			<td>with luer lock</td>
		</tr>
		<tr>
			<td>Cell Culture Inserts for 6-well plates, 8.0 &#181;m, transparent PET membrane&#160;</td>
			<td>BD Falcon&#8482;&#160;</td>
			<td>353093</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri dish (35 and 100mm)</td>
			<td></td>
			<td></td>
			<td>disposable, polystyrene</td>
		</tr>
		<tr>
			<td>tubing and connectors</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>super glue</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Holders and Chambers</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Slice holder</td>
			<td></td>
			<td></td>
			<td>submerged</td>
		</tr>
		<tr>
			<td>Staining chamber</td>
			<td></td>
			<td></td>
			<td>interface chamber</td>
		</tr>
		<tr>
			<td>recording chamber</td>
			<td></td>
			<td></td>
			<td>e.g. MEA probes</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Imaging setup</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Titanium Sapphire laser (Chameleon)</td>
			<td>Coherent</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Trim Scope</td>
			<td>LaVision</td>
			<td></td>
			<td>64 beams</td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Leica</td>
			<td></td>
			<td>High NA lense</td>
		</tr>
	</tbody>
</table>

functional calcium imaging in developing cortical networks

A hallmark pattern of activity in developing nervous systems is spontaneous, synchronized network activity. Synchronized activity has been observed in intact spinal cord, brainstem, retina, cortex and dissociated neuronal culture preparations. During periods of spontaneous activity, neurons depolarize to fire single or bursts of action potentials, activating many ion channels. Depolarization activates voltage-gated calcium channels on dendrites and spines that mediate calcium influx. Highly synchronized electrical activity has been measured from local neuronal networks using field electrodes. This technique enables high temporal sampling rates but lower spatial resolution due to integrated read-out of multiple neurons at one electrode. Single cell resolution of neuronal activity is possible using patch-clamp electrophysiology on single neurons to measure firing activity. However, the ability to measure from a network is limited to the number of neurons patched simultaneously, and typically is only one or two neurons. The use of calcium-dependent fluorescent indicator dyes has enabled the measurement of synchronized activity across a network of cells. This technique gives both high spatial resolution and sufficient temporal sampling to record spontaneous activity of the developing network.
A key feature of newly-forming cortical and hippocampal networks during pre- and early postnatal development is spontaneous, synchronized neuronal activity (Katz &amp; Shatz, 1996; Khaziphov &amp; Luhmann, 2006). This correlated network activity is believed to be essential for the generation of functional circuits in the developing nervous system (Spitzer, 2006). In both primate and rodent brain, early electrical and calcium network waves are observed pre- and postnatally in vivo and in vitro (Adelsberger et al., 2005; Garaschuk et al., 2000; Lamblin et al., 1999). These early activity patterns, which are known to control several developmental processes including neuronal differentiation, synaptogenesis and plasticity (Rakic &amp; Komuro, 1995; Spitzer et al., 2004) are of critical importance for the correct development and maturation of the cortical circuitry.
In this JoVE video, we demonstrate the methods used to image spontaneous activity in developing cortical networks. Calcium-sensitive indicators, such as Fura 2-AM ester diffuse across the cell membrane where intracellular esterase activity cleaves the AM esters to leave the cell-impermeant form of indicator dye. The impermeant form of indicator has carboxylic acid groups which are able to then detect and bind calcium ions intracellularly.. The fluorescence of the calcium-sensitive dye is transiently altered upon binding to calcium. Single or multi-photon imaging techniques are used to measure the change in photons being emitted from the dye, and thus indicate an alteration in intracellular calcium. Furthermore, these calcium-dependent indicators can be combined with other fluorescent markers to investigate cell types within the active network.

Watch this Scientific Journal Video about Functional Calcium Imaging in Developing Cortical Networks at JoVE.com

Functional Calcium Imaging in Developing Cortical Networks

Spontaneous activity of developing neuronal networks can be measured using AM-ester forms of calcium-sensitive indicator dyes. Changes in intracellular calcium, indicating neuronal activation, are detected as transient changes in indicator fluorescence with one- or two-photon imaging. This protocol can be adapted for a range of developmentally-dependent neuronal networks in vitro.

A hallmark pattern of activity in developing nervous systems is spontaneous, synchronized network activity. Synchronized activity has been observed in ...

functional-calcium-imaging-in-developing-cortical-networks

Research

JoVE Journal

Neuroscience

38.8K 조회수.  VU University, Amsterdam. 의 연결을 네트워크를 개발 자발적인 활동은 칼슘에 민감한 지표 염료의 AM - 에스테르 양식을 사용하여 측정할 수 있습니다. 의 연결을 활성화를 나타내는 세포 내 칼슘의 변화는 하나 또는 두 개의 광자 이미징과 형광 표시기에 일시적 변화로 감지됩니다. 이 프로토콜은 발달 종속의 연결 네트워크의 범위에 대해 적용할 수 있습니다 체외에서.

비디오: Functional Calcium Imaging in Developing Cortical Networks

Functional Mapping with Simultaneous MEG and EEG

We use magnetoencephalography (MEG) and electroencephalography (EEG) to locate and determine the temporal evolution in brain areas involved in the processing of simple sensory stimuli. We will use somatosensory stimuli to locate the hand somatosensory areas, auditory stimuli to locate the auditory cortices, visual stimuli in four quadrants of the visual field to locate the early visual areas. These type of experiments are used for functional mapping in epileptic and brain tumor patients to locate eloquent cortices. In basic neuroscience similar experimental protocols are used to study the orchestration of cortical activity. The acquisition protocol includes quality assurance procedures, subject preparation for the combined MEG/EEG study, and acquisition of evoked-response data with somatosensory, auditory, and visual stimuli. We also demonstrate analysis of the data using the equivalent current dipole model and cortically-constrained minimum-norm estimates. Anatomical MRI data are employed in the analysis for visualization and for deriving boundaries of tissue boundaries for forward modeling and cortical location and orientation constraints for the minimum-norm estimates.

We use magnetoencephalography (MEG) and electroencephalography (EEG) to map brain areas involved in the processing of simple sensory stimuli.

We use magnetoencephalography (MEG) and electroencephalography (EEG) to locate and determine the temporal evolution in brain areas involved in the ...

Time-lapse Live Imaging of Clonally Related Neural Progenitor Cells in the Developing Zebrafish Forebrain

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To understand the behavior of neural progenitor cells in the complex in vivo environment, time-lapse live imaging of neural progenitor cells in an intact brain is critically required. In this video, we exploit the unique features of zebrafish embryos to visualize the development of forebrain neural progenitor cells in vivo. We use electroporation to genetically and sparsely label individual neural progenitor cells. Briefly, DNA constructs coding for fluorescent markers were injected into the forebrain ventricle of 22 hours post fertilization (hpf) zebrafish embryos and electric pulses were delivered immediately. Six hours later, the electroporated zebrafish embryos were mounted with low melting point agarose in glass bottom culture dishes. Fluorescently labeled neural progenitor cells were then imaged for 36hours with fixed intervals under a confocal microscope using water dipping objective lens. The present method provides a way to gain insights into the in vivo development of forebrain neural progenitor cells and can be applied to other parts of the central nervous system of the zebrafish embryo.

The present video demonstrates a method which takes advantage of the combination of electroporation and confocal microscopy to perform live imaging on individual neural progenitor cells in the developing zebrafish forebrain. In vivo analysis of the development of forebrain neural progenitor cells at a clonal level can be achieved in this way.

Precise patterns of division, migration and differentiation of neural progenitor cells are crucial for proper brain development and function1,2. To ...

In vivo Neuronal Calcium Imaging in C. elegans

The nematode worm C. elegans is an ideal model organism for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, well-characterized nervous system allows identification and fluorescence imaging of any neuron within the intact animal. Simple immobilization techniques with minimal impact on the animal's physiology allow extended time-lapse imaging. The development of genetically-encoded calcium sensitive fluorophores such as cameleon 1 and GCaMP 2 allow in vivo imaging of neuronal calcium relating both cell physiology and neuronal activity. Numerous transgenic strains expressing these fluorophores in specific neurons are readily available or can be constructed using well-established techniques. Here, we describe detailed procedures for measuring calcium dynamics within a single neuron in vivo using both GCaMP and cameleon. We discuss advantages and disadvantages of both as well as various methods of sample preparation (animal immobilization) and image analysis. Finally, we present results from two experiments: 1) Using GCaMP to measure the sensory response of a specific neuron to an external electrical field and 2) Using cameleon to measure the physiological calcium response of a neuron to traumatic laser damage. Calcium imaging techniques such as these are used extensively in C. elegans and have been extended to measurements in freely moving animals, multiple neurons simultaneously and comparison across genetic backgrounds. C. elegans presents a robust and flexible system for in vivo neuronal imaging with advantages over other model systems in technical simplicity and cost.

In vivo Neuronal Calcium Imaging in C. elegans

With its small transparent body, well-documented neuroanatomy and a host of amenable genetic techniques and reagents, C. elegans makes an ideal model organism for in vivo neuronal imaging using relatively simple, low-cost techniques. Here we describe single neuron imaging within intact adult animals using genetically encoded fluorescent calcium indicators.

The nematode worm C. elegans is an ideal model organism  for relatively simple, low cost neuronal imaging in vivo. Its small transparent body and simple, ...

Fast Micro-iontophoresis of Glutamate and GABA: A Useful Tool to Investigate Synaptic Integration

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of the dendritic tree, which eventually leads to neuronal output of action potentials at the axon. The influence of diverse spatial and temporal parameters of specific synaptic input on neuronal output is currently under investigation, e.g. the distance-dependent attenuation of dendritic inputs, the location-dependent interaction of spatially segregated inputs, the influence of GABAergig inhibition on excitatory integration, linear and non-linear integration modes, and many more.
With fast micro-iontophoresis of glutamate and GABA it is possible to precisely investigate the spatial and temporal integration of glutamatergic excitation and GABAergic inhibition. Critical technical requirements are either a triggered fluorescent lamp, light-emitting diode (LED), or a two-photon scanning microscope to visualize dendritic branches without introducing significant photo-damage of the tissue. Furthermore, it is very important to have a micro-iontophoresis amplifier that allows for fast capacitance compensation of high resistance pipettes. Another crucial point is that no transmitter is involuntarily released by the pipette during the experiment.
Once established, this technique will give reliable and reproducible signals with a high neurotransmitter and location specificity. Compared to glutamate and GABA uncaging, fast iontophoresis allows using both transmitters at the same time but at very distant locations without limitation to the field of view. There are also advantages compared to focal electrical stimulation of axons: with micro-iontophoresis the location of the input site is definitely known and it is sure that only the neurotransmitter of interest is released. However it has to be considered that with micro-iontophoresis only the postsynapse is activated and presynaptic aspects of neurotransmitter release are not resolved. In this article we demonstrate how to set up micro-iontophoresis in brain slice experiments.

In this article we introduce fast micro-iontophoresis of neurotransmitters as a technique to investigate integration of postsynaptic signals with high spatial and temporal precision.

One of the fundamental interests in neuroscience is to understand the integration of excitatory and inhibitory inputs along the very complex structure of ...

In Vivo Functional Brain Imaging Approach Based on Bioluminescent Calcium Indicator GFP-aequorin

Functional in vivo imaging has become a powerful approach to study the function and physiology of brain cells and structures of interest. Recently a new method of Ca2+-imaging using the bioluminescent reporter GFP-aequorin (GA) has been developed. This new technique relies on the fusion of the GFP and aequorin genes, producing a molecule capable of binding calcium and&#160;&#8212; with the addition of its cofactor coelenterazine &#8212; emitting bright light that can be monitored through a photon collector. Transgenic lines carrying the GFP-aequorin gene have been generated for both mice and Drosophila. In Drosophila, the GFP-aequorin gene has been placed under the control of the GAL4/UAS binary expression system allowing for targeted expression and imaging within the brain. This method has subsequently been shown to be capable of detecting both inward Ca2+-transients and Ca2+-released from inner stores. Most importantly it allows for a greater duration in continuous recording, imaging at greater depths within the brain, and recording at high temporal resolutions (up to 8.3 msec). Here we present the basic method for using bioluminescent imaging to record and analyze Ca2+-activity within the mushroom bodies, a structure central to learning and memory in the fly brain.

In Vivo Functional Brain Imaging Approach Based on Bioluminescent Calcium Indicator GFP-aequorin

Here we present a novel Ca2+-imaging approach using a bioluminescent reporter. This approach uses a fused construct GFP-aequorin which binds to Ca2+ and emits light, eliminating the need for light excitation. Significantly this method permits long continuous imaging, access to deep brain structures and high temporal resolution.

Functional in vivo imaging has become a powerful approach to study the function and physiology of brain cells and structures of interest. Recently a new ...

Simultaneous Recordings of Cortical Local Field Potentials, Electrocardiogram, Electromyogram, and Breathing Rhythm from a Freely Moving Rat

Monitoring the physiological dynamics of the brain and peripheral tissues is necessary for addressing a number of questions about how the brain controls body functions and internal organ rhythms when animals are exposed to emotional challenges and changes in their living environments. In general experiments, signals from different organs, such as the brain and the heart, are recorded by independent recording systems that require multiple recording devices and different procedures for processing the data files. This study describes a new method that can simultaneously monitor electrical biosignals, including tens of local field potentials in multiple brain regions, electrocardiograms that represent the cardiac rhythm, electromyograms that represent awake/sleep-related muscle contraction, and breathing signals, in a freely moving rat. The recording configuration of this method is based on a conventional micro-drive array for cortical local field potential recordings in which tens of electrodes are accommodated, and the signals obtained from these electrodes are integrated into a single electrical board mounted on the animal's head. Here, this recording system was improved so that signals from the peripheral organs are also transferred to an electrical interface board. In a single surgery, electrodes are first separately implanted into the appropriate body parts and the target brain areas. The open ends of all of these electrodes are then soldered to individual channels of the electrical board above the animal's head so that all of the signals can be integrated into the single electrical board. Connecting this board to a recording device allows for the collection of all of the signals into a single device, which reduces experimental costs and simplifies data processing, because all data can be handled in the same data file. This technique will aid the understanding of the neurophysiological correlates of the associations between central and peripheral organs.

This study introduces a method for the simultaneous recording of local field potentials in the brain, electrocardiograms, electromyograms, and breathing signals of a freely moving rat. This technique, which reduces experimental costs and simplifies data analysis, will contribute to the understanding of the interactions between the brain and peripheral organs.

Monitoring the physiological dynamics of the brain and peripheral tissues is necessary for addressing a number of questions about how the brain controls ...

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 ...

In vivo Calcium Imaging in Mouse Inferior Olive

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the cerebellum. IO has long been proposed to be crucial for motor control and its activity is currently considered to be at the center of many hypotheses of both motor and cognitive functions of the cerebellum. While its physiology and function have been relatively well studied on single-cell level in vitro, presently there are no reports on the organization of the IO network activity in living animals. This is largely due to the extremely challenging anatomical location of the IO, making it difficult to subject to conventional fluorescent imaging methods, where an optic path must be created through the entire brain located dorsally to the region of interest.
Here we describe an alternative method for obtaining state-of-the-art -level calcium imaging data from the IO network. The method takes advantage of the extreme ventral location of the IO and involves a surgical procedure for inserting a gradient-refractive index (GRIN) lens through the neck viscera to come into contact with the ventral surface of the calcium sensor GCaMP6s-expressing IO in anesthetized mice. A representative calcium imaging recording is shown to demonstrate the feasibility to record IO neuron activity after the surgery. While this is a non-survival surgery and the recordings must be conducted under anesthesia, it avoids damage to life-critical brainstem nuclei and allows conducting large variety of experiments investigating spatiotemporal activity patterns and input integration in the IO. This procedure with modifications could be used for recordings in other, adjacent regions of the ventral brainstem.

In vivo Calcium Imaging in Mouse Inferior Olive

We present a protocol to expose the brainstem of adult mouse from the ventral side. By using a gradient-refractive index lens with a miniature microscope, calcium imaging can be used to examine the activity of inferior olive neural somata in vivo.

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the ...

Reliable Acquisition of Electroencephalography Data during Simultaneous Electroencephalography and Functional MRI

Simultaneous electroencephalography (EEG) and functional magnetic resonance imaging (fMRI), EEG-fMRI, combines the complementary properties of scalp EEG (good temporal resolution) and fMRI (good spatial resolution) to measure neuronal activity during an electrographic event, through hemodynamic responses known as blood-oxygen-level-dependent (BOLD) changes. It is a non-invasive research tool that is utilized in neuroscience research and is highly beneficial to the clinical community, especially for the management of neurological diseases, provided that proper equipment and protocols are administered during data acquisition. Although recording EEG-fMRI is apparently straightforward, the correct preparation, especially in placing and securing the electrodes, is not only important for safety but is also critical in ensuring the reliability and analyzability of the EEG data obtained. This is also the most experience-demanding part of the preparation. To address these issues, a straightforward protocol that ensures data quality was developed. This article provides a step-by-step guide for acquiring reliable EEG data during EEG-fMRI using this protocol that utilizes readily available medical products. The presented protocol can be adapted to different applications of EEG-fMRI in research and clinical settings, and may be beneficial to both inexperienced and expert operators.

This article provides a straightforward protocol for acquiring good quality electroencephalography (EEG) data during simultaneous EEG and functional magnetic resonance imaging by utilizing readily available medical products.

Simultaneous electroencephalography (EEG) and functional magnetic resonance imaging (fMRI), EEG-fMRI, combines the complementary properties of scalp EEG ...

In Vivo Calcium Imaging of Neuronal Ensembles in Networks of Primary Sensory Neurons in Intact Dorsal Root Ganglia

Ca2+ imaging can be used as a proxy for cellular activity, including action potentials and various signaling mechanisms involving Ca2+ entry into the cytoplasm or the release of intracellular Ca2+ stores. Pirt-GCaMP3-based Ca2+ imaging of primary sensory neurons of the dorsal root ganglion (DRG) in mice offers the advantage of simultaneous measurement of a large number of cells. Up to 1,800 neurons can be monitored, allowing neuronal networks and somatosensory processes to be studied as an ensemble in their normal physiological context at a populational level in vivo. The large number of neurons monitored allows the detection of activity patterns that would be challenging to detect using other methods. Stimuli can be applied to the mouse hindpaw, allowing the direct effects of stimuli on the DRG neuron ensemble to be studied. The number of neurons producing Ca2+ transients as well as the amplitude of Ca2+ transients indicates sensitivity to specific sensory modalities. The diameter of neurons provides evidence of activated fiber types (non-noxious mechano vs. noxious pain fibers, A&#946;, A&#948;, and C fibers). Neurons expressing specific receptors can be genetically labeled with td-Tomato and specific Cre recombinases together with Pirt-GCaMP. Therefore, Pirt-GCaMP3 Ca2+ imaging of DRG provides a powerful tool and model for the analysis of specific sensory modalities and neuron subtypes acting as an ensemble at the populational level to study pain, itch, touch, and other somatosensory signals.

In Vivo Calcium Imaging of Neuronal Ensembles in Networks of Primary Sensory Neurons in Intact Dorsal Root Ganglia

This protocol describes the surgical exposure of the dorsal root ganglion (DRG) followed by GCaMP3 (genetically-encoded Ca2+ indicator; Green Fluorescent Protein-Calmodulin-M13 Protein 3) Ca2+ imaging of the neuronal ensembles using Pirt-GCaMP3 mice while applying a variety of stimuli to the ipsilateral hind paw.

Ca2+ imaging can be used as a proxy for cellular activity, including action potentials and various signaling mechanisms involving Ca2+ entry into the ...