This work was supported by CNPq (Conselho Nacional de Desenvolvimento Cient&#237;fico e Tecnol&#243;gico), CAPES (Coordena&#231;&#227;o de Aperfei&#231;oamento de Pessoal de N&#237;vel Superior) and FAPERN (Funda&#231;&#227;o de Amparo a Pesquisa do Rio Grande do Norte).

All experiments involving live animals described in this protocol are conducted according to the National and International laws and were approved by the local University Animal Care and Use Committee (CEUA/UFRN), under the license 009/2014. The following protocol is performed in a sterile environment. Familiarity with basic cell culture is expected.
1. Dorsolateral Telencephalon Microdissection
<ol>
	<li>Prepare the dissection medium (100 mL): 98.5 mL Hank&#39;s Balanced Salt Solution (HBSS), 0.5 mL 5 M HEPES pH 7.4, 1 mL penicillin/streptomycin (10,000 units/mL and 10,000 &#181;g/mL).</li>
	<li>Filter-sterilize the dissection medium.</li>
	<li>Remove embryonic day 14 (E14) embryos from a pregnant mouse C57/Bl6 (Mus musculus) under anesthesia with isoflurane. 
	NOTE: Consider E0 as the day of vaginal plug detection. Dissection should be completed no later than 1 h after removal from the pregnant mouse. This protocol also applies for embryos E11-E17.</li>
	<li>Remove the embryos by hysterectomy under sterile conditions.</li>
	<li>Transfer the embryos to a Petri dish with cold dissection medium (4 &#176;C).</li>
	<li>Remove the brains from the skull by cutting along the longitudinal fissure. The number of embryos usually varies from five to ten. Each E14 brain yields approximately 2 x 106 cells, including progenitors and postmitotic neurons (1:1 ratio) 
	NOTE: Use the stereomicroscope for the following steps.</li>
	<li>Remove the meninges using dissection forceps. Split the telencephalon in the middle to separate the hemispheres. To isolate the dorsolateral telencephalon, cut along the dorsomedial curve and pallial-subpallial boundary using dissection forceps. Transfer the dorsolateral telencephalon to a 2 mL tube with cold dissection medium (4 &#176;C) until all brains are dissected.</li>
</ol>
2. Cell Dissociation and Plating
<ol>
	<li>Prepare the proliferation medium (50 mL of DMEM+10% FCS): 44 mL Dulbecco Modified Eagle&#39;s Medium (DMEM), 5mL Fetal Calf Serum (FCS), 0.5 mL 40% glucose, 0.5 mL penicillin/streptomycin (10,000 units/mL and 10,000 &#181;g/mL). Filter-sterilize the proliferation medium.</li>
	<li>Prepare differentiation medium (50 mL of DMEM+2% B27): 48 mL DMEM, 1 mL B27, 0.5 mL 40% glucose, 0.5 mL penicillin/streptomycin (10,000 units/mL and 10,000 &#181;g/mL). Filter-sterilize the differentiation medium.</li>
	<li>After micro-dissection of the dorsolateral telencephalon from E14 mice, centrifuge the tube with the collected tissue and cold dissection medium (for 5 min at 4 &#176;C, 340 x g) to precipitate the tissue.</li>
	<li>Remove the supernatant with a pipette and add 1 mL of pre-warmed (37 &#176;C) Trypsin-EDTA (0.05%) for chemical digestion. Incubate for 15 min at 37 &#176;C. Add 2 mL of proliferation medium to stop trypsin activity.</li>
	<li>Polish the tip of a glass Pasteur pipette over a gas burner or Bunsen burner for few seconds to slightly narrow the aperture. Wash the pipette by aspirating FCS to coat the tip. Avoiding bubbles, dissociate mechanically the cells with a fire-polished and FCS-coated Pasteur pipette.</li>
	<li>Centrifuge the cells (for 5 min at 4 &#176;C, 340 x g). Remove supernatant with a pipette. Add 1 mL of proliferation medium and resuspend the cells using a pipette. Repeat this step once.</li>
	<li>Prepare a 1:1 dilution of the cell suspension using a 0.4% Trypan Blue solution. Non-viable cells will be blue. Count the number of viable cells (unstained) using a Neubauer chamber.</li>
	<li>Dilute the cells in proliferation medium to get 106 cells/mL. Add 500 &#181;L of the cell suspension to each well of a 24 well tissue culture plate (approximately 2.5 &#215; 105&#160;cells/cm2). Incubate cells at 37 &#176;C and 5% CO2. 
	NOTE: Do not use glass coverslips, as they can move during the experiment and change the field of observation. Alternatively, use glass bottom tissue culture plates. 
	NOTE: If you are not using cell-culture treated multidished, it is important to pre-treat your plates with Poly-D-Lysin (50 &#181;g/ml) 2 h at 37 &#176;C. PDL-treated plates can be stored at 4 &#176;C after washing with distilled water.</li>
</ol>
3. Retroviral-mediated Transfection
NOTE: This is a simple method to transfect only progenitor cells. However, other viral vectors or chemical/electrical transfection can be used to insert genes of interest in cultured cells.
<ol>
	<li>To manipulate gene expression and label cells with fluorescent reporter proteins, add retroviruses carrying plasmids with genes of interest 2 h after cell plating. 
	NOTE: Retroviral vectors only integrate into the genome of dividing cells. Therefore, it is important to infect cells when proliferation is high. 
	NOTE: We used a plasmid containing the Beta-actin/CMV fusion sequence (CAG) and an internal ribosomal entry site (IRES) followed by the coding sequence for the green fluorescent protein (GFP), designated pCAG-IRES-GFP44. Genes of interest could be cloned between the CAG promoter and IRES sequence37.</li>
	<li>Dilute the retrovirus in differentiation medium to obtain the desired number of particles per mL. Add 500 &#181;L of differentiation medium with retrovirus to each well. 
	NOTE: The amount of retroviruses used is dependent on its titer and should be calculated to obtain 50 to 100 infected cells per well. This titer allows the identification of isolated GFP-expressing clones, whose relationship can be further confirmed by tracking phase contrast images (see section 6). The titration of retrovirus should be performed using the same cell population45.</li>
</ol>
4. Time-lapse Video-microscopy
NOTE: This step requires an inverted fluorescence microscope with incubation chamber (see Table of Materials).
<ol>
	<li>After the retrovirus infection, place the tissue culture plate into an incubator connected to temperature and CO2 controllers. 
	NOTE: Preheating the chamber for 3 h prior to starting imaging could help minimize drift and defocus during the acquisition. 
	NOTE: The chamber should maintain the cells at constant conditions of 37 &#176;C and 5% CO2.</li>
	<li>Select a position in the XY axis to serve as a zero-point (XY = 0). This will allow a reference to find the positions again in the future, for instance after post-imaging immunocytochemistry (see section 5). 
	NOTE: It is advisable to draw a sign in the plate that can be used as a reference to determine the zero-point.</li>
	<li>Select the positions to be imaged at 10X magnification. 
	NOTE: Select 10-15 positions per well to increase the likelihood to observe retrovirally-transfected cells. 
	NOTE: Long-distance high magnification objectives (20X or 40X) can be used to gain higher resolution images. Depending on the working distance of objectives, glass-bottom plates may be required.</li>
	<li>Acquire phase contrast images every 5 min and fluorescence images every 3 h to decrease phototoxicity. Check and adjust the focus in the first 3 h of experiment, as it may change while the temperature equilibrates. Check and adjust the focus daily. 
	NOTE: We observe that the focus becomes stable after the first 24 h. However, it is advisable to monitor the focus.&#160;Alternatively, use a software-based autofocus system.</li>
	<li>After 7 days, stop acquisition and proceed to post-imaging immunocytochemistry (see section 5).</li>
	<li>After immunocytochemistry, replace the plate in the microscope incubator, find the zero-point, reset XYZ = 0 and reload the experiment with saved positions. Acquire the fluorescence images of each position using appropriate fluorescence filters.</li>
</ol>
5. Post-imaging Immunocytochemistry
<ol>
	<li>After 7 days in culture, fix the cell culture with 4% Paraformaldehyde (PFA) fixative for 15 min at room temperature. 
	CAUTION: PFA is toxic and should be handle in a chemical fume hood.</li>
	<li>Rinse with 0.5 mL PBS 0.1 M for 5 min. Repeat 3 times.</li>
	<li>Incubate in primary antibodies overnight at 4 &#176;C in 0.5% triton and 10% of normal goat serum in PBS 0.1 M. 
	NOTE: Use anti-Microtubule Associated Protein 2 (MAP2) antibody to confirm the neuronal phenotype (mouse IgG1, 1:1,000) and anti-GFP to label the transduced cells (chicken antibody, 1:500). Other antibodies can be used to detect progenitor or glial cells.</li>
	<li>Rinse with 0.5 mL PBS 0.1 M for 5 min. Repeat 3 times.</li>
	<li>Dilute secondary antibodies in 0.5% triton and 10% of normal goat serum in PBS 0.1 M and incubate for 2 h at room temperature. Recommended dilution: 1:1,000.</li>
	<li>Rinse with 0.5 mL PBS 0.1 M for 5 min. Repeat 3 times.</li>
	<li>For nuclei staining, incubate the cells for 5 min with 0.1 &#181;g/mL 4&#8242;6&#8242;-diamino-2-phenylindone (DAPI) in PBS 0.1 M.</li>
	<li>Rinse with 0.5 mL PBS 0.1 M for 5 min. Repeat 3 times.</li>
	<li>Keep the cells in PBS 0.1 M and protect from light (e.g., wrap the plate with foil).</li>
</ol>
6. Cell Tracking
NOTE: Here, we briefly describe the main steps to analyze time-lapse video-microscopy data using the software The Tracking Tool (tTt), which is available at the link provided in Table of Materials.
NOTE: If the image acquisition is performed with other software, import the image data with tTt Converter available in the tTt installer (link provided in Table of Materials). Only png, tif or jpg image formats are accepted. Follow the steps shown at Figure 1 A - B after starting the tTt Converter.
<ol>
	<li>Start the software tTt and choose a user name. Click on &#34;add user&#34; and &#34;continue&#34; button (Figure 2A). Browse and select the &#34;TTTWorkFolder&#34; (Figure 2B). Lineages trees, statistics and exported images will be saved in this folder. Next, select the experiment to be analyzed and load it (Figure 2B).</li>
	<li>If the images were converted previously by tTt Converter, click on &#34;LogFileConverter&#34; (Figure 3A) and specify the number of seconds between two consecutive time points in the LogFileConverter window. Click on &#34;Convert log files for whole experiment&#34; (Figure 1C).</li>
	<li>In a list of images, which appear in the tTt window, select the desired images that by loading manually or using the option displayed in the program. Load images (Figure 3).</li>
	<li>Select File &#62; Open &#62; New colony. Click on &#34;Tracking&#34;, followed by &#34;Start Tracking&#34; to start the tracking in a chosen position.</li>
	<li>A movie window will appear (Figure 4). Select the cell using the tracking circle and press the key 0. The software will go to the next frame. Keep on positioning the tracking circle around the tracked cell using the mouse and click 0 in each frame to add a track mark on the cell.</li>
	<li>Use the keys 1 and 3 to move to the previous or next frame, respectively. To delete a track mark, just click on the correct position to mark and press 0.</li>
	<li>If the cell divided, select the button &#34;division&#34; (red rectangle at Figure 4). Continue tracking one daughter cell by pressing Shift+D. To track the second daughter cell, select it in the cell editor and press F2.</li>
	<li>If the cell died during the video microscopy, select &#34;apoptosis&#34; (blue rectangle at Figure 4). 
	NOTE: Although the software labels the event as &#34;apoptosis&#34;, other mechanisms may be responsible for the cell death.</li>
	<li>If the cell leaves the field of observation or intermingles with neighboring cells precluding tracking, select &#34;lost&#34; (green rectangle at Figure 4).</li>
	<li>To stop tracking and continue later click on &#34;interrupt&#34; (purple rectangle at Figure 4). 
	NOTE: In the Cell editor window a tree will be generated during the tracking. To save the lineage tree, select File &#62; Save &#62; Current tree (in the Cell editor window).</li>
	<li>To start tracking a different cell, click on the cell using the right mouse button or select the cell using the left mouse button and click on Tracking &#62; Start Tracking.</li>
	<li>To export the movie with the tracking data, the software should not be in &#34;tracking mode&#34;. To turn off the tracking mode, execute step 6.10. In the Movie window select Export &#62; Movie. Set up the format, range of frames, number of frames per second and bitrate. Click on &#34;Start export&#34;.</li>
</ol>
7. Quantifications
NOTE: Several measurements can be performed using video microscopy data46. Here, we describe three possibilities that are exemplified later in the &#34;representative results&#34; section.
<ol>
	<li>Quantify the cell survival for each cell lineage by dividing the number of cells alive at different time points by the total number of cells generated before these time points were reached within individual clones37.</li>
	<li>Quantify the proportion of symmetric progenitor (SP, both daughter cells continue to proliferate), asymmetric (A, one daughter cell continues to proliferate and the other becomes postmitotic), or symmetric terminal (ST, both daughter cells become postmitotic)15,38.</li>
	<li>Measure the cell cycle length by calculating the time span of proliferating cells between their generation and division15.</li>
</ol>

The authors have nothing to disclose.

Real time observation of primary cerebral cortex cells allows the analysis of cell proliferation, mode of cell division, cell cycle length, cell differentiation and cell survival3,14,15,37. More importantly, it permits the study of single-cell lineages, leading to the identification of the intermediate phases enacted during the progression from NSCs to neurons3. Finally,...

Neural stem cells (NSC) generate neurons and macroglial cells during cerebral cortex development. At early-corticogenesis, NSCs undergo several rounds of symmetric cell division, and expand the progenitor pool. Then, NSCs divide asymmetrically to generate neurons directly or indirectly through intermediates1. Only at mid- to late-corticogenesis, progenitors switch to generate astrocytes and oligodendrocytes2,3,4. However, the complete mechanisms that control cell proliferation and differentiation, as well as the contribution of fate-restricted progenitors to the generation of unique types of neurons or macroglial cells remain a matter of intense debate4,5,6. The potential of individual cortical NSCs to generate neurons, astrocytes and oligodendrocytes has been extensively studied in vitro and in vivo using a myriad of techniques such as: live imaging in single-cell cultures7,8,9,10,11,12,13; live imaging in high-density cultures cultures3,14,15; live imaging in slice cultures16,17,18; clonal analysis using viral vector-mediated genetic labeling in high-density cultures14,15,19,20,21; clonal analysis in vivo using retrovirus22,23,24,25,26,27,28,29,30,31,32,33; and clonal analysis in vivo using transgenic animals34.
Each of these techniques presents pros and cons. For instance, in vivo lineage tracing is susceptible to the lumping and splitting errors3, leading to conflicting conclusions about the potential of individual cortical progenitors. Moreover, both in vitro and in vivo studies based on the labeling of progenitor cells at early time-points and posterior analysis of cell lineages may be influenced by the undetected occurrence of cell death during lineage-progression35. Therefore, a suitable system to analyze the potential of single NSCs must allow the identification of all cells generated, as well as the appropriate characterization of cell fates within the lineage. Combination of primary cell culture and live imaging provides this setting. Using single-cell culture and time-lapse video microscopy, Temple et al. have shown the switch in the lineage of individual cerebral cortex progenitors from neurogenesis to gliogenesis11. Later, they used the same system to show that different types of neurons are generated from a single cortical progenitor12. However, this system presents an important caveat: only 1% of cortical progenitors isolated at early corticogenesis generate clones of 4 or more progeny9. After the addition of FGF2, the frequency of cells generating 4 or more cells increases to 8 - 10%9. Nevertheless, this number is too small considering that virtually all cortical progenitors are proliferative at this stage. Moreover, the potential effects of FGF2 on fate-specification cannot be ruled out36. To circumvent these limitations, we used high-density cell cultures that support the proliferation of both ventricular (Pax6-expressing) and subventricular (Tbr2-expressing) cortical progenitors15. Moreover, the real-time observation of these cultures has shown that several features of NSC lineage progression are reproduced under these conditions, such as mode of cell division, lengthening of cell cycle, potential of single cells to generate neurons and glia, among others3,15. More recently, we have also used this system to show that CREB-signaling affects the cell survival of immature cerebral cortex neurons in mice37. Thus, we believe that time-lapse video-microscopy of primary murine cerebral cortex cells grown in high density is a powerful and accessible tool to study cellular and molecular mechanisms of cell cycle progression, mode of cell division, cell survival and cell fate specification. The latter can be accomplished using transgenic animals, allowing the identification of specific cell fates on real time38,39,40 or the use of post-imaging immunocytochemistry3,38,41,42.
Here, we provide a step-by-step protocol to prepare primary cerebral cortex cell culture supporting proliferation of NSCs and the subsequent generation of neurons and macroglial cells. We also discuss the use of retroviral-mediated transfection to manipulate gene expression of individual cells, which can be identified and tracked at the single-cell level using time-lapse video microscopy. This protocol can be used to study primary cerebral cortex cells isolated from the beginning to the end of the corticogenesis in rodents, but a few adjustments may be required according to the stage14. NSCs isolated from other sources can also be studied using time-lapse video microscopy of 2D cultures, but the appropriate culturing system should be determined by comparing cell behaviors in vitro and in vivo38,43.

<table width="953">
	<tbody>
		<tr height="40">
			<td height="40" style="width:320px;height:40px;">Hank&#39;s Balanced Salt Solution (HBSS)</td>
			<td style="width:123px;">Invitrogen Life Technologies</td>
			<td style="width:344px;">14175129</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">HEPES</td>
			<td style="width:123px;">Sigma-Aldrich</td>
			<td style="width:344px;">H3375-25G</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Penicillin/streptomycin</td>
			<td style="width:123px;">Gibco</td>
			<td>15140122</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Dulbecco Modified Eagle&#39;s Medium (DMEM)</td>
			<td style="width:123px;">Gibco</td>
			<td>12400-024</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Fetal Calf Serum (FCS)</td>
			<td style="width:123px;">Gibco</td>
			<td>10437028</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Glucose</td>
			<td style="width:123px;">Gibco</td>
			<td>A2494001</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">B27</td>
			<td style="width:123px;">Gibco</td>
			<td>17504044</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">trypsin-EDTA (0.05%)</td>
			<td style="width:123px;">Gibco</td>
			<td style="width:344px;">25300054</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Paraformaldehyde</td>
			<td>Sigma</td>
			<td style="width:344px;">16005</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="width:320px;height:40px;">Goat serum</td>
			<td style="width:123px;">Sigma-Aldrich</td>
			<td style="width:344px;">69023</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="width:320px;height:40px;">Triton X-100</td>
			<td style="width:123px;">VWR International Ltd.</td>
			<td style="width:344px;">306324N</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Isoflurane</td>
			<td>Sigma</td>
			<td>792632</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">anti-MAP2, mouse</td>
			<td>Sigma</td>
			<td>M4403</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">anti-GFP chicken</td>
			<td>Aves</td>
			<td>0511FP12</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">DAPI</td>
			<td>Sigma</td>
			<td>D9542</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Goat anti-mouse alexa 594</td>
			<td>Invitrogen</td>
			<td>A11005</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Goat anti-chicken alexa 488</td>
			<td>Invitrogen</td>
			<td>A11039</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="width:320px;height:40px;">ImageJ</td>
			<td style="width:123px;">NIH</td>
			<td style="width:344px;"></td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">tTt</td>
			<td>ETH Zurich</td>
			<td style="width:344px;"></td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Cell observer microscope</td>
			<td>Zeiss</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Pasteur pipette</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PBS</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">The Tracking Tool (tTt) software</td>
			<td></td>
			<td>https://www.bsse.ethz.ch/csd/software/ttt-and-qtfy.html</td>
			<td>download link</td>
		</tr>
	</tbody>
</table>

live imaging of primary cerebral cortex cells using a 2d culture system

During cerebral cortex development, progenitor cells undergo several rounds of symmetric and asymmetric cell divisions to generate new progenitors or postmitotic neurons. Later, some progenitors switch to a gliogenic fate, adding to the astrocyte and oligodendrocyte populations. Using time-lapse video-microscopy of primary cerebral cortex cell cultures, it is possible to study the cellular and molecular mechanisms controlling the mode of cell division and cell cycle parameters of progenitor cells. Similarly, the fate of postmitotic cells can be examined using cell-specific fluorescent reporter proteins or post-imaging immunocytochemistry. More importantly, all these features can be analyzed at the single-cell level, allowing the identification of progenitors committed to the generation of specific cell types. Manipulation of gene expression can also be performed using viral-mediated transfection, allowing the study of cell-autonomous and non-cell-autonomous phenomena. Finally, the use of fusion fluorescent proteins allows the study of symmetric and asymmetric distribution of selected proteins during division and the correlation with daughter cells fate. Here, we describe the time-lapse video-microscopy method to image primary cerebral cortex murine cells for up to several days and analyze the mode of cell division, cell cycle length and fate of newly generated cells. We also describe a simple method to transfect progenitor cells, which can be applied to manipulate genes of interest or simply label cells with reporter proteins.

Primary cultures of cerebral cortex cells isolated from embryos E14 contain both progenitor and neuronal cells. During the period of imaging, progenitors undergo several rounds of cell division, increasing the number of cells (Figure 5 and Video Figure 1).
Retroviral-mediated transfection of a few progenitor cells facilitates the identification of cell clones (F...

Watch this Scientific Journal Video about Live Imaging of Primary Cerebral Cortex Cells Using a 2D Culture System at JoVE.com

Live Imaging of Primary Cerebral Cortex Cells Using a 2D Culture System

Live imaging is a powerful tool to study cellular behaviors in real time. Here, we describe a protocol for time-lapse video-microscopy of primary cerebral cortex cells that allows a detailed examination of the phases enacted during the lineage progression from primary neural stem cells to differentiated neurons and glia.

During cerebral cortex development, progenitor cells undergo several rounds of symmetric and asymmetric cell divisions to generate new progenitors or ...

live-imaging-primary-cerebral-cortex-cells-using-2d-culture

Research

JoVE Journal

Neuroscience

8.0K Views.  Federal University of Rio Grande do Norte. Live imaging is a powerful tool to study cellular behaviors in real time. Here, we describe a protocol for time-lapse video-microscopy of primary cerebral cortex cells that allows a detailed examination of the phases enacted during the lineage progression from primary neural stem cells to differentiated neurons and glia.

Video: Live Imaging of Primary Cerebral Cortex Cells Using a 2D Culture System

Magnetic Resonance Spectroscopy of live Drosophila melanogaster using Magic Angle Spinning

High-Resolution Magic Angle Spinning (HRMAS) proton magnetic resonance spectroscopy (1H-MRS) is a novel non-destructive technique that improves spectral line-widths and allows high-resolution spectra to be obtained from extracts, intact cells, cell cultures, and more importantly intact tissue to investigate relationships between metabolites and cellular processes. In vivo HRMAS 1H-MRS studies have yet to be reported in the live fruit fly Drosophila melanogaster. Drosophila, as a simpler genetic organism, allows the multiple biological functions and various evolutionarily conserved signaling pathways to be examined at the whole organism level and it is a useful model for investigating genetics and physiology. To this end, we developed and implemented an in vivo HRMAS 1H-MRS method to investigate live Drosophila at 14.1 T. Here, we outline an HRMAS 1H-MRS protocol for the molecular characterization of Drosophila with a conventional MR spectrometer equipped with an HRMAS probe. This technique is a novel, in vivo, non-destructive Drosophila metabolite measurement approach, which enables the identification of disease biomarkers and thus may contribute to novel therapeutic development.

Magnetic Resonance Spectroscopy of live Drosophila melanogaster using Magic Angle Spinning

This technique enables the use of high-resolution magic angle spinning proton MR spectroscopy (HRMAS 1H-MRS) for molecular characterization of live Drosophila melanogaster with a conventional 14.1 tesla spectrometer equipped with an HRMAS probe.

High-Resolution Magic Angle Spinning (HRMAS) proton magnetic resonance spectroscopy (1H-MRS) is a novel non-destructive technique that improves spectral ...

Chronic Imaging of Mouse Visual Cortex Using a Thinned-skull Preparation

In vivo imaging using two-photon laser scanning microscopy (2PLSM) allows the study of living cells and neuronal processes in the intact brain. The technique presented here allows the imaging of the same area of the brain at several time points (chronic imaging) with microscopic resolution allowing the tracking of dendritic spines which are the small structures that represent the majority of postsynaptic excitatory sites in the CNS. The ability to clearly resolve fine cortical structures over several time points has many advantages, specifically in the study of brain plasticity in which morphological changes at synapses and circuit remodeling may help explain underlying mechanisms. In this video and supplementary material, we show a protocol for chronic in vivo imaging of the intact brain using a thinned-skull preparation. The thinned-skull preparation is a minimally invasive approach, which avoids potential damage to the dura and/or cortex, thus reducing the onset of an inflammatory response. When this protocol is performed correctly, it is possible to clearly monitor changes in dendritic spine characteristics in the intact brain over a prolonged period of time.

In this video and supplemental material, we show a protocol for chronic in vivo imaging of the intact brain using a thinned-skull preparation.

In vivo imaging using two-photon laser scanning microscopy (2PLSM) allows the study of living cells and neuronal processes in the intact brain. The ...

Live-imaging of PKC Translocation in Sf9 Cells and in Aplysia Sensory Neurons

Protein kinase Cs (PKCs) are serine threonine kinases that play a central role in regulating a wide variety of cellular processes such as cell growth and learning and memory. There are four known families of PKC isoforms in vertebrates: classical PKCs (&alpha;, &beta;I, &beta;II and &gamma;), novel type I PKCs (&#949; and &eta;), novel type II PKCs (&delta; and &theta;), and atypical PKCs (&zeta; and &iota;). The classical PKCs are activated by Ca2+ and diacylclycerol (DAG), while the novel PKCs are activated by DAG, but are Ca2+-independent. The atypical PKCs are activated by neither Ca2+ nor DAG. In Aplysia californica, our model system to study memory formation, there are three nervous system specific PKC isoforms one from each major class, namely the conventional PKC Apl I, the novel type I PKC Apl II and the atypical PKC Apl III. PKCs are lipid-activated kinases and thus activation of classical and novel PKCs in response to extracellular signals has been frequently correlated with PKC translocation from the cytoplasm to the plasma membrane. Therefore, visualizing PKC translocation in real time in live cells has become an invaluable tool for elucidating the signal transduction pathways that lead to PKC activation. For instance, this technique has allowed for us to establish that different isoforms of PKC translocate under different conditions to mediate distinct types of synaptic plasticity and that serotonin (5HT) activation of PKC Apl II requires production of both DAG and phosphatidic acid (PA) for translocation 1-2. Importantly, the ability to visualize the same neuron repeatedly has allowed us, for example, to measure desensitization of the PKC response in exquisite detail 3. In this video, we demonstrate each step of preparing Sf9 cell cultures, cultures of Aplysia sensory neurons have been described in another video article 4, expressing fluorescently tagged PKCs in Sf9 cells and in Aplysia sensory neurons and live-imaging of PKC translocation in response to different activators using laser-scanning microscopy.

In this video, we demonstrate visualization of PKC translocation in living cells using fluorescently tagged PKCs.

Protein kinase Cs (PKCs) are serine threonine kinases that play a central role in regulating a wide variety of cellular processes such as cell growth and ...

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

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

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

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

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

Detection of Microregional Hypoxia in Mouse Cerebral Cortex by Two-photon Imaging of Endogenous NADH Fluorescence

The brain's ability to function at high levels of metabolic demand depends on continuous oxygen supply through blood flow and tissue oxygen diffusion. Here we present a visualized experimental and methodological protocol to directly visualize microregional tissue hypoxia and to infer perivascular oxygen gradients in the mouse cortex. It is based on the non-linear relationship between nicotinamide adenine dinucleotide (NADH) endogenous fluorescence intensity and oxygen partial pressure in the tissue, where observed tissue NADH fluorescence abruptly increases at tissue oxygen levels below 10 mmHg1. We use two-photon excitation at 740 nm which allows for concurrent excitation of intrinsic NADH tissue fluorescence and blood plasma contrasted with Texas-Red dextran. The advantages of this method over existing approaches include the following: it takes advantage of an intrinsic tissue signal and can be performed using standard two-photon in vivo imaging equipment; it permits continuous monitoring in the whole field of view with a depth resolution of ~50 &mu;m. We demonstrate that brain tissue areas furthest from cerebral blood vessels correspond to vulnerable watershed areas which are the first to become functionally hypoxic following a decline in vascular oxygen supply. This method allows one to image microregional cortical oxygenation and is therefore useful for examining the role of inadequate or restricted tissue oxygen supply in neurovascular diseases and stroke.

Here we describe a method to directly visualize microregional tissue hypoxia in the mouse cortex in vivo. It is based on concurrent two-photon imaging of nicotinamide adenine dinucleotide (NADH) and the cortical microcirculation. This method is useful for high resolution analysis of tissue oxygen supply.

The brain's ability to function at high levels of metabolic demand depends on continuous oxygen supply through blood flow and tissue oxygen diffusion. ...

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date this tool has been widely used to study the regulation of cellular proliferation, differentiation and neuronal migration especially in the developing cerebral cortex 6-8. Here we detail our protocol to electroporate in utero the cerebral cortex and the hippocampus and provide evidence that this approach can be used to study dendrites and spines in these two cerebral regions.
Visualization and manipulation of neurons in primary cultures have contributed to a better understanding of the processes involved in dendrite, spine and synapse development. However neurons growing in vitro are not exposed to all the physiological cues that can affect dendrite and/or spine formation and maintenance during normal development. Our knowledge of dendrite and spine structures in vivo in wild-type or mutant mice comes mostly from observations using the Golgi-Cox method 9. However, Golgi staining is considered to be unpredictable. Indeed, groups of nerve cells and fiber tracts are labeled randomly, with particular areas often appearing completely stained while adjacent areas are devoid of staining. Recent studies have shown that IUE of fluorescent constructs represents an attractive alternative method to study dendrites, spines as well as synapses in mutant / wild-type mice 10-11 (Figure 1A). Moreover in comparison to the generation of mouse knockouts, IUE represents a rapid approach to perform gain and loss of function studies in specific population of cells during a specific time window. In addition, IUE has been successfully used with inducible gene expression or inducible RNAi approaches to refine the temporal control over the expression of a gene or shRNA 12. These advantages of IUE have thus opened new dimensions to study the effect of gene expression/suppression on dendrites and spines not only in specific cerebral structures (Figure 1B) but also at a specific time point of development (Figure 1C).
Finally, IUE provides a useful tool to identify functional interactions between genes involved in dendrite, spine and/or synapse development. Indeed, in contrast to other gene transfer methods such as virus, it is straightforward to combine multiple RNAi or transgenes in the same population of cells. 
In summary, IUE is a powerful method that has already contributed to the characterization of molecular mechanisms underlying brain function and disease and it should also be useful in the study of dendrites and spines.

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

This article describes in detail a protocol to electroporate in utero the cerebral cortex and the hippocampus at E14.5 in mice. We also show that this is a valuable method to study dendrites and spines in these two cerebral regions.

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date ...

Functional Imaging of Auditory Cortex in Adult Cats using High-field fMRI

Current knowledge of sensory processing in the mammalian auditory system is mainly derived from electrophysiological studies in a variety of animal models, including monkeys, ferrets, bats, rodents, and cats. In order to draw suitable parallels between human and animal models of auditory function, it is important to establish a bridge between human functional imaging studies and animal electrophysiological studies. Functional magnetic resonance imaging (fMRI) is an established, minimally invasive method of measuring broad patterns of hemodynamic activity across different regions of the cerebral cortex. This technique is widely used to probe sensory function in the human brain, is a useful tool in linking studies of auditory processing in both humans and animals and has been successfully used to investigate auditory function in monkeys and rodents. The following protocol describes an experimental procedure for investigating auditory function in anesthetized adult cats by measuring stimulus-evoked hemodynamic changes in auditory cortex using fMRI. This method facilitates comparison of the hemodynamic responses across different models of auditory function thus leading to a better understanding of species-independent features of the mammalian auditory cortex.

Functional studies of the auditory system in mammals have traditionally been conducted using spatially-focused techniques such as electrophysiological recordings. The following protocol describes a method of visualizing large-scale patterns of evoked hemodynamic activity in the cat auditory cortex using functional magnetic resonance imaging.

Current knowledge of sensory processing in the mammalian auditory system is mainly derived from electrophysiological studies in a variety of animal ...

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

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

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

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

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

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