We thank members of the Ninov lab for comments on the manuscript, members of Center for Regenerative Therapies Dresden (CRTD) fish and microscopy facility for technical assistance. N.N. is supported by funding from the DFG-CRTD, Cluster of Excellence at TU-Dresden, German Research Foundation (DFG) and the German Center for Diabetes Research (DZD).

All the procedures including animal subjects have been approved by the Animal Welfare Act and with permission of the Landesdirektion Sachsen, Germany (AZ 24&#8211;9168.11-1/2013-14, T12/2016).
1. Preparation
NOTE: This protocol is for ex vivo imaging of zebrafish primary islet from double transgenic Tg(ins:nls-Renilla-mKO2; cryaa:CFP); Tg(ins:GCaMP6s; cryaa:mCherry)19 zebrafish. In this transgenic line, the insulin promoter (ins) drives beta-cell specific expression of two transgenes: nls-Renilla-mKO2, which marks the nucleus of beta-cells with monomeric Kusabira orange 2 (mKO2) fluorescence; and GCaMP6s15, which emits green fluorescence in response to increase in intracellular calcium levels. The beta-cell-specific expression of GCaMP6s enables studying the glucose-responsiveness of beta-cells without interference from calcium changes in surrounding cell types.
<ol>
	<li>Prepare fresh fibrinogen stock (10 mg/mL) by dissolving 10 mg of bovine fibrinogen in 1 mL of Ca2+/Mg2+-containing Hanks&#39; Balanced Salt solution (HBSS) in a 1.5 mL centrifuge tube. Vortex vigorously until the fibrinogen powder dissolves completely. Keep the solution at room temperature for at least 15 min more. 
	NOTE: The stock can be kept at room temperature for 2&#8211;3 h. Discard the stock if the solution starts to polymerize and becomes viscous.</li>
	<li>Prepare the fibrinogen working solution (3.3 mg/mL) by diluting the fibrinogen stock three-fold in HBSS. For example, mix 300 &#181;L of fibrinogen stock in 600 &#181;L of HBSS to prepare 900 &#181;L of fibrinogen working solution.</li>
	<li>Prepare thrombin solution (10 U/mL) by dissolving 10 units of thrombin in 1 mL of HBSS or phosphate buffered saline (PBS). 
	NOTE: This solution can be prepared in advance, aliquoted in 50 &#181;L parts, and frozen at -20 &#176;C.</li>
	<li>Prepare 200 mM D-glucose solution by dissolving 1.8 g of D-Glucose in 50 mL of water. Keep in 4 &#176;C for long-term storage.</li>
	<li>Prepare 300 mM KCl solution by dissolving 1.1 g of KCl in 50 mL of water. Keep in 4 &#176;C for long-term storage.</li>
	<li>Procure 35 mm diameter glass-bottom dishes for mounting the islets.</li>
	<li>For dissection and mounting of the islet, use a stereo microscope equipped with fluorescence lamp and red filter cube (TRITC: excitation: 532&#8211;554 nm, emission: 570&#8211;613 nm; or Texas-Red: excitation: 540&#8211;580 nm, emission: 592&#8211;667 nm).</li>
	<li>For imaging the calcium influx in beta-cells, use an inverted confocal microscope (Zeiss LSM 780 or similar) with 20X (0.8 NA) air objective and equipped with a plate holder for the 35 mm diameter glass-bottom dishes.</li>
	<li>Prepare a 200 mg/L solution of tricaine methane sulfonate (MS222) for euthanizing the zebrafish.</li>
	<li>Procure 90 mm Petri dishes for the zebrafish dissection.</li>
</ol>
2. Zebrafish Primary Islet Dissection and Mounting
<ol>
	<li>Euthanize a fish using prolonged incubation in MS222. For this, gently remove the fish from the fish tank using a fishing net and incubate it in a Petri dish containing the MS222 solution until the animal shows no opercular (gills) movement; typically, this takes 5 min. Transfer the fish to a Petri dish containing the HBSS solution with Ca2+/Mg2+.</li>
	<li>Under a stereo microscope equipped with a fluorescence lamp and red filter cube, dissect the skin covering the right side of the abdomen of the animal to isolate the pancreas.
	<ol>
		<li>For this, cut the skin of the zebrafish from mouth to anal fin using sharp forceps. Peel away the cut skin to expose the abdomen; this snipping exposes the internal organs. By using the red fluorescence of mKO2 expression in beta-cells, ascertain the location of the islets by visual examination under the microscope. If needed, remove the lobes of the liver, as they could cover the islet, making it difficult to find. 
		NOTE: The primary islet is located near the anterior region of the abdomen, typically on the right side.</li>
	</ol>
	</li>
	<li>Clean the primary islet by carefully removing surrounding tissue such as liver and adipocytes. Take precaution not to injure or poke the islet; after clearing of the surrounding tissue, the individual cells on the islet surface become discernable.</li>
	<li>Pipette a 30 &#181;L drop of HBSS onto the center of a glass-bottom dish. Transfer the dissected islet to this drop.</li>
	<li>Carefully wash the islet once with HBSS and once with 30 &#181;L of the fibrinogen working solution (3.3 mg/mL). Make sure to avoid drying of the islet during the washing steps because failure to do so results in cell death.</li>
	<li>Slowly and gently add 10 &#181;L of the thrombin solution (10 U/mL). Leave the islet and the dish untouched for 15&#8211;20 min. Observe that the fibrinogen-thrombin drop will become viscous and slightly opaque at this point.</li>
</ol>
3. Ex Vivo Live-imaging of GCaMP Fluorescence Intensity in Zebrafish Primary Islets
<ol>
	<li>Add 200 &#181;L of HBSS on top of the mold and place the dish carefully on the plate holder of the confocal microscope. Use a 20X, 0.8 NA, air objective for confocal imaging. Locate the islet using the brightfield option.</li>
	<li>Using the filter for red fluorescence to view the nuclear mKO2 fluorescence in beta-cells, focus on the islet. Individual nuclei should be clearly visible.</li>
	<li>Locate a clear imaging plane by manually changing the focal plane of the confocal microscope so as to move through the thickness of the islet along its z-axis. Ensure that the imaging plane contains a sufficient number (50&#8211;100) of beta-cells for imaging, and the brightness of the nuclear mKO2 fluorescence is uniform, especially in the center of islet.</li>
	<li>Set up a sequential acquisition for GCaMP6s and mKO2 fluorescence using the following settings in the &#34;Smart Setup Menu&#34;: GCaMP6s, excitation: 488 nm, emission: 500&#8211;555 nm, false-color: green (Select &#34;GFP&#34;); mKO2, excitation: 561 nm (mCherry), emission: 570&#8211;630 nm, false-color: red (Select &#34;mCherry&#34;). 
	NOTE: With this setting, the red channel will record the position of beta-cell nuclei, while the green channel will record the GCaMP fluorescence intensity.</li>
	<li>In the &#34;Acquisition Mode&#34; set the image resolution at 1,024 x 1,024 pixels, speed at 10 and averaging at 1. Initiate a continuous recording by selecting the option for &#34;Time-series&#34;, and setting the &#34;Duration&#34; for 500 cycles, with approximately 2 s acquisition time per frame. 
	NOTE: The first 50 frames of the time-series correspond to the activity of the beta-cells at 5 mM glucose concentration. This is the baseline response. A responding beta-cell will show waxing and waning in the green fluorescence intensity with time. We have observed that a few (1&#8211;5%) of the beta-cells oscillate at 5 mM glucose.</li>
	<li>Keep an eye on the imaging cycle. After the first 50 frames, increase the glucose concentration of the surrounding solution to 10 mM without stopping the recording.
	<ol>
		<li>Without perturbing the image acquisition, gently pipette 5 &#181;L of the 200 mM D-glucose solution on top of the gel holding the islet. Acquire 150 frames at 10 mM glucose. 
		NOTE: The increase in glucose concentration will increase the number of beta-cells undergoing GCaMP fluorescent oscillations in the green channel.</li>
		<li>Ensure that the nuclei of the cells remain stable during the process. If the islet is shaking extensively during acquisition and the nuclei are moving out of the focal plane, discard the sample (if needed).</li>
		<li>Wait a sufficient period of time to enable polymerization of the fibrinogen-thrombin mold in order to ensure the stability of subsequent samples.</li>
	</ol>
	</li>
	<li>At 200 frames, further increase the concentration of the solution to 20 mM by gently pipetting 10 &#181;L of the 200 mM D-glucose solution. Acquire 150 frames for the 20 mM concentration.</li>
	<li>After 350 frames, depolarize the islet using 30 mM of KCl. For this, add 20 &#181;L of the 300 mM KCl stock solution. At this stage, observe that the GCaMP6s fluorescence oscillations will stop and fix at a high intensity; beta-cells that did not respond to glucose might also display an increase in green-fluorescence intensity upon KCL addition.</li>
</ol>
4. Quantification of GCaMP Fluorescence Trace for Individual Beta-cells
NOTE: To trace and quantify the responses of individual beta-cells to varying levels of glucose, quantify the GCaMP fluorescence intensity for the entire imaging period. Quantification is performed at cellular resolution. For this, use FIJI20 to extract the intensity values of GCaMP fluorescence from the images (steps 4.1&#8211;4.6), and spreadsheet software or R21 to perform analysis (steps 4.8&#8211;4.9).
<ol>
	<li>Open the image file in FIJI using the &#34;LSM Toolbox&#34;. For this, select &#34;Plugin | LSM Toolbox | Show LSM toolbox&#34;. In the &#34;LSM Toolbox&#34;, click &#34;Open LSM&#34; and select the image file. 
	Note: For formats not supported by the &#34;LSM Toolbox&#34;, convert them first to tiff for analysis.</li>
	<li>Extract the responses of cells using the region-of-interest (ROI) tool in FIJI. Open the &#34;ROI Manager&#34; in the &#34;Analyze&#34; menu under &#34;Tools&#34;. Manually draw the ROI using the &#34;polygon selection tool&#34; located in the Toolbar.</li>
	<li>Draw the ROI in the red channel in which the beta-cell nuclei are visible. Select the ROI such that it covers an area that is larger than a cell&#8217;s nucleus in order to include some of the cell&#8217;s cytoplasm. Ensure that the ROI position is consistent between frames and adjust the position if necessary.</li>
	<li>Add the selected ROIs to the &#34;ROI manager&#34; by clicking on the &#34;Add [t]&#34; button. Select and add multiple ROIs to the ROI for obtaining data on multiple cells.</li>
	<li>Next, from the &#34;Analyze&#34; menu, select &#34;Set measurements&#34;. Select &#34;Integrated Density&#34; for specifying extraction of total fluorescence intensity within the area.</li>
	<li>Shift to the green channel containing the GCaMP fluorescence and select &#34;Multi Measure&#34; in the &#34;ROI Manager&#34;. 
	NOTE: This will provide the intensity measurements for the cells throughout the time-series.</li>
	<li>In case the ROI position needs to be adjusted due to movement of the islet, manually compile the intensity measurements in different frames. Copy and paste the values to a separate spreadsheet.</li>
	<li>Obtain the timestamps of the image frames from &#34;LSM Toolbox&#34;. Use &#34;Apply stamps | Apply t stamps | File name | Dump to textfile&#34; to obtain the timestamps. Save timestamps using the &#34;Save as&#34; option, or copy them into the spreadsheet.</li>
	<li>Upon compiling the intensity values for all cells, perform the analysis one cell at a time or automatically (e.g., using Excel Formulas or R).</li>
	<li>Analyze individual cells in two steps.
	<ol>
		<li>In the first step, calculate the fluorescence intensity above the baseline. To this end, calculate the baseline fluorescent intensity (F0) as the average of fluorescence intensities for the first 50 frames (5 mM glucose). Then subtract the baseline (F0) from the entire time-series (F &#8211; F0). 
		NOTE: A few cells can show clear GCaMP oscillations under basal glucose, which typically continue following stimulation with higher concentrations. For such cells, it is only possible to estimate F0 by taking the mean intensity of the initial frames in which the cells showed a drop in fluorescence.</li>
		<li>In the second step of the analysis, obtain the final GCaMP fluorescence intensity by normalizing the fluorescence intensity. 
		NOTE: This is performed to remove the differences between islets from different animals. Individual islets display varying levels of fluorescence emission upon glucose stimulation.</li>
		<li>Normalize the fluorescence intensity by dividing it with the highest intensity value. For this, calculate the peak intensity (Fmax &#8211; F0), and divide the baseline subtracted values by peak intensity to yield the final GCaMP fluorescence intensity (F &#8211; F0)/(Fmax &#8211; F0).</li>
	</ol>
	</li>
	<li>Discard the cells that do not exhibit a change in intensity following KCl stimulation, as they might be unhealthy or damaged.</li>
	<li>For performing the analysis (steps 4.9&#8211;4.10) in R, use the R script (plotcelltrace.R) provided with this manuscript.</li>
</ol>

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The authors declare no competing financial interests.

Here we demonstrate a technique for the quantification of beta-cell glucose responsiveness at single-cell resolution. This is made possible by monitoring intracellular calcium concentration using a genetically-encoded calcium indicator, GCaMP6s. The beta-cell activity is captured ex vivo by mounting the islet in a fibrinogen-thrombin mold. A critical step of the protocol is the stability of the mold. Sufficient time needs to be given for the fibrinogen to dissolve in the HBSS solution. Without this, the mold does not polymerize sufficiently to provide stability during the imaging session. An islet mounted in fibrinogen-thrombin mold and immersed in cell culture media can remain viable for at least a week (data not shown). Alternatives to the fibrinogen-thrombin mold, such as low-melt agarose, can be utilized to mount the islet22. Another critical parameter is the dissection of the islet. During this step, the tissue surrounding the islet needs to be removed without injuring or poking the islet. A skillful dissection comes with practice.
A limitation of the imaging protocol is the restriction to one confocal plane of the islet. This is done to capture the dynamics of calcium influx within individual beta-cells. A Z-stack through the entire thickness of the islet leads to low imaging speeds and loss of oscillatory signal from individual cells. This limitation could be improved by using faster means of confocal-imaging such as spinning-disk microscopy, to enable capturing the calcium dynamics in 3-dimensions. Another frontier would be in vivo calcium imaging12. The transparent nature of zebrafish embryo or the use of pigment-less strains of zebrafish adults23 could open the possibility for in vivo imaging in the future.
The imaging of beta-cell activity at high spatial and temporal resolution enables to investigate the functional heterogeneity among individual beta-cells. This approach can help shed light on the existence of beta-cell sub-populations. Recently, multiple studies have shown the existence of sub-populations in the nominally homogenous beta-cells24,25,26. Ex vivo imaging can be combined with genetic reporters to characterize of the sub-population&#39;s response to glucose. Additionally, combining the imaging setup with pharmacological stimulation can allow for screening of compounds that could improve beta-cell functionality.
In summary, the technique presented here allows quantification and comparison of the glucose responsiveness for individual beta-cells. It provides a direct window into beta-cell functionality, an important parameter in development of diabetes.

Our blood-glucose is maintained in a narrow range, in large part due to the function of the endocrine pancreas. The endocrine role of the pancreas is performed by the islets of Langerhans, which contain hormone-secreting cells. The insulin-secreting beta-cells are responsible for reducing the levels of blood-glucose following a carbohydrate-containing meal. Inadequate insulin secretion from beta-cell can result in diabetes1, which is characterized by sustained high blood-glucose. Type 1 and Type 2 Diabetes, which currently afflicts more than 400 million people worldwide, leads to morbidity and mortality2. By investigating the molecular and environmental factors that contribute to beta-cell dysfunction, we will understand better how type 2 diabetes starts and progresses. In addition, the ability to differentiate human stem cells into functional beta-cells in vitro can provide a source of new beta-cells for cell-replacement therapies in Type 1 Diabetes. To this end, it is important to study beta-cell function and maturation in genetic model organisms in order to gain the necessary knowledge for making functional beta-cells in a dish.
Beta-cell functionality can be monitored at the whole-islet level by quantifying the total amount of insulin secreted in response to glucose-stimulation. This cumulative approach studies the islet as a single group of cells without differentiating a cell&#39;s individual properties. However, the analysis of glucose responses of individual beta-cells revealed a diversity in the functional properties of beta-cells and the presence of heterogeneity3. To assess the function of individual beta-cells, it is possible to monitor the intracellular changes that lead to insulin secretion4. Insulin secretion is preceded by an entry of glucose into the beta-cells. The glucose that enters in the beta-cells is rapidly metabolized to ATP. Higher intracellular ATP concentrations decrease the open probability of ATP-sensitive potassium ion channels leading to beta-cell depolarization. Depolarization opens the voltage-sensitive calcium ion channels and increases intracellular calcium. In turn, calcium triggers exocytosis of insulin, which is released in the circulation and lowers glucose levels by promoting glucose-utilization5,6,7.
Several strategies have been applied to investigate the function of beta-cells, including monitoring of the membrane potential8, direct visualization of insulin-vesicles exocytosis9, and quantification of intracellular Ca2+ influx as a proxy for glucose-responsiveness10. Among them, imaging of intracellular Ca2+ provides the advantage of up-scaling the analysis to multiple individual cells within the same islet11,12, allowing for direct comparison of the glucose-responsiveness between individual beta-cells. Intracellular Ca2+ concentration can be monitored using calcium-sensitive fluorescent dyes13 or genetically-encoded calcium indicators (GECIs)14. Whereas calcium-indicator dyes lack cell-type specificity, GECIs can be expressed in a specific cell type by specific promoters. Additionally, the new generation of GECIs, such as GCaMP6, provides better signal-to-noise ratio along with faster temporal dynamics15. Here we describe the utility of the new-generation of GECIs, in particular GCaMP6s, to visualize calcium in beta-cells at single-cell resolution. We apply this method to the zebrafish primary islet as our model of choice. During embryonic development, beta-cells in the primary islet originate from the dorsal and ventral pancreatic buds16. The primary islet is located at a stereotypical anatomical position within the zebrafish pancreas, enabling its easy identification and isolation. Beta-cells in the primary islet are necessary for glucose-regulation, as their genetic ablation leads to hyperglycemia17,18. Furthermore, these beta-cells become glucose-responsive during early zebrafish development19. This protocol can be also applied to imaging the secondary islets, which form during the post-embryonic stages. The protocol allows to image beta-cells ex vivo, at later stages of development and under defined glucose concentrations.

<table border="0" cellpadding="0" cellspacing="0" width="1034">
	<tbody>
		<tr height="79">
			<td height="79" style="height:79px;width:276px;">Transgenic Zebrafish line: Tg(ins:nls-Renilla-mKO2; cryaa:CFP); Tg(ins:GCaMP6s; cryaa:mCherry)</td>
			<td style="width:123px;"></td>
			<td style="width:344px;"></td>
			<td style="width:291px;">By request from authors</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:276px;">Stereo microscope</td>
			<td style="width:123px;">ZEISS</td>
			<td style="width:344px;">495015-0001-000</td>
			<td style="width:291px;">SteREO Discovery.V8</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:276px;">Fluorescence lamp</td>
			<td style="width:123px;">ZEISS</td>
			<td style="width:344px;">423013-9010-000</td>
			<td style="width:291px;">Illuminator HXP 120 V</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:276px;">Red Filter Cube</td>
			<td style="width:123px;">ZEISS</td>
			<td style="width:344px;">000000-1114-462&#160;</td>
			<td style="width:291px;">Filter set 45 HQ TexasRed</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:276px;">Confocal Microscope</td>
			<td style="width:123px;">ZEISS</td>
			<td style="width:344px;">LSM 780</td>
			<td style="width:291px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Bovine fibrinogen</td>
			<td style="width:123px;">Sigma</td>
			<td>F8630</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">HBSS (Hanks&#39; Balanced Salt solution)</td>
			<td>ThermoFisher</td>
			<td>14025092</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Thrombin</td>
			<td>Sigma</td>
			<td>T4648</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">D-Glucose</td>
			<td>Sigma</td>
			<td>G8270</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">KCl</td>
			<td>Sigma</td>
			<td>P9333</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">35 mm diameter glass-bottom dishes</td>
			<td>ThermoFisher</td>
			<td>150680</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">tricaine methane sulfonate</td>
			<td style="width:123px;">Sigma</td>
			<td style="width:344px;">E10521&#160;</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:276px;">Fine Forceps</td>
			<td style="width:123px;">Fine Science Tools</td>
			<td style="width:344px;">11445-12</td>
			<td style="width:291px;"></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:276px;">FIJI, using ImageJ Version: 2.0.0-rc-43/1.50e</td>
			<td style="width:123px;"></td>
			<td style="width:344px;"></td>
			<td style="width:291px;">https://fiji.sc/</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:276px;">R, version 3.2.4</td>
			<td style="width:123px;"></td>
			<td style="width:344px;"></td>
			<td style="width:291px;">https://www.r-project.org/</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:276px;">RStudio</td>
			<td style="width:123px;"></td>
			<td style="width:344px;"></td>
			<td style="width:291px;">https://www.rstudio.com/</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;width:276px;">plotcelltrace.R</td>
			<td style="width:123px;"></td>
			<td style="width:344px;"></td>
			<td style="width:291px;">A custom script provided with the manuscript</td>
		</tr>
	</tbody>
</table>

analysis of beta-cell function using single-cell resolution calcium imaging in zebrafish islets

Pancreatic beta-cells respond to increasing blood glucose concentrations by secreting the hormone insulin. The dysfunction of beta-cells leads to hyperglycemia and severe, life-threatening consequences. Understanding how the beta-cells operate under physiological conditions and what genetic and environmental factors might cause their dysfunction could lead to better treatment options for diabetic patients. The ability to measure calcium levels in beta-cells serves as an important indicator of beta-cell function, as the influx of calcium ions triggers insulin release. Here we describe a protocol for monitoring the glucose-stimulated calcium influx in zebrafish beta-cells by using GCaMP6s, a genetically encoded sensor of calcium. The method allows monitoring the intracellular calcium dynamics with single-cell resolution in ex vivo mounted islets. The glucose-responsiveness of beta-cells within the same islet can be captured simultaneously under different glucose concentrations, which suggests the presence of functional heterogeneity among zebrafish beta-cells. Furthermore, the technique provides high temporal and spatial resolution, which reveals the oscillatory nature of the calcium influx upon glucose stimulation. Our approach opens the doors to use the zebrafish as a model to investigate the contribution of genetic and environmental factors to beta-cell function and dysfunction.

Using the protocol described above, the glucose responses of beta-cells in an islet from a 45 days post-fertilization (dpf) zebrafish were analyzed. For this, the primary islet was dissected from a euthanized animal and mounted in the fibrinogen-thrombin mold in a glass-bottom dish. The islet was immersed in HBSS containing 5 mM glucose. The glucose concentration was increased in a step-wise manner to 10 mM and 20 mM. The responses of beta-cells to the increasing glucose concentrations were recorded. Finally, beta-cells were depolarized using 30 mM KCl (Figure 1). The depolarization using KCl induces calcium entry in healthy beta-cells.
Using FIJI and a data analysis software, the GCaMP6s fluorescence intensity of individual beta-cells is extracted and normalized (Figure 2). As seen from the trace of fluorescence intensity, individual beta-cells display oscillations in GCaMP6s fluorescence upon glucose stimulation, which stops upon KCl stimulation. The technique provides a cellular resolution of the beta-cell&#39;s glucose-responsiveness and a window into their functionality.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57851/57851fig1.jpg" /> 
Figure 1: Ex vivo live-imaging of calcium influx using GCaMP6s in zebrafish beta-cells. A primary islet from Tg(ins:nls-Renilla-mKO2); Tg(ins:GCaMP6s) zebrafish (45 dpf) was mounted in fibrinogen-thrombin mold and incubated with 5 mM (basal) glucose. Beta-cells were labeled with a red nuclear marker, while the GCaMP6s fluorescence is present in the green channel. The islet was stimulated with a glucose-ramp consisting of sequential incubation with 10 mM and 20 mM D-glucose, and depolarized via the addition of 30 mM KCl. Arrowheads mark individual beta-cells whose activity was analyzed. <a href="https://www.jove.com/files/ftp_upload/57851/57851fig1large.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/57851/57851fig2.jpg" /> 
Figure 2: Normalized GCaMP6s fluorescence intensity-trace for individual beta-cells. Normalized GCaMP6s fluorescence intensity-trace for the beta-cells marked with arrowheads in Figure 1. The X-axis denotes the time in seconds. At the top, bars depict the concentration of glucose and KCl in the HBSS medium. The Y-axis denotes the normalized fluorescence intensity during the time-series. For this, baseline intensity (F0) is calculated as the mean intensity during incubation in 5 mM glucose. This is subtracted from the entire time-series data (F - F0). The intensity-over baseline is normalized by the maximum intensity displayed by the cell (F - F0)/(Fmax- F0). The normalized trace shows an oscillatory response of beta-cells to glucose, which stabilizes when the cells are depolarized with 30 mM KCl. <a href="https://www.jove.com/files/ftp_upload/57851/57851fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Analysis of Beta-cell Function Using Single-cell Resolution Calcium Imaging in Zebrafish Islets at JoVE.com

Analysis of Beta-cell Function Using Single-cell Resolution Calcium Imaging in Zebrafish Islets

Beta-cell functionality is important for blood-glucose homeostasis, which is evaluated at single-cell resolution using a genetically encoded reporter for calcium influx.

Pancreatic beta-cells respond to increasing blood glucose concentrations by secreting the hormone insulin. The dysfunction of beta-cells leads to ...

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14.6K Views.  TU Dresden. Beta-cell functionality is important for blood-glucose homeostasis, which is evaluated at single-cell resolution using a genetically encoded reporter for calcium influx.

Video: Analysis of Beta-cell Function Using Single-cell Resolution Calcium Imaging in Zebrafish Islets

Using the optokinetic response to study visual function of zebrafish

Optokinetic response (OKR) is a behavior that an animal vibrates its eyes to follow a rotating grating around it. It has been widely used to assess the visual functions of larval zebrafish1-5. Nevertheless, the standard protocol for larval fish is not yet readily applicable in adult zabrafish. Here, we introduce how to measure the OKR of adult zebrafish with our simple custom-built apparatus using a new protocol which is established in our lab. Both our apparatus and step-by-step procedure of OKR in adult zebrafish are illustrated in this video. In addition, the measurements of the larval OKR, as well as the optomotor response (OMR) test of adult zebrafish, are also demonstrated in this video. This OKR assay of adult zebrafish in our experiment may last for up to 4 hours. Such OKR test applied in adult fish will benefit to visual function investigation more efficiently when the adult fish vision system is manipulated.
Su-Qi Zou and Wu Yin contributed equally to this paper.

Optokinetic response has been widely used to assess the visual functions of larval zebrafish. Nevertheless, the standard protocol for larval fish is not yet readily applicable in adults1-5. Here, we introduce how to measure the OKR of adult zebrafish using a new protocol which is established in our lab.

Optokinetic response (OKR) is a behavior that an animal vibrates its eyes to follow a rotating grating around it. It has been widely used to assess the ...

Single Cell Fate Mapping in Zebrafish

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, lymphatic, and blood vessel lineages arise in developing embryos requires fate mapping and lineage tracing of undifferentiated precursor cells. Recently, photoactivatable proteins which include: Eos1, 2, PAmCherry3, Kaede4-7, pKindling8, and KikGR9, 10 have received wide interest as cell tracing probes. The fluorescence spectrum of these photosensitive proteins can be easily converted with UV excitation, allowing a population of cells to be distinguished from adjacent ones. However, the photoefficiency of the activated protein may limit long-term cell tracking11. As an alternative to photoactivatable proteins, caged fluorescein-dextran has been widely used in embryo model systems7, 12-14. Traditionally, to uncage fluorescein-dextran, UV excitation from a fluorescence lamp house or a single photon UV laser has been used; however, such sources limit the spatial resolution of photoactivation. Here we report a protocol to fate map, lineage trace, and detect single labeled cells. Single cells in embryos injected with caged fluorescein-dextran are photoactivated with near-infrared laser pulses produced from a titanium sapphire femtosecond laser. This laser is customary in all two-photon confocal microscopes such as the LSM 510 META NLO microscope used in this paper. Since biological tissue is transparent to near-infrared irradiation15, the laser pulses can be focused deep within the embryo without uncaging cells above or below the selected focal plane. Therefore, non-linear two-photon absorption is induced only at the geometric focus to uncage fluorescein-dextran in a single cell. To detect the cell containing uncaged fluorescein-dextran, we describe a simple immunohistochemistry protocol16 to rapidly visualize the activated cell. The activation and detection protocol presented in this paper is versatile and can be applied to any model system. 
Note: The reagents used in this protocol can be found in the table appended at the end of the article.

A method is described to photoactivate single cells containing a caged fluorescent protein using two-photon absorption from a Ti:Sapphire femtosecond laser oscillator. To fate map the photoactivated cell, immunohistochemistry is used. This technique can be applied to any cell type.

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, ...

High Resolution Whole Mount In Situ Hybridization within Zebrafish Embryos to Study Gene Expression and Function

This article focuses on whole-mount in situ hybridization (WISH) of zebrafish embryos. The WISH technology facilitates the assessment of gene expression both in terms of tissue distribution and developmental stage. Protocols are described for the use of WISH of zebrafish embryos using antisense RNA probes labeled with digoxigenin. Probes are generated by incorporating digoxigenin-linked nucleotides through in vitro transcription of gene templates that have been cloned and linearized. The chorions of embryos harvested at defined developmental stages are removed before incubation with specific probes. Following a washing procedure to remove excess probe, embryos are incubated with anti-digoxigenin antibody conjugated with alkaline phosphatase. By employing a chromogenic substrate for alkaline phosphatase, specific gene expression can be assessed. Depending on the level of gene expression the entire procedure can be completed within 2-3 days.

High Resolution Whole Mount In Situ Hybridization within Zebrafish Embryos to Study Gene Expression and Function

The zebrafish, a small tropical fish, has become a popular model for studying gene function during vertebrate development and disease. The temporal and spatial expression of target genes can be determined by in situ hybridization. Our improved protocol allows for the detection of low abundant transcripts with low non-specific background signal.

This article focuses on whole-mount in situ hybridization (WISH) of zebrafish embryos. The WISH technology facilitates the assessment of gene expression ...

Rapid and Efficient Zebrafish Genotyping Using PCR with High-resolution Melt Analysis

Zebrafish is a powerful vertebrate model system for studying development, modeling disease, and performing drug screening. Recently a variety of genetic tools have been introduced, including multiple strategies for inducing mutations and generating transgenic lines. However, large-scale screening is limited by traditional genotyping methods, which are time-consuming and labor-intensive. Here we describe a technique to analyze zebrafish genotypes by PCR combined with high-resolution melting analysis (HRMA).&#160;This approach is rapid, sensitive, and inexpensive, with lower risk of contamination artifacts.&#160;Genotyping by PCR with HRMA can be used for embryos or adult fish, including in high-throughput screening protocols.

PCR combined with high-resolution melt analysis (HRMA) is demonstrated as a rapid and efficient method to genotype zebrafish.

Zebrafish is a powerful vertebrate model system for studying development, modeling disease, and performing drug screening. Recently a variety of genetic ...

Analysis of Nephron Composition and Function in the Adult Zebrafish Kidney

The zebrafish model has emerged as a relevant system to study kidney development, regeneration and disease. Both the embryonic and adult zebrafish kidneys are composed of functional units known as nephrons, which are highly conserved with other vertebrates, including mammals. Research in zebrafish has recently demonstrated that two distinctive phenomena transpire after adult nephrons incur damage: first, there is robust regeneration within existing nephrons that replaces the destroyed tubule epithelial cells; second, entirely new nephrons are produced from renal progenitors in a process known as neonephrogenesis. In contrast, humans and other mammals seem to have only a limited ability for nephron epithelial regeneration. To date, the mechanisms responsible for these kidney regeneration phenomena remain poorly understood. Since adult zebrafish kidneys undergo both nephron epithelial regeneration and neonephrogenesis, they provide an outstanding experimental paradigm to study these events. Further, there is a wide range of genetic and pharmacological tools available in the zebrafish model that can be used to delineate the cellular and molecular mechanisms that regulate renal regeneration. One essential aspect of such research is the evaluation of nephron structure and function. This protocol describes a set of labeling techniques that can be used to gauge renal composition and test nephron functionality in the adult zebrafish kidney. Thus, these methods are widely applicable to the future phenotypic characterization of adult zebrafish kidney injury paradigms, which include but are not limited to, nephrotoxicant exposure regimes or genetic methods of targeted cell death such as the nitroreductase mediated cell ablation technique. Further, these methods could be used to study genetic perturbations in adult kidney formation and could also be applied to assess renal status during chronic disease modeling.

The zebrafish adult kidney is an excellent system for renal regeneration and disease studies. An essential aspect of such research is the assessment of nephron structure and function. This protocol describes several methodologies that can be implemented to assess nephron tubule composition and to evaluate renal reabsorption.

The zebrafish model has emerged as a relevant system to study kidney development, regeneration and disease. Both the embryonic and adult zebrafish kidneys ...

Evaluation of Zebrafish Kidney Function Using a Fluorescent Clearance Assay

The zebrafish embryo offers a tractable model to study organogenesis and model human genetic disease. Despite its relative simplicity, the zebrafish kidney develops and functions in almost the same way as humans. A major difference in the construction of the human kidney is the presence of millions of nephrons compared to the zebrafish that has only two. However, simplifying such a complex system into basic functional units has aided our understanding of how the kidney develops and operates. In zebrafish, the midline located glomerulus is responsible for the initial blood filtration into two pronephric tubules that diverge to run bilaterally down the embryonic axis before fusing to each other at the cloaca. The pronephric tubules are heavily populated by motile cilia that facilitate the movement of filtrate along the segmented tubule, allowing the exchange of various solutes before finally exiting via the cloaca2-4. Many genes responsible for CKD, including those related to ciliogenesis, have been studied in zebrafish5. However, a major draw back has been the difficulty in evaluating zebrafish kidney function after genetic manipulation. Traditional assays to measure kidney dysfunction in humans have proved non translational to zebrafish, mainly due to their aquatic environment and small size. For example, it is not physically possible to extract blood from embryonic staged fish for analysis of urea and creatinine content, as they are too small. In addition, zebrafish do not produce enough urine for testing on a simple proteinuria &#8216;dipstick&#8217;, which is often performed during initial patient examinations. We describe a fluorescent assay that utilizes the optical transparency of the zebrafish to quantitatively monitor the clearance of a fluorescent dye, over time, from the vasculature and out through the kidney, to give a read out of renal function1,6-9.

The zebrafish is a popular tool to model chronic kidney disease (CKD). However, their small size makes it impossible to evaluate renal function using traditional methods. We describe a fluorescent dye kidney clearance assay1 that allows quantitative analysis of zebrafish kidney function in CKD.

The zebrafish embryo offers a tractable model to study organogenesis and model human genetic disease. Despite its relative simplicity, the zebrafish ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

CUBIC Protocol Visualizes Protein Expression at Single Cell Resolution in Whole Mount Skin Preparations

The skin is essential for our survival. The outer epidermal layer consists of the interfollicular epidermis, which is a stratified squamous epithelium covering most of our body, and epidermal appendages such as the hair follicles and sweat glands. The epidermis undergoes regeneration throughout life and in response to injury. This is enabled by K14-expressing basal epidermal stem/progenitor cell populations that are tightly regulated by multiple regulatory mechanisms active within the epidermis and between epidermis and dermis. This article describes a simple method to clarify full thickness mouse skin biopsies, and visualize K14 protein expression patterns, Ki67 labeled proliferating cells, Nile Red labeled sebocytes, and DAPI nuclear labeling at single cell resolution in 3D. This method enables accurate assessment and quantification of skin anatomy and pathology, and of abnormal epidermal phenotypes in genetically modified mouse lines. The CUBIC protocol is the best method available to date to investigate molecular and cellular interactions in full thickness skin biopsies at single cell resolution.

This report describes a CUBIC protocol to clarify full thickness mouse skin biopsies, and visualize protein expression patterns, proliferating cells, and sebocytes at the single cell resolution in 3D. This method enables accurate assessment of skin anatomy and pathology, and of abnormal epidermal phenotypes in genetically modified mouse lines.

The skin is essential for our survival. The outer epidermal layer consists of the interfollicular epidermis, which is a stratified squamous epithelium ...

A Rapid Automated Protocol for Muscle Fiber Population Analysis in Rat Muscle Cross Sections Using Myosin Heavy Chain Immunohistochemistry

Quantification of muscle fiber populations provides a deeper insight into the effects of disease, trauma, and various other influences on skeletal muscle composition. Various time-consuming methods have traditionally been used to study fiber populations in many fields of research. However, recently developed immunohistochemical methods based on myosin heavy chain protein expression provide a quick alternative to identify multiple fiber types in a single section. Here, we present a rapid, reliable and reproducible protocol for improved staining quality, allowing automatic acquisition of whole cross sections and automatic quantification of fiber populations with ImageJ. For this purpose, embedded skeletal muscles are cut in cross sections, stained using myosin heavy chains antibodies with secondary fluorescent antibodies and DAPI for cell nuclei staining. Whole cross sections are then scanned automatically using a slide scanner to obtain high-resolution composite pictures of the entire specimen. Fiber population analyses are subsequently performed to quantify slow, intermediate and fast fibers using an automated macro for ImageJ. We have previously shown that this method can identify fiber populations reliably to a degree of &#177;4%. In addition, this method reduces inter-user variability and time per analyses significantly using the open source platform ImageJ.

Here, we present a protocol for rapid muscle fiber analyses, which allows improved staining quality, and thereby automatic acquisition and quantification of fiber populations using the freely available software ImageJ.

Quantification of muscle fiber populations provides a deeper insight into the effects of disease, trauma, and various other influences on skeletal muscle ...

Single-Cell Resolution Three-Dimensional Imaging of Intact Organoids

Organoid technology, in vitro 3D culturing of miniature tissue, has opened a new experimental window for cellular processes that govern organ development and function as well as disease. Fluorescence microscopy has played a major role in characterizing their cellular composition in detail and demonstrating their similarity to the tissue they originate from. In this article, we present a comprehensive protocol for high-resolution 3D imaging of whole organoids upon immunofluorescent labeling. This method is widely applicable for imaging of organoids differing in origin, size and shape. Thus far we have applied the method to airway, colon, kidney, and liver organoids derived from healthy human tissue, as well as human breast tumor organoids and mouse mammary gland organoids. We use an optical clearing agent, FUnGI, which enables the acquisition of whole 3D organoids with the opportunity for single-cell quantification of markers. This three-day protocol from organoid harvesting to image analysis is optimized for 3D imaging using confocal microscopy.

The entire 3D structure and cellular content of organoids, as well as their phenotypic resemblance to the original tissue can be captured using the single-cell resolution 3D imaging protocol described here. This protocol can be applied to a wide range of organoids varying in origin, size and shape.

Organoid technology, in vitro 3D culturing of miniature tissue, has opened a new experimental window for cellular processes that govern organ development ...