The authors wish to acknowledge funding from the Lundbeck Foundation Research initiative on Brain Barriers and Drug Delivery (RIBBDD) and Simon Hougners Family Foundation.

1. Preparation of buffers and solutions for cell culturing
<ol>
	<li>Prepare collagen stock solution by dissolving 5 mg of collagen IV from human placenta in 50 mL of PBS overnight at 4 &#176;C. Aliquot the stock solution into 5 mL portions and store at -20 &#176;C.</li>
	<li>Prepare fibronectin stock solution by dissolving 5 mg of fibronectin in 5 mL of sterile water overnight. Store the fibronectin stocks in aliquots of 500 &#181;L at -20 &#176;C. When thawing, add PBS to a final volume of 50 mL to prepare the work solution and store it at 4 &#176;C.</li>
	<li>Prepare Dulbecco&#39;s Modified Eagle Medium (DMEM) complete medium by adding 50 mL of fetal bovine serum (FBS), 5 mL of MEM nonessential amino acids and 5 mL of penicillin/streptomycin (0.1 g/L streptomycin sulfate and 100,000 U/L penicillin G sodium) to 500 mL of DMEM.</li>
	<li>Prepare 5 mg/mL heparin stock solution by dissolving heparin sodium salt in PBS and pass it through a 0.2 &#181;m filter for sterilization. Store the stock solution at 4 &#176;C.</li>
	<li>Prepare growth medium (GM) immediately before use; mix 10 mL of DMEM-comp and 250 &#181;L of heparin stock solution per T75-flask.</li>
</ol>
2. Isolation of capillaries from fresh bovine brain
NOTE: Bovine brain capillaries are isolated and cultured as previously described (Helms et al.10).
<ol>
	<li>Collect brains from calves, no older than 12 months of age, from a slaughterhouse and bring directly to the lab on ice.</li>
	<li>Remove the meninges and collect all gray matter from the brain using a scalpel. Identify the meninges as the film covering the brain and the gray matter by its gray color.</li>
	<li>Use a 40 mL Dounce tissue grinder to homogenize the gray matter in Dulbecco&#39;s Modified Eagle Medium (DMEM). Fill the slim part of the tissue grinder 1/5 with gray matter suspension and add DMEM until the slim part is filled.</li>
	<li>Separate the capillaries from free cells and smaller tissue pieces by filtration of the homogenate through a 160 &#181;m nylon net filter. Flush the filters with DMEM-comp. Retrieve the capillaries and pool the suspensions into 50 mL centrifugation tubes.</li>
	<li>Resuspend the capillaries in DMEM-comp and add an enzyme mix of DNase I (170 U/mL), collagenase type III (200 U/mL) and trypsin (90 U/mL). Leave the suspension for 1 h in a 37 &#176;C water bath for digestion of the capillaries.</li>
	<li>Run the suspension through a 200 &#181;m mesh filter and resuspend in FBS with 10% dimethyl sulfoxide (DMSO). Freeze the capillaries overnight at -80 &#176;C and move them to liquid nitrogen the day after for long term storage. 
	NOTE: The protocol can be paused here.</li>
</ol>
3. Seeding and culturing of bovine capillaries
<ol>
	<li>Day 0: Mix 0.7 mL of collagen IV stock with 6.3 mL of PBS. Add the solution to a T75-flask and leave the flask for 2 h at room temperature (RT) or leave it overnight at 4 &#176;C.</li>
	<li>Remove the collagen solution from the flask and wash three times with PBS.</li>
	<li>Add 7 mL of fibronectin work solution and leave the flask for 30 min at RT. Then, remove the fibronectin solution and seed the capillaries immediately after.</li>
	<li>During the 30 min waiting time, thaw one vial of capillaries in a 37 &#176;C water bath.</li>
	<li>When the capillaries are thawed, transfer immediately to a centrifugation tube with 30 mL of DMEM-comp and centrifuge for 5 min at 500 x g and RT. Remove DMEM-comp from the tube and re-suspend the capillary-pellet in 10 mL of fresh DMEM-comp.</li>
	<li>Transfer the 10 mL suspension to the coated T75-flask and leave the capillaries to adhere to the bottom of the flask for 4-6 h in a 37 &#176;C incubator at 10% CO2. 
	NOTE: The cell growth rate is higher at 10% CO2 rather than the conventional 5% CO2.</li>
	<li>After 4-6 h of incubation inspect the flask under a light microscope. Fractions of capillaries should now be attached to the bottom of the flask (Figure 1, day 0).</li>
	<li>Prepare GM and aspirate the DMEN-comp medium very careful from the capillaries and replace it with 10 mL of freshly made GM.</li>
	<li>Day 2: Remove GM from the capillaries and replace with 10 mL of freshly made GM. Cellular outgrowth from the capillaries should be visible under a light microscope at this point (Figure 1, day 2-3).</li>
</ol>
4. Isolation of primary pericytes from bovine brain capillaries
<ol>
	<li>Day 4: Inspect the capillaries under a light microscope. 
	NOTE: The flask should now be approximately 60-70% confluent to provide an appropriate amount of pericytes (Figure 1, day 4). If this is not the case; replace the GM with 10 mL of fresh medium and leave the flask in the incubator for another day.</li>
	<li>Aspirate the medium and wash the cells gently in PBS.</li>
	<li>Add 2 mL of thawed Trypsin-EDTA for endothelial cells and leave the flask in the incubator for 1-3 min. Take out the flask frequently and observe with the microscope during this time period. 
	NOTE: The endothelial cells should round up and detach from the flask; pericytes should be visible as cells with a &#34;ghost&#34;-morphology and still be attached to the surface of the flask. This is a tricky and important step. It is essential to remove most endothelial cells to avoid contamination of the pericyte monoculture, but prolonged trypsinization can also detach the pericytes. The trypsinization time can vary slightly from time to time, and it is therefore of utmost importance to observe the flask frequently with the microscope during the treatment.</li>
	<li>Gently tap the flask, when the endothelial cells have started to round up, to detach the loosened endothelial cells.</li>
	<li>To stop the trypsinization, add 10 mL of DMEM-comp to the flask. Flush the flask carefully a few times with the medium to remove the endothelial cells. Aspirate the endothelial cell suspension from the flask. The endothelial cells can now be used for other purposes.</li>
	<li>Add 10 mL of DMEM-comp to the flask. Look under the light microscope to assure the pericytes are still present and attached to the bottom. Put the flask back into the incubator to allow the pericyte-enriched culture to grow. 
	NOTE: It is important to observe the culture during the following days. If there is still a fair amount of endothelial cells growing another trypsin-treatment can be performed.</li>
	<li>Allow the pericyte monoculture to grow with change of DMEM-comp. medium every second day. Check the growth of the cells under the light microscope (Figure 1, day 5-8).</li>
</ol>
5. Generation and storage of a monoculture of primary bovine pericytes
<ol>
	<li>Day 8-9: Inspect the capillaries under a light microscope 
	NOTE: The pericytes should now have reached 70-80% confluency and grow in islands in the flask (Figure 1, day 9). If the confluency of the pericytes is less than 70%, allow the cells to grow for another day. The pericytes will not form a complete monolayer as the endothelial cells would.</li>
	<li>Aspirate DMEM-comp and wash the pericytes with 7 mL of PBS.</li>
	<li>Add 2 mL of trypsin-EDTA to the flask and leave it in the incubator for 2-3 min. Place the flask frequently under the light microscope to observe when the pericytes round up and detach from the flask. When the pericytes have started to round up, the flask can be gently tapped to detach the cells.</li>
	<li>Gently tap the flask, when the pericytes have started to round up, to detach the cells.</li>
	<li>Add 10 mL of DMEM-comp to the flask to stop the trypsinization process. Flush the flask a few times with the medium to help detach the last pericytes.</li>
	<li>Transfer the 12 mL cell suspension to a 50 mL centrifugation tube and fill up to 30 mL with DMEM-comp.</li>
	<li>Centrifuge the cell suspension for 5 min at 500 x g and RT. Aspirate the DMEM-comp. carefully without touching the cell pellet. Resuspend the cell pellet in 3 mL of FBS with 10% DMSO.</li>
	<li>Transfer the cell suspension into cryovials; add 1 mL to each, so there will be a total of 3 vials per T75-flask of pericytes. Freeze the pericytes at -80 &#176;C overnight and move them to liquid nitrogen the day after for long term storage. 
	NOTE: Cells may be counted before freezing for a later estimate of survival percentage. The protocol can be paused here.</li>
</ol>
6. Setting up a pericyte monoculture for experiments
<ol>
	<li>Coat a T75-flask with collagen IV and fibronectin using the same procedure as mentioned in section 3.1-3.4.</li>
	<li>While the flask is being coated with fibronectin, thaw one vial of pericytes in a 37 &#176;C water bath.</li>
	<li>Transfer the now thawed pericytes from the cryovial to a centrifugation tube with 30 mL of DMEM-comp. Centrifuge the cell suspension for 5 min at 500 x g, RT.</li>
	<li>Carefully aspirate the medium, leaving the cell pellet at the bottom of the tube. Re-suspend the pellet in 10 mL DMEM-comp.</li>
	<li>Collect and transfer the cell suspension to the coated flask. Leave the flask with pericytes to grow in a 37 &#176;C incubator at 10% CO2.</li>
	<li>Every second day, refresh the medium with 10 mL of fresh DMEM-comp. 
	NOTE: After 5 days of growth, the pericytes should have reached approximately 80% confluency. If the confluency is less, leave the cells to grow for another day or two. The cells should now be ready for seeding for further experiments.</li>
</ol>
7. Seeding of pericytes in a coated 96-well plate
<ol>
	<li>Dilute collagen IV as described in step 3.1. Add 100 &#181;L to each well in a 96-well plate and incubate for 2 h at RT or overnight at 4 &#176;C.</li>
	<li>Aspirate the solution and wash the wells three times with PBS.</li>
	<li>Add 100 &#181;L of diluted fibronectin to each well and incubate at RT for 30 min. Remove the fibronectin solution and use the plate immediately. 
	NOTE: Depending on how well the pericyte batch is growing, there should be enough cells for seeding two plates.</li>
	<li>Take out the pericytes from the incubator and aspirate the medium. Wash the cells with PBS.</li>
	<li>Add 2 mL of trypsin-EDTA to the pericytes and follow same procedure as in step 5.3-5.6.</li>
	<li>Aspirate the medium, without harming the cell pellet and re-suspend the pellet in 1 mL of fresh DMEM-comp.</li>
	<li>Take out 12 &#181;L of cell suspension and add to a counting chamber. Under the light microscope, count at least 3 of 3x3 grids and use the average cell count per grid.</li>
	<li>Use the equation below to calculate the volume of cell suspension that should be added to each well to seed 10.000 cells per well, in the 96-well plate. 
	<img src="/files/ftp_upload/61253/61253eq1v2.jpg" /></li>
	<li>Add DMEM-comp and the calculated volume of cell suspension in each well to reach a final volume of 200 &#181;L.</li>
	<li>Place the 96-well plate in a 37 &#176;C incubator at 10% CO2. Leave the cells to grow for 4 days with a change of medium after 2 days.</li>
</ol>
8. Preparation of buffers and solutions for Ca2+-imaging
<ol>
	<li>Autoclave coverslip cell chambers and coverslips.</li>
	<li>Assay buffer: Add 1.19 g of HEPES to 500 mL of HBSS buffer for a final concentration of 10 mM HEPES. Adjust the pH to 7.4.</li>
	<li>Prepare 20% (w/v) Pluronic F127 + 1% (v/v) polyethoxylated castor oil stock solution by dissolving 0.5 g of Pluronic F127 solution in 2.5 mL of anhydrous DMSO in a glass vial. Heat to 40 &#176;C for approximately 30 min or until dissolved and vortex. Add 25 &#181;L of polyethoxylated castor oil and store at RT. Do not freeze.</li>
	<li>Prepare 2 mM Fura-2 AM stock by dissolving 1 mg in 500 &#181;L of anhydrous DMSO. Store in aliquots of 20 &#181;L at -20 &#176;C protected from light.</li>
	<li>Prepare 5 &#181;M Fura-2 AM loading solution by mixing 20 &#181;L of 20% Pluronic F-127 + 1% polyethoxylated castor oil stock solution with the 20 &#181;L of 2 mM Fura-2 AM aliquot. Add 500 &#181;L of assay buffer and vortex. Add assay buffer to a final volume of 8 mL. The solution should be prepared immediately before use and protected from light.</li>
	<li>Prepare 4 mM Cal-520 AM by dissolving 1 mg in 226.7 &#181;L of anhydrous DMSO. Store in aliquots of 20 &#181;L at -20 &#176;C protected from light.</li>
	<li>Prepare 20 &#181;M Cal-520 AM loading solution by mixing 20 &#181;L of 20% Pluronic F-127 + 1% polyethoxylated castor oil stock solution with the 20 &#181;L 4 mM Cal-520 aliquot. Add 500 &#181;L of assay buffer and vortex. Add assay buffer to a final volume of 4 mL. The solution should be prepared immediately before use and protected from light.</li>
</ol>
9. Loading of pericytes with Fura-2 AM calcium indicator dye in a plate-reader setup
NOTE: All solutions should be at RT before the experiment starts.
<ol>
	<li>Take out the 96-well plate with cells from the incubator and aspirate the medium from the wells. Wash the cells twice with assay buffer.</li>
	<li>Add 100 &#181;L of loading solution to each well and wrap the plate with tinfoil, to avoid photo bleaching. Incubate for 45 min with 30 rpm shaking at RT. 
	NOTE: Do not load Fura-2 AM at 37 &#176;C, as it may load internal compartments. Remember to leave wells with cells in assay buffer instead of loading buffer; these are&#160;the &#34;blanks&#34; used for measuring background fluorescence.</li>
	<li>Aspirate the loading buffer and wash the cells with assay buffer twice. Add 100 &#181;L of fresh assay buffer and leave the cells to incubate for 30 min at RT; this allows for continuous cleavage of the AM-ester and thereby trapping Fura-2 AM inside the cells.</li>
	<li>Prior to the Ca2+-imaging, wash and replace the buffer with 100 &#181;L of fresh assay buffer.</li>
</ol>
10. Well-plate fluorescence reading of pericytes in a plate-reader setup
<ol>
	<li>Set the temperature of the plate reader to 37 &#176;C and transfer the 96-well plate with cells to the sample plate position. Place the reagent plate with agonist at the reagent plate position.</li>
	<li>Start by measuring loading of the cells to ensure equal loading of Fura-2 AM in all wells.</li>
	<li>Perform the measurements with excitation fluorescence wavelength at 340 nm/380 nm and the emission wavelength at 510 nm. Add 50 &#181;L of agonist at speed 150 &#181;L/s from the reagent plate to each well with cells in the sample plate position.</li>
	<li>Save the data and export as xlsx files for further analysis. Figure 2 shows the cytosolic Ca2+-response measured as the ratio between the two excitation wavelengths over time, where background fluorescence is subtracted. 
	NOTE: The plate-reader need to be a dual microplate reader with room for a &#34;cell tray&#34; and a &#34;sample tray&#34; and an integrated pipettor system.</li>
</ol>
11. Seeding of pericytes in a coated cell chamber for live imaging
NOTE: Coverslips may also be placed in the bottom of culture wells, coated and seeded with pericytes as described above, and then mounted in the chamber prior to experiments.&#160;&#160;
<ol>
	<li>Mount a coverslip into the cell chamber and make it tight to avoid leakiness.</li>
	<li>Dilute collagen IV as described in step 3.1. Add 500 &#181;L to each cell chamber and incubate for 2 h at RT or overnight at 4 &#176;C.</li>
	<li>Aspirate the collagen solution and wash the chambers three times with 500 &#181;L of PBS.</li>
	<li>Add 500 &#181;L of diluted fibronectin to each well and incubate at RT for 30 min. Remove the fibronectin solution and use the cell chamber straight afterwards.</li>
	<li>In the meantime, take out the flask with confluent pericytes and wash with 7 mL of PBS.</li>
	<li>Add 2 mL of trypsin-EDTA to the pericytes and follow same procedure as in step 5.3-5.6.</li>
	<li>Proceed by following the same steps as in step 8.6-8.7.</li>
	<li>Use the equation below to calculate the volume of cell suspension, which should be added to each chamber to seed 90.000 cells per chamber. 
	<img src="/files/ftp_upload/61253/61253eq2v2.jpg" /></li>
	<li>Add DMEM-comp and the calculated volume of cell suspension in each chamber to reach a final volume of 500 &#181;L.</li>
	<li>Place the cell chambers in the incubator at 37 &#176;C, 10% CO2. Leave the cells to grow for 6 days (or until confluent). 
	NOTE: The pericytes grow slower on glass-slides compared to plastic; more days of growth are necessary.</li>
</ol>
12. Loading of pericytes with Cal-520 AM calcium indicator dye for live imaging
NOTE: All solutions should be at RT before the experiment starts.
<ol>
	<li>Prepare the 20 &#181;M Cal-520 AM loading buffer: Mix 20 &#181;L of 20% Pluronic F-127 + 1% polyethoxylated castor oil stock solution with the 20 &#181;L 4 mM Cal-520 aliquot. Add 500 of &#181;L assay buffer and vortex. Add assay buffer to a final volume of 4 mL. The solution should be prepared immediately before use and protected from light. 
	NOTE: Protect solutions containing Cal-520 AM from light exposure.</li>
	<li>Take out the cell chambers from the incubator and aspirate the medium. Wash the cells twice with assay buffer.</li>
	<li>Add 500 &#181;L of loading buffer to each chamber and incubate at RT for 45 min.</li>
	<li>Aspirate the loading buffer and wash the cells twice with assay buffer.</li>
	<li>Add 500 &#181;L of fresh assay buffer to each chamber and incubate for 30 min at RT to allow cleavage of the AM-ester.</li>
	<li>Replace the buffer with 500 &#181;L of fresh assay buffer before performing the live imaging at a confocal microscope.</li>
</ol>
13. Live imaging of intracellular Ca2+-levels
NOTE: A variety of microscope types can be used for the imaging. Upright or inverted conventional fluorescence microscopes, as well as upright or inverted confocal laser scanning microscopes with appropriate excitation source (488 nm) and emission filters (510-520 nm) can be used. Objectives should be suited for fluorescence and be of a high quality and with high numerical aperture (NA).
<ol>
	<li>Mount the cell chamber on the stage of the confocal microscope as gentle as possible, in order to avoid disturbance of the cells.</li>
	<li>Select an excitation wavelength of 488 nm, emission at 515 nm, sequential image acquisition with 5 second intervals, an XY image size of 512 x 512 pixels and measure for 2 min to measure baseline calcium signals.</li>
	<li>Add 3 &#181;L of 100 mM ATP to the cell chamber with a pipette, and continue the sequential image acquisition. Perform the addition slowly and gently to not disturb the preparation and move the cells out of focus.</li>
	<li>Observe the degree of changes and increase the time interval over time as needed for approximately 18 min until no further morphological change is noted (Figure 3).</li>
	<li>Save time-lapse images and export them as TIFF and/or AVI files for further analysis. 
	NOTE: One vial of pericytes should give enough cells for seeding in 1-2 96-well plates and several coverslips, meaning you can prepare cells for both types of calcium-measurements.</li>
</ol>

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The authors declare no competing financial interests.

In this study, we have presented a method to isolate primary pericytes from bovine brains. The described protocol allows culture of this otherwise rather inaccessible cell type. The subsequently obtained cell culture was a nearly homogenous population of pericytes, with little or no contamination with endothelial cells&#160;and glial cells based on cell morphology and protein expression12. Furthermore, we demonstrated a simple and straightforward method to load the pericytes with calcium dyes for Ca2+-imaging using two different methods, depending on the intended outcome.
One of the main issues when isolating primary cells from brain tissue is the limited access to tissue. In several earlier studies, rats and mice are the traditional model animals used, but the brain tissue from these small animals are sparse6,13. Isolation from human brain has also been conducted14; however, due to ethical issues this is not easily accessible. Hence, a high cell yield from an available source is preferred. The number of cell vials obtained from isolation from a single bovine brain outnumbers the amount obtained from either mouse or rat by far, making the bovine tissue advantageous compared to the other mentioned sources. Another advantage of the method described here is the low passage number to obtain a pure culture of pericytes. Passaging of mural cells can lead to de-differentiation15 and therefore, should preferably be avoided. In this protocol, gentle trypsinization to remove the endothelial cell layer is a single step used to isolate the pericytes and therefore also one of the most critical steps described in this protocol. If the trypsinization is prolonged, the pericytes will start to detach from the flask together with the endothelial cells, leading to only a small yield. On the other hand, only allowing a very short trypsinization time can cause an impure culture of pericytes. It is therefore of utmost importance to observe the cells very frequently during the trypsinization step. One should be aware of the morphological&#160;differences between pericytes and endothelial cells to be able to distinguish the two cell types during the trypsinization procedure. Although, this step might require some training, the method is less time consuming and inexpensive compared to other methods used to obtain pure pericyte cultures7,8,14. Furthermore, the obtained culture of pericytes here showed the typical &#34;ghost&#34;-like morphology with finger-like processes and expressed several specific markers such as &#945;-SMA, PDGFR-&#946; and Nestin (data not shown here)12, which are well known markers for primary pericytes in culture1.
Here, we also presented a simple method for Ca2+-imaging using fluorescent calcium indicator dyes. Loading of the pericytes with the dyes (Fura-2 and Cal-520) should be performed at RT and not at 37 &#176;C. Several studies have performed loading of AM esters at 37 &#176;C16,17, but loading at this temperature can lead to the esters entering intracellular compartments such as the endoplasmic reticulum. This is not preferable, as the Fura-2 AM can bind to free Ca2+ trapped in the internal stores and thereby result in a lower increase in the fluorescent measurements. Hence, it can cause false results. Although, we have not experienced any major difficulties with loading of the pericytes at RT, if one is experiencing problems anyhow, one might consider optimizing loading by sonication of the loading solution for 5 min in an ultrasound bath prior to loading of the cells.
With the described method, intracellular Ca2+-signaling together with contraction of the pericytes is readily observed during live recordings, which can also be quantified for further analysis. The method has previously been used to make the first demonstration of in vitro Ca2+-signaling and contraction of brain capillary pericytes12. In previous studies various methods have been used to measure and quantify cell contraction as a consequence of extracellular stimulus18,19,20. However, observing the preliminary intracellular response directly followed by contraction of the cells was not a possibility in these studies. In the current study it is possible to observe the intracellular signaling and the following contraction of the cells and both measurements can be quantified in the same experiment. Thus, it eliminates additional experiments, which are more time consuming as well, and shows the direct link between intracellular Ca2+-signaling and the morphological changes.
In conclusion, this study represents a simple and effective method for isolating and culturing primary brain pericytes. In addition, we demonstrate an easy and reproducible method to study intracellular Ca2+-signaling in cultured pericytes. The methods described here should provide other researchers in the field with strong tools to study the pericyte biology and intracellular signaling in pericytes in vitro.

Brain capillary pericytes, together with endothelial cells and astrocytes, constitute the NVU1,2,3. The endothelial cells, which form the structural basis of the capillaries, form long cylindrical tubes with a diameter of 5-8 &#181;m. The endothelial cells are sporadically covered with pericytes and surrounded by protrusions from astrocytes; the astrocyte endfeet.
The blood-brain barrier (BBB), situated at the brain capillaries, is the main site for exchange of nutrients, gases and waste products between the brain and the blood. The BBB also protects the brain from endogenous and exogenous neurotoxins and serves as a barrier for the delivery of a large number of drug compounds. The barrier function is a focus area, as well as an obstacle, for drug companies developing central nervous system (CNS) medicines. This has spurred a large interest in investigating the cells of the NVU in culture4. Brain astrocytes and endothelial cells have been cultured and characterized in a number of studies, whereas the studies and protocols for pericyte culture are sparse.
Previously published protocols have described generation of brain capillary pericyte cultures to some degree, using a range of different approaches such as immunopanning5, high- and low-glucose media6, fluorescent-activated cell sorting7, density gradient centrifugation8, etc. Although these methods seem sufficient to obtain cultures of pericytes, some are time consuming, cost expensive and the pericytes obtained might not be ideal due to the number of culture passages that can de-differentiate the pericytes9. Furthermore, the potential of cultured pericytes in in vitro signaling studies has been fairly unexplored until now.
The present work focuses on the generation of pericyte cultures from isolated bovine brain capillaries and the subsequent setup for measurements and imaging studies of changes in intracellular calcium, an important intracellular second messenger. We briefly describe the isolation of capillaries from cortical gray matter (for details see Helms et al.10) and the isolation and culture of pericytes in pure monoculture without contamination with endothelial or glial cells. We then provide a protocol for seeding of pericytes in 96-well plates and loading protocols for the calcium probe Fura-2 AM. Finally, we show how pericytes can be used in real-time confocal imaging in microscope culture chambers and describe the protocols for this.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>ATP</td>
			<td>Tocris</td>
			<td>3245</td>
			<td></td>
		</tr>
		<tr>
			<td>Cal-520 AM</td>
			<td>AAT Bioquest</td>
			<td>21130</td>
			<td></td>
		</tr>
		<tr>
			<td>Cell incubator</td>
			<td>Thermo Fisher</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge</td>
			<td>Thermo Fisher</td>
			<td>Heraeus Multifuge 3SR+</td>
			<td>Standard large volume centrifuge for spinning down cells</td>
		</tr>
		<tr>
			<td>Collagen IV</td>
			<td>Sigma Aldrich</td>
			<td>C5533</td>
			<td></td>
		</tr>
		<tr>
			<td>Confocal laser scanning microscope</td>
			<td>Carl Zeiss</td>
			<td>Zeiss LSM 510</td>
			<td>Inverted microscope</td>
		</tr>
		<tr>
			<td>Counting chamber</td>
			<td>FastRead</td>
			<td>102</td>
			<td></td>
		</tr>
		<tr>
			<td>Coverslip cell chamber</td>
			<td>Airekacells</td>
			<td>SC15022</td>
			<td></td>
		</tr>
		<tr>
			<td>Cremophor EL</td>
			<td>Sigma Aldrich</td>
			<td>C5135</td>
			<td>Formerly known as Kolliphor EL</td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Sigma Aldrich</td>
			<td>471267</td>
			<td></td>
		</tr>
		<tr>
			<td>Dulbecco&#39;s Modified Eagles Medium</td>
			<td>Sigma Aldrich</td>
			<td>D0819</td>
			<td></td>
		</tr>
		<tr>
			<td>Fetal bovine serum (FBS)</td>
			<td>PAA/GE Healthcare</td>
			<td>A15-101</td>
			<td></td>
		</tr>
		<tr>
			<td>Fibronectin</td>
			<td>Sigma Aldrich</td>
			<td>F1141</td>
			<td></td>
		</tr>
		<tr>
			<td>Fura-2 AM</td>
			<td>Thermo Fisher</td>
			<td>F1201</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass coverslips 22x22 mm</td>
			<td>VWR International</td>
			<td>631-0123</td>
			<td></td>
		</tr>
		<tr>
			<td>HBSS</td>
			<td>Gibco</td>
			<td>14065-049</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>Sigma Aldrich</td>
			<td>H3149</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>AppliChem Panreac</td>
			<td>A1069</td>
			<td></td>
		</tr>
		<tr>
			<td>Light microscope</td>
			<td>Olympus</td>
			<td>Olympus CK2</td>
			<td>Upright light microscope with phase contrast</td>
		</tr>
		<tr>
			<td>MEM nonessential amino acids</td>
			<td>Sigma Aldrich</td>
			<td>M7145</td>
			<td></td>
		</tr>
		<tr>
			<td>Microplate Reader</td>
			<td>BMG LabTech</td>
			<td>NOVOstar</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Sigma Aldrich</td>
			<td>D8537</td>
			<td>Phosphate-buffered saline</td>
		</tr>
		<tr>
			<td>penicillin G sodium/streptomycin sulfate</td>
			<td>Sigma Aldrich</td>
			<td>P0781</td>
			<td></td>
		</tr>
		<tr>
			<td>Pluronic F127</td>
			<td>Sigma Aldrich</td>
			<td>P2443</td>
			<td></td>
		</tr>
		<tr>
			<td>Trypsin-EDTA</td>
			<td>Sigma Aldrich</td>
			<td>T4299</td>
			<td></td>
		</tr>
		<tr>
			<td>T-75 flask</td>
			<td>Sigma Aldrich</td>
			<td>CLS3972</td>
			<td></td>
		</tr>
		<tr>
			<td>96-well plate</td>
			<td>Corning incorporated</td>
			<td>3603</td>
			<td></td>
		</tr>
	</tbody>
</table>

culture of brain capillary pericytes for cytosolic calcium measurements and calcium imaging studies

Pericytes are associated with endothelial cells and astrocytic endfeet in a structure known as the neurovascular unit (NVU). Brain capillary pericyte function is not fully known. Pericytes have been suggested to be involved in capillary development, regulation of endothelial barrier tightness and trancytosis activity, regulation of capillary tone and to play crucial roles in certain brain pathologies.
Pericytes are challenging to investigate in the intact brain due to the difficulties in visualizing processes in the brain parenchyma, as well as the close proximity to the other cells of the NVU. The present protocol describes a method for isolation and culture of primary bovine brain capillary pericytes and their following usage in calcium imaging studies, where effects of agonists involved in brain signaling and pathologies can be investigated. Cortical capillary fragments are allowed to attach to the bottom of culture flasks and, after 6 days, endothelial cells and pericytes have grown out from the capillary fragments. The endothelial cells are removed by gentle trypsinization and pericytes are cultured for 5 additional days before passaging.
Isolated pericytes are seeded in 96-well culture plates and loaded with the calcium indicator dye (Fura-2 acetoxymethyl (AM)) to allow for measurements of intracellular calcium levels in a plate reader setup. Alternatively, pericytes are seeded on coverslips and mounted in cell chambers. Following loading with the calcium indicator (Cal-520 AM), calcium live-imaging can be performed using confocal microscopy at an excitation wavelength of 488 nm and emission wavelength of 510-520 nm.
The method described here has been used to obtain the first intracellular calcium measurements from primary brain capillary pericytes, demonstrating that pericytes are stimulated via ATP and are able to contract in vitro.

Bovine brain capillaries were isolated from fresh brain tissue and Figure 1 presents the capillary seeding and cellular outgrowth over days and subsequent purification&#160;of pericytes. The capillaries are fully attached to the flask at day 1 and on day 2 endothelial sprouting has become visible (Figure 1, day 2). After 4 days, the cellular outgrowth is highly distinctive (Figure 1, day 4a) and the endothelial cells are removed by gentle trypsinization as according to the described protocol. Remnants of the capillaries can be present after the trypsinization, but will disappear from the flask in the following days (Figure 1, day 4b-6). After removal of the endothelial-layer, pericytes are easily detected with morphology distinct from the endothelial cells. The pericytes present finger-like processes that attach strongly to the flask (Figure 1, day 4b). Subsequently, pericytes are allowed to grow until confluency (Figure 1, day 4b-9) and on day 9, the pericytes have reached approximately 80% confluency and grow in islands. This is in contrast to the endothelial cells that do create a contact inhibited&#160;monolayer observed at day 4.
ATP is a well-known endogenous inducer of intracellular Ca2+-signaling11 and was used as an extracellular stimulant to induce cytosolic changes in Ca2+-levels in pericytes. Addition of ATP to the Fura-2 loaded pericytes, as according to the described protocol, resulted in an increase in cytosolic Ca2+-levels measured as the fluorescent ratio as shown in Figure 2. The ATP-induced response occurs immediately after addition of ATP to the pericytes and declines slowly over the measured time period.
Using this protocol for real-time confocal imaging of intracellular Ca2+-responses, pericytes were seeded on coated coverslips, loaded with Cal-520 AM and placed at the confocal microscope. Figure 3 (0 s) shows the pericytes with baseline levels of fluorescence prior to treatment. During live-recording, ATP is added to the pericytes and a strong intracellular Ca2+-response is evident shortly after (64 s). Soon after, the cytosolic Ca2+ compartmentalizes in the cells and a reduction in cell area is visible (189 s). At 300 s. post start of the recordings, the cell area is heavily reduced and the Ca2+-signal has declined to intensity close to the baseline fluorescence.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61253/61253fig1.jpg" /> 
Figure 1: Culturing of capillaries and isolation of pericytes. Capillaries have been isolated from fresh bovine brain and seeded in culture flasks on day 0. Outgrowth from the bovine brain capillaries and the following isolation of pericytes were followed over days with a light microscope. Day 4a shows the endothelial cell growth prior to treatment with trypsin to remove endothelial cells and day 4b shows the remnants immediately after the treatment. Day 8a shows the focus plane, where any capillary remnants would be visible, whereas day 8b are focused on the plane where the growth of the pericytes is visible. &#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61253/61253fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61253/61253fig2.jpg" /> 
Figure 2: Representative example of intracellular calcium-measurement using Fura-2 calcium indicator dye. Primary pericytes have been seeded in 96 well plates and loaded with Fura-2 AM in order to visualize changes in intracellular calcium. 10 &#181;M ATP is added to the pericytes at 30 s. and the cytosolic Ca2+-response is measured as the ratio between the two excitation wavelengths; 340 nm and 380 nm over time. Scale bars are defined as standard deviation (N=3, n=1). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61253/61253fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61253/61253fig3.jpg" /> 
Figure 3: Representative example of intracellular calcium live-imaging using Cal-520 calcium indicator dye. Primary pericytes have been seeded in a cell chamber and loaded with Cal-520 in order to visualize changes in intracellular calcium and cell morphology. 600 &#181;M ATP is added to the pericytes and snapshots from different time points during the live-imaging are presented here. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61253/61253fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Culture of Brain Capillary Pericytes for Cytosolic Calcium Measurements and Calcium Imaging Studies at JoVE.com

Culture of Brain Capillary Pericytes for Cytosolic Calcium Measurements and Calcium Imaging Studies

Brain capillary pericytes are essential players in the regulation of blood-brain barrier properties and blood flow. This protocol describes how brain capillary pericytes can be isolated, cultured, characterized with respect to cell type and applied for investigations of intracellular calcium signaling with fluorescent probes.

Pericytes are associated with endothelial cells and astrocytic endfeet in a structure known as the neurovascular unit (NVU). Brain capillary pericyte ...

culture-brain-capillary-pericytes-for-cytosolic-calcium-measurements

Nanyang Technological University

Research

JoVE Journal

Neuroscience

4.7K Views.  University of Copenhagen. Brain capillary pericytes are essential players in the regulation of blood-brain barrier properties and blood flow. This protocol describes how brain capillary pericytes can be isolated, cultured, characterized with respect to cell type and applied for investigations of intracellular calcium signaling with fluorescent probes.

Video: Culture of Brain Capillary Pericytes for Cytosolic Calcium Measurements and Calcium Imaging Studies

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.

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 ...

Preparation of Acute Subventricular Zone Slices for Calcium Imaging

The subventricular zone (SVZ) is one of the two neurogenic zones in the postnatal brain. The SVZ contains densely packed cells, including neural progenitor cells with astrocytic features (called SVZ astrocytes), neuroblasts, and intermediate progenitor cells. Neuroblasts born in the SVZ tangentially migrate a great distance to the olfactory bulb, where they differentiate into interneurons. Intercellular signaling through adhesion molecules and diffusible signals play important roles in controlling neurogenesis. Many of these signals trigger intercellular calcium activity that transmits information inside and between cells. Calcium activity is thus reflective of the activity of extracellular signals and is an optimal way to understand functional intercellular signaling among SVZ cells.
Calcium activity has been studied in many other regions and cell types, including mature astrocytes and neurons. However, the traditional method to load cells with calcium indicator dye (i.e. bath loading) was not efficient at loading all SVZ cell types. Indeed, the cellular density in the SVZ precludes dye diffusion inside the tissue. In addition, preparing sagittal slices will better preserve the three-dimensional arrangement of SVZ cells, particularly the stream of neuroblast migration on the rostral-caudal axis.
Here, we describe methods to prepare sagittal sections containing the SVZ, the loading of SVZ cells with calcium indicator dye, and the acquisition of calcium activity with time-lapse movies. We used Fluo-4 AM dye for loading SVZ astrocytes using pressure application inside the tissue. Calcium activity was recorded using a scanning confocal microscope allowing a precise resolution for distinguishing individual cells. Our approach is applicable to other neurogenic zones including the adult hippocampal subgranular zone and embryonic neurogenic zones. In addition, other types of dyes can be applied using the described method.

A method to load subventricular zone (SVZ) cells with calcium indicator dyes for recording calcium activity is described. The postnatal SVZ contains tightly packed cells including neural progenitor cells and neuroblasts. Rather than using bath loading we injected the dye by pressure inside the tissue allowing better dye diffusion.

The subventricular zone (SVZ) is one of the two neurogenic zones in the postnatal brain. The SVZ contains densely packed cells, including neural ...

Simultaneous Electrophysiological Recording and Calcium Imaging of Suprachiasmatic Nucleus Neurons

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in regulating the intracellular calcium concentration. Changing environmental conditions, such as the light-dark cycle, can modify neuronal and neural network activity and the expression of a family of circadian clock genes within the suprachiasmatic nucleus (SCN), the location of the master circadian clock in the mammalian brain. Excitatory synaptic transmission leads to an increase in the postsynaptic Ca2+ concentration that is believed to activate the signaling pathways that shifts the rhythmic expression of circadian clock genes. Hypothalamic slices containing the SCN were patch clamped using microelectrodes filled with an internal solution containing the calcium indicator bis-fura-2. After a seal was formed between the microelectrode and the SCN neuronal membrane, the membrane was ruptured using gentle suction and the calcium probe diffused into the neuron filling both the soma and dendrites. Quantitative ratiometric measurements of the intracellular calcium concentration were recorded simultaneously with membrane potential or current. Using these methods it is possible to study the role of changes of the intracellular calcium concentration produced by synaptic activity and action potential firing of individual neurons. In this presentation we demonstrate the methods to simultaneously record electrophysiological activity along with intracellular calcium from individual SCN neurons maintained in brain slices.

Procedures are described to perform simultaneous recordings of membrane potential or current and changes of intracellular calcium concentration. Suprachiasmatic nucleus neurons are filled with the calcium indicator bis-fura-2 using a patch clamp electrode in the whole cell patch clamp configuration.

Simultaneous electrophysiological and fluorescent imaging recording methods were used to study the role of changes of membrane potential or current in ...

Two-photon Calcium Imaging in Neuronal Dendrites in Brain Slices

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ fluctuations can occur through a variety of membrane and intracellular mechanisms and play a crucial role in the induction of synaptic plasticity and regulation of dendritic excitability. Hence, the ability to record different types of Ca2+ signals in dendritic branches is valuable for groups studying how dendrites integrate information. The advent of two-photon microscopy has made such studies significantly easier by solving the problems inherent to imaging in live tissue, such as light scattering and photodamage. Moreover, through combination of conventional electrophysiological techniques with two-photon Ca2+ imaging, it is possible to investigate local Ca2+ fluctuations in neuronal dendrites in parallel with recordings of synaptic activity in soma. Here, we describe how to use this method to study the dynamics of local Ca2+ transients (CaTs) in dendrites of GABAergic inhibitory interneurons. The method can be also applied to studying dendritic Ca2+ signaling in different neuronal types in acute brain slices.

We present a method combining whole-cell patch-clamp recordings and two-photon imaging to record Ca2+ transients in neuronal dendrites in acute brain slices.

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ ...

Ablation of a Neuronal Population Using a Two-photon Laser and Its Assessment Using Calcium Imaging and Behavioral Recording in Zebrafish Larvae

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ablation of neurons is an effective method for this purpose when neurons are selectively labeled with fluorescent probes. In the present study, protocols for laser ablating a subpopulation of neurons using a two-photon microscope and testing of its functional and behavioral consequences are described. In this study, prey capture behavior in zebrafish larvae is used as a study model. The pretecto-hypothalamic circuit is known to underlie this visually-driven prey catching behavior. Zebrafish pretectum were laser-ablated, and neuronal activity in the inferior lobe of the hypothalamus (ILH; the target of the pretectal projection) was examined. Prey capture behavior after pretectal ablation was also tested.

Here, we present a protocol to ablate a genetically labeled subpopulation of neurons by a two-photon laser from Zebrafish larvae.

To identify the role of a subpopulation of neurons in behavior, it is essential to test the consequences of blocking its activity in living animals. Laser ...

Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila

Organ-to-organ communication by endocrine signaling, for example, from the periphery to the brain, is essential for maintaining homeostasis. As a model animal for endocrine research, Drosophila melanogaster, which has sophisticated genetic tools and genome information, is being increasingly used. This article describes a method for the calcium imaging of Drosophila brain explants. This method enables the detection of the direct signaling of a hormone to the brain. It is well known that many peptide hormones act through G-protein-coupled receptors (GPCRs), whose activation causes an increase in the intracellular Ca2+concentration. Neural activation also elevates intracellular Ca2+ levels, from both Ca2+ influx and the release of Ca2+ stored in the endoplasmic reticulum (ER). A calcium sensor, GCaMP, can monitor these Ca2+ changes. In this method, GCaMP is expressed in the neurons of interest, and the GCaMP-expressing larval brain is dissected and cultured ex vivo. The test peptide is then applied to the brain explant, and the fluorescent changes in GCaMP are detected using a spinning disc confocal microscope equipped with a CCD camera. Using this method, any water-soluble molecule can be tested, and various cellular events associated with neural activation can be imaged using the appropriate fluorescent indicators. Moreover, by modifying the imaging chamber, this method can be used to image other Drosophila organs or the organs of other animals.

Ex Vivo Calcium Imaging for Visualizing Brain Responses to Endocrine Signaling in Drosophila

This paper describes a protocol for ex vivo calcium imaging of the Drosophila brain. In this method, natural or synthetic compounds can be applied to the buffer to test their ability to activate particular neurons in the brain.

Organ-to-organ communication by endocrine signaling, for example, from the periphery to the brain, is essential for maintaining homeostasis. As a model ...

Brain Pericyte Calcium and Hemodynamic Imaging in Transgenic Mice In Vivo

Recent advances in protein biology and mouse genetics have made it possible to measure intracellular calcium fluctuations of brain cells in vivo and to correlate this with local hemodynamics. This protocol uses transgenic mice that have been prepared with a chronic cranial window and express the genetically encoded calcium indicator, RCaMP1.07, under the &#945;-smooth muscle actin promoter to specifically label mural cells, such as vascular smooth muscle cells and ensheathing pericytes. Steps are outlined on how to prepare a tail vein catheter for intravenous injection of fluorescent dyes to trace blood flow, as well as how to measure brain pericyte calcium and local blood vessel hemodynamics (diameter, red blood cell velocity, etc.) by two photon microscopy in vivo through the cranial window in ketamine/xylazine anesthetized mice. Finally, details are provided for the analysis of calcium fluctuations and blood flow movies via the image processing algorithms developed by Barrett et al. 2018, with an emphasis on how these processes can be adapted to other cellular imaging data.

Brain Pericyte Calcium and Hemodynamic Imaging in Transgenic Mice In Vivo

This protocol presents steps to acquire and analyze fluorescent calcium images from brain ensheathing pericytes and blood flow data from nearby blood vessels in anesthetized mice. These techniques are useful for studies of mural cell physiology and can be adapted to investigate calcium transients in any cell type.

Recent advances in protein biology and mouse genetics have made it possible to measure intracellular calcium fluctuations of brain cells in vivo and to ...

Stereotaxic Viral Injection and Gradient-Index Lens Implantation for Deep Brain In Vivo Calcium Imaging

A miniature fluorescence microscope (miniscope) is a potent tool for in vivo calcium imaging from freely behaving animals. It offers several advantages over conventional multi-photon calcium imaging systems: (1) compact; (2) light-weighted; (3) affordable; and (4) allows recording from freely behaving animals. This protocol describes brain surgeries for deep brain in vivo calcium imaging using a custom-developed miniscope recording system. The preparation procedure consists of three steps, including (1) stereotaxically injecting the virus at the desired brain region of a mouse brain to label a specific subgroup of neurons with genetically encoded calcium sensor; (2) implantation of gradient-index (GRIN) lens that can relay calcium image from deep brain region to the miniscope system; and (3) affixing the miniscope holder over the mouse skull where miniscope can be attached later. To perform in vivo calcium imaging, the miniscope is fastened onto the holder, and neuronal calcium images are collected along with simultaneous behavior recordings. The present surgery protocol is compatible with any commercial or custom-built single-photon and two-photon imaging systems for deep brain in vivo calcium imaging.

Stereotaxic Viral Injection and Gradient-Index Lens Implantation for Deep Brain In Vivo Calcium Imaging

Miniscope in vivo calcium imaging is a powerful technique to study neuronal dynamics and microcircuits in freely behaving mice. This protocol describes performing brain surgeries to achieve good in vivo calcium imaging using a miniscope.

A miniature fluorescence microscope (miniscope) is a potent tool for in vivo calcium imaging from freely behaving animals. It offers several advantages ...

TACI: An ImageJ Plugin for 3D Calcium Imaging Analysis

Research in neuroscience has evolved to use complex imaging and computational tools to extract comprehensive information from data sets. Calcium imaging is a widely used technique that requires sophisticated software to obtain reliable results, but many laboratories struggle to adopt computational methods when updating protocols to meet modern standards. Difficulties arise due to a lack of programming knowledge and paywalls for software. In addition, cells of interest display movements in all directions during calcium imaging. Many approaches have been developed to correct the motion in the lateral (x/y) direction.
This paper describes a workflow using a new ImageJ plugin, TrackMate Analysis of Calcium Imaging (TACI), to examine motion on the z-axis in 3D calcium imaging. This software identifies the maximum fluorescence value from all the z-positions a neuron appears in and uses it to represent the neuron&#39;s intensity at the corresponding t-position. Therefore, this tool can separate neurons overlapping in the lateral (x/y) direction but appearing&#160;on distinct z-planes. As an ImageJ plugin, TACI is a user-friendly, open-source computational tool for 3D calcium imaging analysis. We validated this workflow using fly larval thermosensitive neurons that displayed movements in all directions during temperature fluctuation and a 3D calcium imaging dataset acquired from the fly brain.

TrackMate Analysis of Calcium Imaging (TACI) is an open-source ImageJ plugin for 3D calcium imaging analysis that examines motion on the z-axis and identifies the maximum value of each z-stack to represent a cell's intensity at the corresponding time point. It can separate neurons overlapping in the lateral (x/y) direction but on different z-planes.

Research in neuroscience has evolved to use complex imaging and computational tools to extract comprehensive information from data sets. Calcium imaging ...

Subcellular Imaging of Neuronal Calcium Handling In Vivo

Calcium (Ca2+) imaging has been largely used to examine neuronal activity, but it is becoming increasingly clear that subcellular Ca2+ handling is a crucial component of intracellular signaling. The visualization of subcellular Ca2+ dynamics in vivo,&#160;where neurons can be studied in their native, intact circuitry, has proven technically challenging in complex nervous systems. The transparency and relatively simple nervous system of the nematode Caenorhabditis elegans enable the cell-specific expression and in vivo visualization of fluorescent tags and indicators. Among these are fluorescent indicators that have been modified for use in the cytoplasm as well as various subcellular compartments, such as the mitochondria. This protocol enables non-ratiometric Ca2+ imaging in vivo with a subcellular resolution that permits the analysis of Ca2+ dynamics down to the level of individual dendritic spines and mitochondria. Here, two available genetically encoded indicators with different Ca2+ affinities are used to demonstrate the use of this protocol for measuring relative Ca2+ levels within the cytoplasm or mitochondrial matrix in a single pair of excitatory interneurons (AVA). Together with the genetic manipulations and longitudinal observations possible in C. elegans, this imaging protocol may be useful for answering questions regarding how Ca2+ handling regulates neuronal function and plasticity.

Subcellular Imaging of Neuronal Calcium Handling In Vivo

The current methods describe a non-ratiometric approach for high-resolution, sub-compartmental calcium imaging in vivo in Caenorhabditis elegans using readily available genetically encoded calcium indicators.

Calcium (Ca2+) imaging has been largely used to examine neuronal activity, but it is becoming increasingly clear that subcellular Ca2+ handling is a ...