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><tbody><tr><td>Transgenic Zebrafish line: Tg(ins:nls-Renilla-mKO2; cryaa:CFP); Tg(ins:GCaMP6s; cryaa:mCherry)</td><td></td><td></td><td>By request from authors</td></tr><tr><td>Stereo microscope</td><td>ZEISS</td><td>495015-0001-000</td><td>SteREO Discovery.V8</td></tr><tr><td>Fluorescence lamp</td><td>ZEISS</td><td>423013-9010-000</td><td>Illuminator HXP 120 V</td></tr><tr><td>Red Filter Cube</td><td>ZEISS</td><td>000000-1114-462&amp;nbsp;</td><td>Filter set 45 HQ TexasRed</td></tr><tr><td>Confocal Microscope</td><td>ZEISS</td><td>LSM 780</td><td></td></tr><tr><td>Bovine fibrinogen</td><td>Sigma</td><td>F8630</td><td></td></tr><tr><td>HBSS (Hanks&#039; Balanced Salt solution)</td><td>ThermoFisher</td><td>14025092</td><td></td></tr><tr><td>Thrombin</td><td>Sigma</td><td>T4648</td><td></td></tr><tr><td>D-Glucose</td><td>Sigma</td><td>G8270</td><td></td></tr><tr><td>KCl</td><td>Sigma</td><td>P9333</td><td></td></tr><tr><td>35 mm diameter glass-bottom dishes</td><td>ThermoFisher</td><td>150680</td><td></td></tr><tr><td>tricaine methane sulfonate</td><td>Sigma</td><td>E10521&amp;nbsp;</td><td></td></tr><tr><td>Fine Forceps</td><td>Fine Science Tools</td><td>11445-12</td><td></td></tr><tr><td>FIJI, using ImageJ Version: 2.0.0-rc-43/1.50e</td><td></td><td></td><td>https://fiji.sc/</td></tr><tr><td>R, version 3.2.4</td><td></td><td></td><td>https://www.r-project.org/</td></tr><tr><td>RStudio</td><td></td><td></td><td>https://www.rstudio.com/</td></tr><tr><td>plotcelltrace.R</td><td></td><td></td><td>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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/files/ftp_upload/57851/57851fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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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 ...

analysis-beta-cell-function-using-single-cell-resolution-calcium

Nanyang Technological University

Research

JoVE Journal

Biology

14.7K Views.  TU Dresden. Bèta-cel functie is belangrijk voor de homeostase van de bloedglucose, die bij eencellige resolutie met behulp van een genetisch gecodeerde verslaggever voor calcium toestroom wordt geëvalueerd.

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

Isolation and Characterization of Single Cells from Zebrafish Embryos

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been described at the morphological level, little is known about the molecular basis of cellular changes that occur as the organism develops. With recent advancements in microfluidics and multiplexing technologies, it is now possible to characterize gene expression in single cells. This allows for investigation of heterogeneity between individual cells of specific cell populations to identify and classify cell subtypes, characterize intermediate states that occur during cell differentiation, and explore differential cellular responses to stimuli. This study describes a protocol to isolate viable, single cells from zebrafish embryos for high throughput multiplexing assays. This method may be rapidly applied to any zebrafish embryonic cell type with fluorescent markers. An extension of this method may also be used in combination with high throughput sequencing technologies to fully characterize the transcriptome of single cells. As proof of principle, the relative abundance of cardiac differentiation markers was assessed in isolated, single cells derived from nkx2.5 positive cardiac progenitors. By evaluation of gene expression at the single cell level and at a single time point, the data support a model in which cardiac progenitors coexist with differentiating progeny. The method and work flow described here is broadly applicable to the zebrafish research community, requiring only a labeled transgenic fish line and access to microfluidics technologies.

This protocol describes a method for isolating single cells from zebrafish embryos, enriching for cells of interest, capturing zebrafish cells in microfluidic based single cell multiplex systems, and assessing gene expression from single cells.

The zebrafish (Danio rerio) is a powerful model organism to study vertebrate development. Though many aspects of zebrafish embryonic development have been ...

Developmental Biology

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Simultaneous Calcium Imaging and Glucose Stimulation in Living Zebrafish to Investigate In Vivo &#946;-Cell Function

The pancreatic &#946;-cells sustain systemic glucose homeostasis by producing and secreting insulin according to the blood glucose levels. Defects in &#946;-cell function are associated with hyperglycemia that can lead to diabetes. During the process of insulin secretion, &#946;-cells experience an influx of Ca2+. Thus, imaging the glucose-stimulated Ca2+ influx using genetically encoded calcium indicators (GECIs) provides an avenue to studying &#946;-cell function. Previously, studies showed that isolated zebrafish islets expressing GCaMP6s exhibit significant Ca2+ activity upon stimulation with defined glucose concentrations. However, it is paramount to study how &#946;-cells respond to glucose not in isolation, but in their native environment, where they are systemically connected, vascularized, and densely innervated. To this end, the study leveraged the optical transparency of the zebrafish larvae at early stages of development to illuminate &#946;-cell activity in vivo. Here, a detailed protocol for Ca2+ imaging and glucose stimulation to investigate &#946;-cell function in vivo is presented. This technique allows to monitor the coordinated Ca2+ dynamics in &#946;-cells with single-cell resolution. Additionally, this method can be applied to work with any injectable solution such as small molecules or peptides. Altogether, the protocol illustrates the potential of the zebrafish model to investigate islet coordination in vivo and to characterize how environmental and genetic components might affect &#946;-cell function.

Here, the study presents a protocol for calcium imaging and glucose stimulation of the pancreatic &#946;-cells of the zebrafish in vivo.

The pancreatic &#946;-cells sustain systemic glucose homeostasis by producing and secreting insulin according to the blood glucose levels. Defects in ...

In Vivo Calcium Imaging of Lateral-line Hair Cells in Larval Zebrafish

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory stimuli that mechanically deflect apical protrusions called hair bundles. Deflection opens mechanotransduction (MET) channels in hair bundles, leading to an influx of cations, including calcium. This cation influx depolarizes the cell and opens voltage-gated calcium channels located basally at the hair-cell presynapse. In mammals, hair cells are encased in bone, and it is challenging to functionally assess these activities in vivo. In contrast, larval zebrafish are transparent and possess an externally located lateral-line organ that contains hair cells. These hair cells are functionally and structurally similar to mammalian hair cells and can be functionally assessed in vivo. This article outlines a technique that utilizes a genetically encoded calcium indicator (GECI), GCaMP6s, to measure stimulus-evoked calcium signals in zebrafish lateral-line hair cells. GCaMP6s can be used, along with confocal imaging, to measure in vivo calcium signals at the apex and base of lateral-line hair cells. These signals provide a real-time, quantifiable readout of both mechanosensation- and presynapse-dependent calcium activities within these hair cells. These calcium signals also provide important functional information regarding how hair cells detect and transmit sensory stimuli. Overall, this technique generates useful data about relative changes in calcium activity in vivo. It is less well-suited for quantification of the absolute magnitude of calcium changes. This in vivo technique is sensitive to motion artifacts. A reasonable amount of practice and skill are required for proper positioning, immobilization, and stimulation of larvae. Ultimately, when properly executed, the protocol outlined in this article provides a powerful way to collect valuable information about the activity of hair-cells in their natural, fully integrated states within a live animal.

The zebrafish is a model system that has many valuable features including optical clarity, rapid external development, and, of particular importance to the field of hearing and balance, externally located sensory hair cells. This article outlines how transgenic zebrafish can be used to assay both hair-cell mechanosensation and presynaptic function in toto.

Sensory hair cells are mechanoreceptors found in the inner ear that are required for hearing and balance. Hair cells are activated in response to sensory ...

Neuroscience

In Vivo Whole-Brain Imaging of Zebrafish Larvae

In Vivo Whole-Brain Imaging of Zebrafish Larvae Using Three-Dimensional Fluorescence Microscopy

As a vertebrate model animal, larval zebrafish are widely used in neuroscience and provide a unique opportunity to monitor whole-brain activity at the cellular resolution. Here, we provide an optimized protocol for performing whole-brain imaging of larval zebrafish using three-dimensional fluorescence microscopy, including sample preparation and immobilization, sample embedding, image acquisition, and visualization after imaging. The current protocol enables in vivo imaging of the structure and neuronal activity of a larval zebrafish brain at a cellular resolution for over 1 h using confocal microscopy and custom-designed fluorescence microscopy. The critical steps in the protocol are also discussed, including sample mounting and positioning, preventing bubble formation and dust in the agarose gel, and avoiding motion in images caused by incomplete solidification of the agarose gel and paralyzation of the fish. The protocol has been validated and confirmed in multiple settings. This protocol can be easily adapted for imaging other organs of a larval zebrafish.

Author Spotlight: In Vivo Whole-Brain Imaging of Zebrafish Larvae Using Three-Dimensional Fluorescence Microscopy  

Presented here is a protocol for in vivo whole-brain imaging of larval zebrafish using three-dimensional fluorescence microscopy. The experimental procedure includes sample preparation, image acquisition, and visualization.

As a vertebrate model animal, larval zebrafish are widely used in neuroscience and provide a unique opportunity to monitor whole-brain activity at the ...

Behavior Tracking and Neural Activity Pattern in the Zebrafish Forebrain

Two-Photon Calcium Imaging of Forebrain Activity in Behaving Adult Zebrafish

Adult zebrafish (Danio rerio) exhibit a rich repertoire of behaviors for studying cognitive functions. They also have a miniature brain that can be used for measuring activities across brain regions through optical imaging methods. However, reports on the recording of brain activity in behaving adult zebrafish have been scarce. The present study describes procedures to perform two-photon calcium imaging in the dorsal forebrain of adult zebrafish. We focus on steps to restrain adult zebrafish from moving their heads, which provides stability that enables laser scanning imaging of the brain activity. The head-restrained animals can freely move their body parts and breathe without aids. The procedure aims to shorten the time of head restraint surgery, minimize brain motion, and maximize the number of neurons recorded. A setup for presenting an immersive visual environment during calcium imaging is also described here, which can be used to study neural correlates underlying visually triggered behaviors.

Author Spotlight: Advancements in Adult Zebrafish Brain Research 

Here, we present a protocol to perform two-photon calcium imaging in the dorsal forebrain of adult zebrafish.

Adult zebrafish (Danio rerio) exhibit a rich repertoire of behaviors for studying cognitive functions. They also have a miniature brain that can be used ...

Imaging Calcium Dynamics in Subpopulations of Mouse Pancreatic Islet Cells

Pancreatic islet hormones regulate blood glucose homeostasis. Changes in blood glucose induce oscillations of cytosolic calcium in pancreatic islet cells that trigger secretion of three main hormones: insulin (from &#946;-cells), glucagon (&#945;-cells) and somatostatin (&#948;-cells). &#946;-Cells, which make up the majority of islet cells and are electrically coupled to each other, respond to the glucose stimulus as one single entity. The excitability of the minor subpopulations, &#945;-cells and &#948;-cells (making up around 20% (30%) and 4% (10%) of the total rodent1 (human2) islet cell numbers, respectively) is less predictable and is therefore of special interest.
Calcium sensors are delivered into the peripheral layer of cells within the isolated islet. The islet or a group of islets is then immobilized and imaged using a fluorescence microscope. The choice of the imaging mode is between higher throughput (wide-field) and better spatial resolution (confocal). Conventionally, laser scanning confocal microscopy is used for imaging tissue, as it provides the best separation of the signal between the neighboring cells. A wide-field system can be utilized too, if the contaminating signal from the dominating population of &#946;-cells is minimized.
Once calcium dynamics in response to specific stimuli have been recorded, data are expressed in numerical form as fluorescence intensity vs. time, normalized to the initial fluorescence and baseline-corrected, to remove the effects linked to bleaching of the fluorophore. Changes in the spike frequency or partial area under the curve (pAUC) are computed vs. time, to quantify the observed effects. pAUC is more sensitive and quite robust whereas spiking frequency provides more information on the mechanism of calcium increase.
Minor cell subpopulations can be identified using functional responses to marker compounds, such as adrenaline and ghrelin, that induce changes in cytosolic calcium in a specific populations of islet cells.

Here, we present a protocol for imaging and quantifying calcium dynamics in heterogeneous cell populations, such as pancreatic islet cells. Fluorescent reporters are delivered into the peripheral layer of cells within the islet, which is then immobilized and imaged, and per-cell analysis of the dynamics of fluorescence intensity is performed.

Pancreatic islet hormones regulate blood glucose homeostasis. Changes in blood glucose induce oscillations of cytosolic calcium in pancreatic islet cells ...

In Vivo Confocal Fluorescence Imaging of Neural Activity Induced by Sensory Stimulation in Partially Restrained Larval Zebrafish

Zebrafish larvae are a promising vertebrate model system for studying the neural mechanisms of behavior. Their translucence and relatively simple neural circuitry facilitate the use of optogenetic techniques in cellular analyses of behavior. Fluorescent indicators of in vivo neural activity, such as GCaMP6s, have been widely used to study the neural activity associated with simple behaviors in larval zebrafish. Here, we present a protocol for detecting sensory-induced activity in semi-restrained zebrafish larvae using the transgenic line Tg(elav3:GCaMP6s). In particular, we use the chemical agent allyl isothiocyanate to induce a robust, reproducible fluorescent response in a brain region at the border of the hindbrain and spinal cord. We discuss the potential uses of GCaMP6s for optical monitoring of neural activity during a range of behavioral paradigms and the limitations of this technique. Our protocol outlines an accessible approach for monitoring dynamic, behavior-related in vivo neural activity in the larval zebrafish brain.

Here, we present a detailed protocol to examine neural activity in brain regions of transgenic zebrafish that express GCaMP calcium indicators using confocal microscopy.

Zebrafish larvae are a promising vertebrate model system for studying the neural mechanisms of behavior. Their translucence and relatively simple neural ...

Confocal Laser Scanning Microscopy of Calcium Dynamics in Acute Mouse Pancreatic Tissue Slices

The acute mouse pancreatic tissue slice is a unique in situ preparation with preserved intercellular communication and tissue architecture that entails significantly fewer preparation-induced changes than isolated islets, acini, ducts, or dispersed cells described in typical in vitro studies. By combining the acute pancreatic tissue slice with live-cell calcium imaging in confocal laser scanning microscopy (CLSM), calcium signals can be studied in a large number of endocrine and exocrine cells simultaneously, with a single-cell or even subcellular resolution. The sensitivity permits the detection of changes and enables the study of intercellular waves and functional connectivity as well as the study of the dependence of physiological responses of cells on their localization within the islet and paracrine relationship with other cells. Finally, from the perspective of animal welfare, recording signals from a large number of cells at a time lowers the number of animals required in experiments, contributing to the 3R-replacement, reduction, and refinement-principle.

We present the preparation of acute pancreatic tissue slices and their use in confocal laser scanning microscopy to study calcium dynamics simultaneously in a large number of live cells, over long time periods, and with high spatiotemporal resolution.

The acute mouse pancreatic tissue slice is a unique in situ preparation with preserved intercellular communication and tissue architecture that entails ...

Intracellular Signaling Events and Hormone Secretion in Human Pseudoislets

Human Pseudoislet System for Synchronous Assessment of Fluorescent Biosensor Dynamics and Hormone Secretory Profiles

The pancreatic islets of Langerhans, which are small 3D collections of specialized endocrine and supporting cells interspersed throughout the pancreas, have a central role in the control of glucose homeostasis through the secretion of insulin by beta cells, which lowers blood glucose, and glucagon by alpha cells, which raises blood glucose. Intracellular signaling pathways, including those mediated by cAMP, are key for regulated alpha and beta cell hormone secretion. The 3D islet structure, while essential for coordinated islet function, presents experimental challenges for mechanistic studies of the intracellular signaling pathways in primary human islet cells. To overcome these challenges and limitations, this protocol describes an integrated live-cell imaging and microfluidic platform using primary human pseudoislets generated from donors without diabetes that resemble native islets in their morphology, composition, and function. These pseudoislets are size-controlled through the dispersion and reaggregation process of primary human islet cells. In the dispersed state, islet cell gene expression can be manipulated; for example, biosensors such as the genetically encoded cAMP biosensor, cADDis, can be introduced. Once formed, pseudoislets expressing a genetically encoded biosensor, in combination with confocal microscopy and a microperifusion platform, allow for the synchronous assessment of fluorescent biosensor dynamics and alpha and beta cell hormone secretory profiles to provide more insight into cellular processes and function.

Author Spotlight: Investigating Islet Abnormalities and Function with a Pseudoislet Protocol 

This protocol describes a method for the synchronous acquisition and co-registration of intracellular signaling events and the secretion of insulin and glucagon by primary human pseudoislets using the adenoviral delivery of a cyclic adenosine monophosphate (cAMP) biosensor, a cAMP difference detector in situ (cADDis), and a microperifusion system.

The pancreatic islets of Langerhans, which are small 3D collections of specialized endocrine and supporting cells interspersed throughout the pancreas, ...

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

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