Name: staining the cytoplasmic ca2+ with fluo-4/am in apple pulp
Uploaded: 2021-11-06T13:00:00.000+00:00
Duration: 8 min 5 s
Description: Watch this Scientific Journal Video about Staining the Cytoplasmic Ca2+ with Fluo-4/AM in Apple Pulp at JoVE.com

This&#160;work was supported by the Agricultural Variety Improvement Project of Shandong Province (2019LZGC007)&#160;and Fruit tree innovation team of Shandong modern agricultural industry&#160;technology system (SDAIT-06-05).

1. Protoplast extraction
<ol>
	<li>Prepare the basic solution: 20 mM CaCl2, 5 mM 2-(N-morpholino)ethanesulfonic acid, and 0.4 M D-sorbitol. 
	NOTE: The pH of the basic solution was adjusted to 5.8 with 0.1 M Tris buffer, filtered through 0.22 &#181;m water-soluble filters, and stored at 4 &#176;C.</li>
	<li>Prepare the enzymatic solution: Mix 0.3%(w/v) Macerozyme R-10 and 0.5%(w/v) cellulase R-10 with the basic solution.</li>
	<li>Add 0.5 mL of enzymatic solution into a 1.5 mL centrifuge tube. Pick a healthy and ripe apple. Then slice the pulp into 10 x 5 x 1 mm3 size (Figure 1A-1C).</li>
	<li>Place the apple fruit pulp pieces into a 1.5 mL centrifuge tube containing enzymatic solution and then close the tube (Figure 1D).</li>
	<li>Incubate the tube at 28 &#176;C for 1 h, shaking at 70 rpm/min in a shaker in the dark.</li>
	<li>Wash the pulp pieces. Aspirate all the enzymatic solution and then add 0.5 mL of the basic solution.</li>
	<li>Centrifuge at 300 x g for 2 min at room temperature.</li>
	<li>To obtain a protoplast suspension, aspirate the solution from the bottom of the centrifuge tube (Figure 1E).</li>
</ol>
2. Small-molecule calcium ion fluorescence staining
<ol>
	<li>Prepare the Fluo-4/AM loading solution with 2 mM Fluo-4/AM, 20% F-127, and 10x phosphate-buffered saline (PBS:80 mM Na2HPO4, 1.36 M NaCI, 20 mM KH2PO4, and 26 mM KCI) in a 1:1:2 ratio.</li>
	<li>Add 1 &#181;L of Fluo-4/AM loading solution to 99 &#181;L of the protoplast suspension present in the 1.5 mL centrifuge tubes. Ensure that the final concentration of the fluorescent dye is 5 &#181;M. Mix the solution and then close the tube.</li>
	<li>La3+ treatment: Prepare 100 &#181;M La3+ solution with 98 &#181;L of the protoplast suspension, 1 &#181;L of 10 mM La3+, and 1 &#181;L of Fluo-4/AM loading solution. Mix the solution and close the tube.</li>
	<li>Incubate for 30 min at 37 &#176;C in the dark.</li>
	<li>Wash the protoplasts by centrifugation at 300 x g for 2 min at room temperature. Aspirate 70 &#181;L of the solution and add 70 &#181;L of the basic solution.</li>
	<li>Incubate the protoplast suspension at 37 &#176;C for 30 min to completely de-esterify.</li>
	<li>Aspirate 15 &#181;L of the protoplast suspension and drip onto a slide.</li>
	<li>Observe under a fluorescence microscope (Supplementary Figure S1). 
	NOTE: Use a color camera with high sensitivity, i.e., 3.2 MP (2048 x 1536) CMOS sensor with 3.45 &#181;m pixel resolution.</li>
	<li>Select the GFP channel for imaging (20x). Set the brightness to 0.5. 
	&#8203;NOTE:Illumination is adjustable-intensity LED light cubes with an integrated hard-coated filter set. The excitation wavelength of Fluo-4/AM is 490 nm.</li>
</ol>
3. Protoplast viability assay
<ol>
	<li>Prepare the Fluorescein diacetate (FDA) stock solution: Dissolve FDA in acetone until the final concentration is 1 mg/mL.</li>
	<li>Prepare the FDA working solution with 1 &#181;L of the stock solution and 99 &#181;L of acetone.</li>
	<li>Add 1 &#181;L of FDA working solution to 99 &#181;L of protoplast suspension. Mix the solution by pipetting up and down and then close the tube. 
	NOTE: The final concentration of the FDA is 100 &#181;g/L.</li>
	<li>Stain at room temperature for 5 min in the dark.</li>
	<li>Prepare the slides and observe under a fluorescence microscope.</li>
	<li>Select the GFP channel for imaging (Figure 1F).</li>
</ol>
4. Image analysis
<ol>
	<li>Analyze the acquired images using image analysis and spreadsheet software (e.g., Image-Pro Plus and Excel 2010).</li>
	<li>Select two vertical diameters from the protoplasts to calculate fluorescence intensity (Supplementary Figure S2). Measure the fluorescence intensity of all protoplasts under different treatments was measured. For final processing, use photo editing software.</li>
</ol>
5. Statistical analysis
<ol>
	<li>Perform statistical analysis using statistical software (Supplementary Figure S3). Data are presented in Mean &#177; SD. The student&#39;s t-test was used to analyze the differences between the experimental groups.</li>
</ol>

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The authors declare that they have no conflicts of interest with the contents of this article.

In this study, viable protoplasts were obtained by enzymatic hydrolysis. Note that this method requires fresh apples. The present protocol allows for the rapid isolation of a large number of protoplasts from fruit pulp for use in research studies. The applicability of this method is not limited to &#39;Fuji&#39;; the protoplasts of the apple pulp of &#39;Dounan&#39; and &#39;Honey Crisp&#39; can also be extracted through the same protocol (Supplementary Figure S4). The protoplast solution after enzymolysis contains cell debris, which was somewhat improved compared to previous methods. As an essential cell material, apple pulp protoplasts can be used for cell protein expression technology, single-cell sequencing, and other research.
There are many methods regarding chemical fluorescent reagent staining in plant cells30. For example, Fluo-3/AM was loaded at a low temperature (4 &#176;C), such that it would enter the root tip cells31. Fluo-3/AM can also be loaded at a high temperature (37 &#176;C) into the pollen tube32. Fluo-4/AM has successfully loaded into the pollen tube of Pyrus pyrifolia (25 &#176;C for 15 min)18 and the root hair of Arabidopsis (4 &#176;C for 30 min)33. However, these methods are not suitable for staining apple pulp protoplasts. In this study, we successfully loaded fluorescent probes into protoplasts at a high temperature (37 &#176;C). In addition, the present method is simple, fast, accurate, and does not affect protoplast vitality. A critical step in the proposed method is to wash the stained protoplasts and incubate them for 30 min. After staining, the protoplasts should be washed. The washing will reduce the background fluorescence during observation so that the fluorescence intensity of the protoplasts can be counted more accurately. Incubate for 30 min to allow more probes to be loaded into the protoplasts. Note that it is essential to avoid light exposure during the loading.
La3+ is an important tool for studying Ca2+. It binds to the Mg2+ catalytic site on the cytoplasmic membrane Ca2+-ATPase, thereby inhibiting the steady-state turnover of Ca2+-ATPase and blocking the Ca2+ transmembrane functioning34. La3+ treatment reduced cytoplasmic Ca2+, and Fluo-4/AM could reflect this change, excluding the interference of other divalent cations and verifying the feasibility of chemical fluorescence reagents.
Although this loading method offers more advantages than other methods, it still needs to be further improved in transforming data results. Here, we estimated the relative cytoplasmic Ca2+ content by detecting the fluorescence value of protoplasts, which is qualitative data rather than quantitative. Therefore, it is particularly important to translate the staining results into specific cytoplasmic Ca2+ content in future studies, which could provide a research basis for analyzing fruit Ca2+ signaling.
In summary, the method described herein can detect cytoplasmic Ca2+ in apple pulp cells, thereby providing technical support for the related studies of fruit pulp-cell calcium.

Ca2+ plays an important role in plant signal transduction and metabolism1,2. Further, it regulates fruit quality traits3,4, including hardness, sugar content, and susceptibility to physiological disorders during storage5,6. Cytoplasmic Ca2+ plays an important role in signal transduction and regulates plant growth and development7. Disturbance of cellular calcium homeostasis can induce bitter pit in apples8, brown spot disease in pears9, and umbilical rot in tomatoes10, affecting fruit quality and causing severe economic losses3,11. Calcium imaging has sufficient spatial and temporal resolution and is an important method for observing Ca2+ dynamics in living cells12,13.
At present, there are two main methods for intracellular calcium imaging in live cells: one employs chemical small-molecular fluorescent probes14, and the other is the gene encoding sensor (GECI)15,16. Given the difficulty of establishing a stable transgenic system in fruit trees and longer fruit development, GECIS is unsuitable for fruit Ca2+ fluorescence imaging.
Small-molecule fluorescent probes such as Fluo-4/AM have a particular advantage: their AM ester form (cell-permeable acetoxymethyl ester derivative) can be readily bulk-loaded into living cells without the need for transfection, which makes it flexible, rapid, and non-cytotoxic17. Fluo-4/AM could successfully be loaded into the pollen tube of Pyrus pyrifolia18 and Petunia,19 as well as into guard cells20 and root hair of Arabidopsis21.
At present, there are few reports on the calcium fluorescence staining of pulp cells22. As an important mineral element, calcium plays a key role in the growth and quality control of tree fruits such as apples. Apple trees are globally recognized as an important economic species, and apples are considered a healthy food23. In this study, we obtained viable protoplasts from apple fruit pulp through enzymatic hydrolysis and then loaded small-molecule fluorescent reagents into the cytoplasm to detect Ca2+.

<table><tbody><tr><td>10&amp;times; phosphate-buffered saline</td><td>Solarbio</td><td>P1022</td><td>PBS (phosphate buffered solution) is a phosphate buffer solution, which can provide a relatively stable ionic environment and pH buffering capacity. It is a buffer salt solution often used in biology for molecular cloning and cell culture. The pH is 7.4.&amp;nbsp;</td></tr><tr><td>2-(N-morpholino)ethanesulfonic acid</td><td>Solarbio</td><td>M8010</td><td>Biological buffer</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Solarbio</td><td>C8370</td><td>Calcium chloride dihydrate is a white or gray chemical, mostly in granular form.</td></tr><tr><td>Cellulase R-10</td><td>Yakult Honsha</td><td>MX7352</td><td>Degrade plant cell walls.</td></tr><tr><td>D-sorbitol</td><td>Solarbio</td><td>S8090</td><td>It has good moisturizing properties, prevents drying, and prevents sugar, salt, etc. from crystallizing.</td></tr><tr><td>F-127</td><td>Thermo Fisher Scientific</td><td>P6867</td><td>Pluronic F-127 is a non-ionic, surfactant polyol (molecular weight of approximately 12500 Daltons), which has been found to be beneficial to promote the dissolution of water-insoluble dyes and other materials in physiological media.&amp;nbsp;</td></tr><tr><td>FDA</td><td>Thermo Fisher Scientific</td><td>F1303</td><td>FDA is a cell-permeant esterase substrate that can serve as a viability probe that measures both enzymatic activity, which is require to activate its fluorescence, and cell-membrane integrity, which is required for intracellular retention of their fluorescent product.&amp;nbsp;</td></tr><tr><td>Fluo-4/AM</td><td>Thermo Fisher Scientific</td><td>F14201</td><td>The green fluorescent calcium indicator Fluo-4/AM is an improved version of the calcium indicator Fluo-3/AM. The Fluo-4/AM loads faster and is brighter at the same concentration. It can be well excited with a 488 nm argon ion laser.</td></tr><tr><td>Fluorescence microscope</td><td>Thermo Fisher</td><td>EVOS Auto 2</td><td>Observe the fluorescence image.</td></tr><tr><td>Macerozyme R-10</td><td>Yakult Honsha</td><td>MX7351</td><td>Degrade plant tissue to separate single cells.</td></tr><tr><td>Tris</td><td>Solarbio</td><td>T8060</td><td>It is widely used in the preparation of buffers in biochemistry and molecular biology experiments.</td></tr></tbody></table>

staining the cytoplasmic ca2+ with fluo-4/am in apple pulp

Cytosolic Ca2+ plays a key role in plant development. Calcium imaging is the most versatile method to detect dynamic changes in Ca2+ in the cytoplasm. In this study, we obtained viable protoplasts of pulp cells by enzymatic hydrolysis. Isolated protoplasts were incubated with the small-molecule fluorescent reagent (Fluo-4/AM) for 30 min at 37 &#176;C. The fluorescent probes successfully stained cytosolic Ca2+ but did not accumulate in vacuoles. La3+, a Ca2+ channel blocker, decreased cytoplasmic fluorescence intensity. These results suggest that Fluo-4/AM can be used to detect changes in cytosolic Ca2+ in the fruit flesh. In summary, we present a method to effectively isolate protoplasts from flesh cells of the fruit and detect Ca2+ by loading a small-molecule calcium fluorescent reagent in the cytoplasm of pulp cells.

Following the protocol described above, we used the enzymatic method to obtain viable protoplasts from the pulp (Figure 1). Some protoplasts had vacuoles, while others did not. While the protoplasts exhibited no fluorescence when the Ca2+ fluorescent indicator was not loaded into them. When Fluo-4/AM was loaded into the protoplasts, the cytoplasm, but not the vacuole, became fluorescent (Figure 2). This result indicated that Fluo-4/AM successfully stained Ca2+ in the cytoplasm and that no compartmentalization was observed24. Protoplasts were stained with FDA for 5 min and showed cytoplasmic fluorescence. This indicated that high temperature (37 &#176;C) does not affect protoplast viability.
La3+, a blocking agent of the Ca2+ channel25, was added when Fluo-4/AM was loaded into protoplasts. At 100 &#181;M, La3+ decreased calcium fluorescence intensity (Figure 3).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62526/62526fig01.jpg" /> 
Figure 1: Protoplasts obtained by enzymatic hydrolysis. (A) A piece was cut from a mature apple. (B) Protoplasts were extracted from pulp pieces. (C) The pulp was cut into 10 &#215; 5 &#215; 1 mm3 pieces. (D) Pulp pieces were placed in centrifuge tubes containing 0.5 mL enzyme solution. (E) Protoplasts. Arrowheads point to the protoplast, and arrows indicate the vacuole in the protoplast. (F) Protoplasts were stained with FDA for 5 min at room temperature. (This figure has been modified from reference 22 [Horticulture research: Loading calcium fluorescent probes into protoplasts to detect calcium in the flesh tissue cells of Malus domestica]. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62526/62526fig01large.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/62526/62526fig02.jpg" /> 
Figure 2: Loading Fluo-4/AM and La3+ into the protoplasts. (A) Control: Intact protoplast without any loaded fluorescent probe. (B) Protoplasts loaded with Fluo-4/AM. (C) La3+ was added when the protoplasts were loaded with Fluo-4/AM. The final concentration of La3+ was 100 &#181;M. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62526/62526fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/62526/62526fig03.jpg" /> 
Figure 3: Statistical analysis of fluorescence intensity in the protoplasts. *** Indicate significant difference as per Student&#180;s t-test (P &#60;0.001). Vertical bars indicate &#177; SD. Each data point represents the mean of 20 protoplasts. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62526/62526fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure S1: Fluorescence microscope. (A) The overall appearance of the fluorescence microscope. (B) Settings page. Select 20x objective, GFP channel, and uniformly adjust the brightness to 0.5. (C) Fluorescence excitation region. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62526/Supplementary Fig S1.docx" target="_blank">Please click here to download this File.</a>
Supplementary Figure S2: Steps used to calculate the Ca2+ fluorescence intensity at protoplast of fruit cell. Fluorescence intensity units used in this study are based on previous reports26,27,28,29. The calculation process is as follows:&#160;(A)&#160;Open the protoplast fluorescence image in the software. Click on the Measure tool. From the dropdown menu select Profile Line. (B) Select Circle in the Line Profile window. Using this draw an ellipse at the protoplast. (C)&#160;In the Line Profile window now select File&#160;| Export data (D) Ensure that the blank spreadsheet is opened and import data by clicking Data Export. Use the &#39;average&#39; function to calculate the average fluorescence intensity. Each treatment was repeated three times with more than 20 protoplasts each. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62526/Supplementary Fig S2.docx" target="_blank">Please click here to download this File.</a>
Supplementary Figure S3: Data statistics were performed using statistical analysis software. Paste the data into the table and click Analyze for data analysis. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62526/Supplementary Fig S3.docx" target="_blank">Please click here to download this File.</a>
Supplementary Figure S4: Extraction of protoplasts from the pulp of other varieties of apples. (A) Dounan. (B) Honey Crisp. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/62526/Supplementary Fig S4.docx" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Staining the Cytoplasmic Ca2+ with Fluo-4/AM in Apple Pulp at JoVE.com

Staining the Cytoplasmic Ca2+ with Fluo-4/AM in Apple Pulp

Isolated protoplasts of apple pulp cells were loaded with a calcium fluorescent reagent to detect cytoplasmic Ca2+ concentration.

Staining the Cytoplasmic Ca2+ with Fluo-4/AM in Apple Pulp

Cytosolic Ca2+ plays a key role in plant development. Calcium imaging is the most versatile method to detect dynamic changes in Ca2+ in the cytoplasm. In ...

staining-the-cytoplasmic-ca2-with-fluo-4am-in-apple-pulp

Nanyang Technological University

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4.3K Views.  Qingdao Agricultural University. Als een belangrijk mineraal element speelt calcium een sleutelrol in de groei en de kwaliteitscontrole van boomvruchten zoals appel, het kan de hardheid, het suikergehalte, de opslagperiode en de fysiologische aandoening tijdens opslag reguleren. Calciumbeeldvorming is de meest westerse methode om dynamische veranderingen in calciumionen in het cytoplasma te detecteren. Fluorescentie-indicatoren met kleine moleculen, zoals fluo-4 /AM, hebben calciumionspecifieke selectiviteit en kunnen niet-i...

Video: Staining the Cytoplasmic Ca2+ with Fluo-4/AM in Apple Pulp

Functional Calcium Imaging in Developing Cortical Networks

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

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

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

Neuroscience

Calcium Imaging of Cortical Neurons using Fura-2 AM

Calcium imaging is a common technique that is useful for measuring calcium signals in cultured cells. Calcium imaging techniques take advantage of calcium indicator dyes, which are BAPTA-based organic molecules that change their spectral properties in response to the binding of Ca2+ ions. Calcium indicator dyes fall into two categories, ratio-metric dyes like Fura-2 and Indo-1 and single-wavelength dyes like Fluo-4. Ratio-metric dyes change either their excitation or their emission spectra in response to calcium, allowing the concentration of intracellular calcium to be determined from the ratio of fluorescence emission or excitation at distinct wavelengths.  The main advantage of using ratio-metric dyes over single wavelength probes is that the ratio signal is independent of the dye concentration, illumination intensity, and optical path length allowing the concentration of intracellular calcium to be determined independently of these artifacts. One of the most common calcium indicators is Fura-2, which has an emission peak at 505 nM and changes its excitation peak from 340 nm to 380 nm in response to calcium binding.  Here we describe the use of Fura-2 to measure intracellular calcium elevations in neurons and other excitable cells.

Calcium signals play a key role in many cellular processes including gene expression, survival and differentiation. Here we demonstrate how to perform calcium imaging using Fura-2 AM. Calcium imaging is a valuable tool to study the regulation of intracellular calcium in real time and its regulation of signaling cascades.

Calcium imaging is a common technique that is useful for measuring calcium signals in cultured cells. Calcium imaging techniques take advantage of calcium ...

Preparation of Acute Subventricular Zone Slices for Calcium Imaging

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

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

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

Examination of Synaptic Vesicle Recycling Using FM Dyes During Evoked, Spontaneous, and Miniature Synaptic Activities

Synaptic vesicles in functional nerve terminals undergo exocytosis and endocytosis. This synaptic vesicle recycling can be effectively analyzed using styryl FM dyes, which reveal membrane turnover. Conventional protocols for the use of FM dyes were designed for analyzing neurons following stimulated (evoked) synaptic activity. Recently, protocols have become available for analyzing the FM signals that accompany weaker synaptic activities, such as spontaneous or miniature synaptic events. Analysis of these small changes in FM signals requires that the imaging system is sufficiently sensitive to detect small changes in intensity, yet that artifactual changes of large amplitude are suppressed. Here we describe a protocol that can be applied to evoked, spontaneous, and miniature synaptic activities, and use cultured hippocampal neurons as an example. This protocol also incorporates a means of assessing the rate of photobleaching of FM dyes, as this is a significant source of artifacts when imaging small changes in intensity.

We describe the use of styryl FM dyes to image synaptic vesicle recycling in functional nerve terminals. This protocol can be applied not only to evoked, but also spontaneous and miniature synaptic activities. The protocol expands the variety of synaptic events that can be effectively evaluated.

Synaptic vesicles in functional nerve terminals undergo exocytosis and endocytosis. This synaptic vesicle recycling can be effectively analyzed using ...

Direct Imaging of ER Calcium with Targeted-Esterase Induced Dye Loading (TED)

Visualization of calcium dynamics is important to understand the role of calcium in cell physiology. To examine calcium dynamics, synthetic fluorescent Ca2+ indictors have become popular. Here we demonstrate TED (= targeted-esterase induced dye loading), a method to improve the release of Ca2+ indicator dyes in the ER lumen of different cell types. To date, TED was used in cell lines, glial cells, and neurons in vitro. TED bases on efficient, recombinant targeting of a high carboxylesterase activity to the ER lumen using vector-constructs that express Carboxylesterases (CES). The latest TED vectors contain a core element of CES2 fused to a red fluorescent protein, thus enabling simultaneous two-color imaging. The dynamics of free calcium in the ER are imaged in one color, while the corresponding ER structure appears in red. At the beginning of the procedure, cells are transduced with a lentivirus. Subsequently, the infected cells are seeded on coverslips to finally enable live cell imaging. Then, living cells are incubated with the acetoxymethyl ester (AM-ester) form of low-affinity Ca2+ indicators, for instance Fluo5N-AM, Mag-Fluo4-AM, or Mag-Fura2-AM. The esterase activity in the ER cleaves off hydrophobic side chains from the AM form of the Ca2+ indicator and a hydrophilic fluorescent dye/Ca2+ complex is formed and trapped in the ER lumen. After dye loading, the cells are analyzed at an inverted confocal laser scanning microscope. Cells are continuously perfused with Ringer-like solutions and the ER calcium dynamics are directly visualized by time-lapse imaging. Calcium release from the ER is identified by a decrease in fluorescence intensity in regions of interest, whereas the refilling of the ER calcium store produces an increase in fluorescence intensity. Finally, the change in fluorescent intensity over time is determined by calculation of &Delta;F/F0.

Direct Imaging of ER Calcium with Targeted-Esterase Induced Dye Loading TED

Targeted-esterase induced dye loading (TED) supports the analysis of intracellular calcium store dynamics by fluorescence imaging. The method bases on targeting of a recombinant Carboxylesterase to the endoplasmic reticulum (ER), where it improves the local unmasking of synthetic low-affinity Ca2+ indicator dyes in the ER lumen.

Visualization of calcium dynamics is important to understand the role of calcium in cell physiology. To examine calcium dynamics, synthetic fluorescent ...

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, electrical excitability and cell proliferation. The diversity and specificity of Ca2+ signaling derives from mechanisms by which Ca2+ signals are generated to act over different time and spatial scales, ranging from cell-wide oscillations and waves occurring over the periods of minutes to local transient Ca2+ microdomains (Ca2+ puffs) lasting milliseconds. Recent advances in electron multiplied CCD (EMCCD) cameras now allow for imaging of local Ca2+ signals with a 128 x 128 pixel spatial resolution at rates of &#62;500 frames sec-1 (fps). This approach is highly parallel and enables the simultaneous monitoring of hundreds of channels or puff sites in a single experiment. However, the vast amounts of data generated (ca. 1 Gb per min) render visual identification and analysis of local Ca2+ events impracticable. Here we describe and demonstrate the procedures for the acquisition, detection, and analysis of local IP3-mediated Ca2+ signals in intact mammalian cells loaded with Ca2+ indicators using both wide-field epi-fluorescence (WF) and total internal reflection fluorescence (TIRF) microscopy. Furthermore, we describe an algorithm developed within the open-source software environment Python that automates the identification and analysis of these local Ca2+ signals. The algorithm localizes sites of Ca2+ release with sub-pixel resolution; allows user review of data; and outputs time sequences of fluorescence ratio signals together with amplitude and kinetic data in an Excel-compatible table.

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Here we present techniques for imaging local IP3-mediated Ca2+ events using fluorescence microscopy in intact mammalian cells loaded with Ca2+ indicators together with an algorithm that automates identification and analysis of these events.

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, ...

Measurement of Calcium Fluctuations Within the Sarcoplasmic Reticulum of Cultured Smooth Muscle Cells Using FRET-based Confocal Imaging

Maintenance of steady-state calcium (Ca2+) levels in the sarcoplasmic reticulum (SR) of vascular smooth muscle cells (VSMCs) is vital to their overall health. A significant portion of intracellular Ca2+ content is found within the SR stores in VSMCs. As the only intracellular organelle with a close association to the surrounding extracellular space through plasma membrane-SR junctions, the SR can be considered to constitute the first line of response to any irregularity in Ca2+ transients, or stress experienced by the cell. Among its many functions, one of the most important is its role in the transmission of Ca2+-regulated signals throughout the cell to induce further cell-wide reactions downstream. The more common use of cytoplasmic Ca2+ indicators in this regard is overall insufficient for research into the highly dynamic changes to the intraluminal SR Ca2+ store that have yet to be fully characterized. Here, we provide a detailed protocol for the direct and clear measurement of luminal SR Ca2+. This tool is useful for investigation into the nuanced changes in SR Ca2+ that have significant subsequent effects on the normal function and health of the cell. Fluctuations in SR Ca2+ content specifically can provide us with a significant amount of information pertaining to cellular responses to disease or stress conditions experienced by the cell. In this method, a modified version of a SR-targeted Ca2+ indicator, D1SR, is used to detect Ca2+ fluctuations in response to the introduction of agents to cultured rat aortic smooth muscle cells (SMCs). Following incubation with the D1SR indicator, confocal fluorescence microscopy and fluorescence resonance energy transfer (FRET)-based imaging are used to directly observe changes to intraluminal SR Ca2+ levels under control conditions and with the addition of agonist agents that function to induce intracellular Ca2+ movement.

Currently, most available calcium indicators are used to quantify cytoplasmic calcium transients as indirect measures of calcium released from the sarcoplasmic reticulum in cultured smooth muscle cells. This protocol describes the use of a specific FRET-based indicator that allows direct measurement of calcium signals within the sarcoplasmic reticulum lumen.

Maintenance of steady-state calcium (Ca2+) levels in the sarcoplasmic reticulum (SR) of vascular smooth muscle cells (VSMCs) is vital to their overall ...

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ concentration. To do this, mitochondria utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester Ca2+. The influx of Ca2+ into the mitochondria stimulates three rate-limiting dehydrogenases of the citric acid cycle, increasing electron transfer through the oxidative phosphorylation (OXPHOS) complexes. This stimulation maintains &#916;&#936;m, which is temporarily dissipated as the positive calcium ions cross the mitochondrial inner membrane into the mitochondrial matrix.
We describe here a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using confocal microscopy. By permeabilizing the cells, mitochondrial Ca2+ can be measured using the fluorescent Ca2+ indicator Fluo-4, AM, with measurement of &#916;&#936;m using the fluorescent dye tetramethylrhodamine, methyl ester, perchlorate (TMRM). The benefit of this system is that there is very little spectral overlap between the fluorescent dyes, allowing accurate measurement of mitochondrial Ca2+ and &#916;&#936;m simultaneously. Using the sequential addition of Ca2+ aliquots, mitochondrial Ca2+ uptake can be monitored, and the concentration at which Ca2+ induces mitochondrial membrane permeability transition and the loss of &#916;&#936;m determined.

Mitochondria can utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester calcium (Ca2+), allowing them to shape cytosolic Ca2+ signaling within the cell. We describe a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using fluorescent dyes and confocal microscopy.

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ ...

Resorption Pit Assay: An In Vitro Technique to Quantify Calcium Resorption Activity of Mature Osteoclasts using Calcium Phosphate-Coated Culture Plates

Source: Cen, W., et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/v/64016">A Simple Pit Assay Protocol to Visualize and Quantify Osteoclastic Resorption In Vitro.</a> J. Vis. Exp. (2022)
In this video, we describe an in vitro osteoclastic resorption assay on calcium phosphate-coated cell culture plates cultured with human peripheral blood mononuclear cell-derived osteoclasts. The osteoclasts form a characteristic actin ring on the calcium phosphate-coated matrix of the cell culture plate and degrade the matrix within the actin ring region to release calcium and phosphate ions, resulting in the formation of distinct resorption pits within the degraded matrix.

Source: Cen, W., et al. A Simple Pit Assay Protocol to Visualize and Quantify Osteoclastic Resorption In Vitro. J. Vis. Exp. (2022)
In this video, we ...

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Cell-based assays

Calcium Influx Assay to Measure Mitochondrial Calcium Uptake in Cultured Cells

Source: Maxwell, J.T., et al., <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/57225">Analyses of Mitochondrial Calcium Influx in Isolated Mitochondria and Cultured Cells.</a> J. Vis. Exp. (2018).
This video describes a confocal microscopy imaging-based assay to measure mitochondrial calcium uptake in cultured cells. The assay uses the calcium-sensitive fluorescent dye Rhod-2/AM to measure the mitochondrial calcium in permeabilized cultured cells.

Source: Maxwell, J.T., et al., Analyses of Mitochondrial Calcium Influx in Isolated Mitochondria and Cultured Cells. J. Vis. Exp. (2018).
This video ...

Staining the Cytoplasmic Ca²⁺ with Fluo-4/AM in Apple Pulp

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