Cell Culture
Cells can be grown using established techniques but must be plated on #1 glass coverslips coated with a cellular adhesive (like polylysine, polyornithine or laminin) to prevent the cells from detaching or moving during imaging experiments.
Solutions
Calcium imaging experiments can be performed using a variety of physiological solutions including cell culture media. It is important, however, to make sure that the solutions are free of phenol red, which greatly increases the fluorescent background. We use Tyrodes solution, which is easily made and mimics cerebrospinal fluid, and we supplement it with 0.1% Bovine Serum Albumin. We use depolarization with 60-90 mM potassium chloride to activate voltage gated calcium channels and 1&mu;M Thapsigargin (1 mM stock in DMSO) or 2&mu;M Ionomycin (1 mM stock in DMSO) to activate store operated CRAC channels. It is often convenient to remove calcium from the extracellular solution to show that calcium elevations are due to calcium influx. When removing calcium it is necessary to maintain the total concentration of divalent cations (Mg2+ and Ca2+) constant. When substituting potassium for sodium it is necessary to maintain the osmotic balance.
Tyrodes solutions:
<table border="1">
<tbody>
<tr>
<td>&nbsp;</td>
<td>Low Potassium 2mM Ca2+Tyrodes (mM)</td>
<td>Low Potassium 0 Ca2+ Tyrodes (mM)</td>
<td>High Potassium 2mM Ca2+Tyrodes (mM)</td>
<td>High Potassium 0 Ca2+Tyrodes (mM)</td>
</tr>
<tr>
<td>NaCl</td>
<td>129</td>
<td>129</td>
<td>5</td>
<td>5</td>
</tr>
<tr>
<td>KCl</td>
<td>5</td>
<td>5</td>
<td>129</td>
<td>129</td>
</tr>
<tr>
<td>CaCl2</td>
<td>2</td>
<td>0</td>
<td>2</td>
<td>0</td>
</tr>
<tr>
<td>MgCl2</td>
<td>1</td>
<td>3</td>
<td>1</td>
<td>3</td>
</tr>
<tr>
<td>Glucose</td>
<td>30</td>
<td>30</td>
<td>30</td>
<td>30</td>
</tr>
<tr>
<td>Hepes</td>
<td>25</td>
<td>25</td>
<td>25</td>
<td>25</td>
</tr>
</tbody>
</table>
Adjust pH to 7.4 with NaOH
Loading of Fura-2 calcium dye
We load cells with acetoxy-methyl-ester Fura-2 (Fura-2 AM), which diffuses across the cell membrane and is de-esterified by cellular esterases to yield Fura-2 free acid. The exact parameters for Fura-2 loading vary widely across cell types. We recommend testing various conditions by preparing several loading solutions containing a multiple concentrations of Fura-2 raging from 1- 4 &micro;M, incubating cells in the loading solution for a variety of times from 15 minutes to 2 hours and testing the loading at room temperature and at 37 deg. A simplified protocol for cortical neurons is given below:
<ol>
<li>First, prepare the 1 mM Fura-2 AM stock by adding 50&micro;l of DMSO to a 50&micro;g vial available from Invitrogen. It is important to use dry DMSO packed under nitrogen and it is necessary to remove the DMSO with a needle by puncturing the septum to prevent hydration of the DMSO. After preparing the Fura-2 AM solution keep it in a dark dry place. Fura-2 AM in DMSO is stable at RT for 24 hours and is stable at - 20 degrees in a dry container for several months.</li>
<li>Aliquot 2 mls of culture media into a 15 ml conical tube, warm to 37 deg. and add 2&micro;l of Fura-2 AM stock to generate a 1&micro;M Fura-2 AM solution. Vortex the solution vigorously for 1 minute.</li>
<li>Transfer the loading solution to a 35 mm tissue culture dish and transfer the coverslip with the cells into the dish.</li>
<li>Incubate the neurons at 37 degrees for 30 minutes in a dark incubator. Time the incubation precisely.</li>
<li>Prepare a 35 mm dish containing 2 mls of tissue culture media without Fura-2 AM. Remove the coverslip from the loading solution and place in the new dish.</li>
<li>Mount the coverslip on the imaging chamber. Remove the coverslip from the 35 mm dish and rapidly mount onto the chamber making sure to prevent drying of the cells. We use an imaging chamber manufactured by Warner Instruments that allows a 10mm coverslip containing the cells to be mounted on the bottom and a second coverslip to be mounted on the top forming a sandwich. The two coverslips are secured with vacuum grease to the chamber and two tubes at either end of the chamber allow for perfusion of solutions through the chamber. The input line is connected to a syringe and the output line is connected to a well that is emptied by a suction line connected to a vacuum trap.</li>
</ol>
The microscope
We use an inverted Nikon Eclipse TE2000-U microscope equipped with a xenon arc lamp (Sutter Instruments), an automated stage (Ludl), an excitation filter wheel (Ludl), and a cooled charge-couple device (CCD) camera (Hamamatsu Orka II). The microscope is controlled by a Macintosh computer running Open Lab software (Improvision). Several other software packages are available for ratiometric imaging. For imaging we use 40, 60 or 100x Nikon Fluor oil immersion objectives with an NA exceeding 1.2.
Imaging Protocol
<ol>
<li>Calibrate the microscope stage.</li>
<li> Load Tyrodes solution into the input line taking care to prevent the formation of air bubbles.</li>
<li>Connect the chamber to the perfusion lines and perfuse Tyrodes solution through the chamber once again, taking care to prevent the formation of bubbles in the chamber</li>
<li>Place a drop of oil on the objective, place the chamber on the microscope stage and focus on the cells using transmitted light.</li>
<li>Examine the fluorescence of the cells using illumination at 340 and at 380 nm using the eyepieces. Resting cells should be dim at 340 and bright at 380. In general, cells should not be illuminated with UV light for more than 10 or 15 seconds and the intensity of the excitation light should be reduced with a neutral density filter to prevent phototoxicity.</li>
<li>Examine the cells using the camera and set the gain and exposure of the camera to generate an image that is close to saturation (but not saturated) when illuminated at 380 nm and well below saturation when illuminated at 340 nm. Keep the exposure below 200ms if possible. Once set, do not change the camera gain or exposure unless you plan to also alter the background, RMin and RMax (see below).</li>
<li>Collect an image at each wavelength and use the region of interest (ROI) tool to measure the intensity of the background in a variety of locations in the images for each wavelength.</li>
<li>Average the background values and enter the background values into the appropriate locations in the imaging program. The background value will be subtracted from each pixel in the field.</li>
<li>Collect a practice ratio image and adjust the Threshold values for each wavelength to generate a ratio image that includes only the cells and not the background and that is not noisy in the region close to the edges of the cells.</li>
<li>Set the minimum ratio value (RMin) to be about 10% below the lowest ratio of any cell in the field.</li>
<li>Set the RMax to be about 12 times the RMin. Do not change the RMin or RMax between experiments that you plan to compare as it will make the comparison difficult. We seldom change the RMin and Rmax values on our imaging rig.</li>
<li>Use the automated stage to find the fields that you plan to image, and record the location of each field using the software. We generally collect five fields of view during an experiment. The automated stage moves to a field, collects a ratio image and moves on to the next field.</li>
<li>Set up the time-lapse interval to collect images between 0.1 and 10 seconds apart depending on the type of signals that you expect to see.</li>
<li>Start the experiment.</li>
<li>When testing the calcium imaging system it is often useful to use stimuli like high potassium tyrodes (65 mM KCl) or ionomycin (2&micro;M) that will cause an intracellular calcium rise and calcium free extracellular solution that will (containing the calcium buffers like EGTA and BAPTA) that will reduce the calcium concentration.</li>
</ol>
Analysis
Once the experiment is complete you will want to convert the set of ratio images into time-lapse calcium measurements for individual cells or regions of interest within cells. To do this:
<ol>
<li>Use the region of interest tool (ROI) to define the areas of the image in which you want to measure calcium. It is generally useful to have at least one ROI that covers the cell body of the cell. When defining the ROIs, it is useful to play the movie of the images to verify that the cells do not move during the course of the experiment.</li>
<li>Use the software to collect time-lapse ratio measurements for each ROI in each image.</li>
<li>Import the ratio measurements into an analysis program. You will need to view the calcium traces for specific cells or ROIs, average multiple cells or ROIs and convert the ratio measurements to intracellular calcium values. We use a set of macros written in Igor Pro which allow us to do all these common analysis tasks but other programs like Excel, though less convenient can also be used. </li>
<li>The equation [Ca] = (R-Rmin)/(Rmax-R)Sf*Kd can be used to convert the Fura-2 ratio values to intracellular calcium concentrations. [Ca] is the calcium concentration, R is the Fura-2 340/380 ratio, RMin and RMax are the 340/380 ratios in the absence of calcium or in the presence of a saturating concentration of calcium respectively; and Sf*Kd is the product of the Kd of Fura-2 (approximately 120 nM at RT) and a scaling value. To measure RMin, RMax and Sf*Kd it is necessary to perform either an in vivo or an in vitro calibration. In vivo calibration requires a combination of patch clamping and calcium imaging, which can be complicated but is also very precise. In vitro calibration can be done using a home-made calibration chamber that consists of two coverslips separated by a thin coverslip spacer to generate a thin layer of solution in which to perform the calibration. The most convenient way of measuring Fura-2 ratio values as a function of calcium concentrations is to use a calibration kit that contain multiple buffered calcium solutions and Fura-2 free acid. These can be obtained from Invitrogen (Molecular Probes) and contain precise instructions for use.</li>
</ol>

<ol>
	<li>Grynkiewicz, . <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+generation+of+Ca2++indicators+with+greatly+improved+fluorescence+properties.">A new generation of Ca2+ indicators with greatly improved fluorescence properties.</a> J. Biol. Chem. 260, 3440-3450 (1985).</li><li>Lewis, . <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitogen-induced+oscillations+of+cytosolic+Ca2++and+transmembrane+Ca2++current+in+human+leukemic+T+cells.">Mitogen-induced oscillations of cytosolic Ca2+ and transmembrane Ca2+ current in human leukemic T cells.</a> Cell Regul. 1, 99-112 (1989).</li><li>Dolmetsch, . <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Signaling+between+intracellular+Ca2++stores+and+depletion-activated+Ca2++channels+generates+[Ca2+]i+oscillations+in+T+lymphocytes.">Signaling between intracellular Ca2+ stores and depletion-activated Ca2+ channels generates [Ca2+]i oscillations in T lymphocytes.</a> J Gen Physiol. 103, 365-388 (1994).</li></ol>

In this presentation we've gone through all the steps for performing calcium imaging using Fura-2 AM. We used cortical neurons as a model but calcium imaging can be used to measure intracellular calcium in real time in a variety of cells. When doing this procedure it is important to use glass coverslips instead of plastic, since plastic is often fluorescent at ultraviolet wavelengths, which makes imaging difficult. Also, it is important to pre-coat the coverslips with polyornithine and laminin in order to prevent cells from detaching. Finally, it's important to remember to avoid creating air bubbles when perfusing solutions through the imaging chamber.

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<td>New Item</td>
<td>Fura-2 AM</td>
<td>Invitrogen</td>
<td>F1221</td>
<td>Protect from light</td>
</tr>
<tr>
<td>New Item</td>
<td>DMSO</td>
<td>EM Science</td>
<td>MX1457-6</td>
<td>Keep in dry place to prevent hydration</td>
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<tr>
<td>New Item</td>
<td>Albumin from bovine serum</td>
<td>Sigma</td>
<td>A8022-100G</td>
<td>Store at 4&#176;</td>
</tr>
<tr>
<td>New Item</td>
<td>Imaging Chamber</td>
<td>Warner Instruments</td>
<td>Model RC-20H</td>
<td>&nbsp;</td>
<tr>
<td>New Item</td>
<td>Microscope</td>
<td>Nikon</td>
<td>Eclipse TE2000-U</td>
<td>&nbsp;</td>
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<td>New Item</td>
<td>Potassium chloride</td>
<td>SIGMA</td>
<td>P9333-500G</td>
<td>&nbsp;</td>
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<td>New Item</td>
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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.

Watch this Scientific Journal Video about Calcium Imaging of Cortical Neurons using Fura-2 AM at JoVE.com

Calcium Imaging of Cortical Neurons using Fura-2 AM

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 ...

calcium-imaging-of-cortical-neurons-using-fura-2-am

Nanyang Technological University

Research

JoVE Journal

Biology

60.8K Views.  Stanford University. 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.

Video: Calcium Imaging of Cortical Neurons using Fura-2 AM

Preparing E18 Cortical Rat Neurons for Compartmentalization in a Microfluidic Device

In this video, we demonstrate the preparation of E18 cortical rat neurons. E18 cortical rat neurons are obtained from E18 fetal rat cortex previously dissected and prepared. The E18 cortex is, upon dissection, immediately dissociated into individual neurons. It is possible to store E18 cortex in Hibernate E buffer containing B27 at 4&deg;C for up to a week before the dissociation is performed. However, there will be a drop in cell viability. Typically we obtain our E18 Cortex fresh. It is transported to the lab in ice cold Calcium free Magnesium free dissection buffer (CMFM). Upon arrival, trypsin is added to the cortex to a final concentration of 0.125%. The cortex is then incubated at 37&deg;C for 8 minutes. DMEM containing 10% FBS is added to the cortex to stop the reaction. The cortex is then centrifuged at 2500 rpm for 2 minutes. The supernatant is removed and 2 ml of Neural Basal Media (NBM) containing 2% B27 (vol/vol) and 0.25% Glutamax (vol/vol) is added to the cortex which is then re-suspended by pipetting up and down. Next, the cortex is triturated with previously fire polished glass pipettes, each with a successive smaller opening. After triturating, the cortex is once again centrifuged at 2500 rpm for 2 minutes. The supernatant is then removed and the cortex pellet re-suspended with 2 ml of NBM containing B27 and Glutamax. The cell suspension is then passed through a 40 um nylon cell strainer. Next the cells are counted. The neurons are now ready for loading into the neuron microfluidic device.

In this video we demonstrate the preparation of E18 Cortical Rat Neurons.

In this video, we demonstrate the preparation of E18 cortical rat neurons. E18 cortical rat neurons are obtained from E18 fetal rat cortex previously ...

In Vivo 2-Photon Calcium Imaging in Layer 2/3 of Mice

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons . In-vivo two-photon calcium imaging is the only method that allows one to record the activity of a dense neuronal population with single-cell resolution . The method consists in implanting a cranial imaging window, injecting a fluorescent calcium indicator dye that can be taken up by large numbers of neurons and finally recording the activity of neurons with time lapse calcium imaging using an in-vivo two photon microscope. Co-injection of astrocyte-specific dyes allows one to differentiate neurons from astrocytes. The technique can be performed in mice expressing fluorescent molecules in specific subpopulations of neurons to better understand the network interactions of different groups of cells.

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons . In-vivo two-photon calcium imaging is the only method that allows one to record the activity of a dense neuronal population with single-cell resolution .

To understand network dynamics of microcircuits in the neocortex, it is essential to simultaneously record the activity of a large number of neurons .  ...

In vivo Imaging of Deep Cortical Layers using a Microprism

We present a protocol for in vivo imaging of cortical tissue using a deep-brain imaging probe in the shape of a microprism.  Microprisms are 1-mm in size and have a reflective coating on the hypotenuse to allow internal reflection of excitation and emission light.  The microprism probe simultaneously images multiple cortical layers with a perspective typically seen only in slice preparations.  Images are collected with a large field-of-view (~900 &mu;m).  In addition, we provide details on the non-survival surgical procedure and microscope setup.  Representative results include images of layer V pyramidal neurons from Thy-1 YFP-H mice showing their apical dendrites extending through the superficial cortical layer and extending into tufts.  Resolution was sufficient to image dendritic spines near the soma of layer V neurons.  A tail-vein injection of fluorescent dye reveals the intricate network of blood vessels in the cortex.  Line-scanning of red blood cells (RBCs) flowing through the capillaries reveals RBC velocity and flux rates can be obtained. This novel microprism probe is an elegant, yet powerful new method of visualizing deep cellular structures and cortical function in vivo.

In vivo Imaging of Deep Cortical Layers using a Microprism

Right-angle microprisms inserted into the mouse neocortex allows for deep imaging of multiple cortical layers with a viewpoint typically found in slice. One-millimeter microprisms offer a wide field-of-view (~900 &mu;m) and spatial resolutions sufficient to resolve dendritic spines. We demonstrate layer V neuronal imaging and neocortical vascular imaging using microprisms.

We present a protocol for in vivo imaging of cortical tissue using a deep-brain imaging probe in the shape of a microprism.  Microprisms are 1-mm in size ...

Monitoring Dynamic Changes In Mitochondrial Calcium Levels During Apoptosis Using A Genetically Encoded Calcium Sensor

Dynamic changes in intracellular calcium concentration in response to various stimuli regulates many cellular processes such as proliferation, differentiation, and apoptosis1. During apoptosis, calcium accumulation in mitochondria promotes the release of pro-apoptotic factors from the mitochondria into the cytosol2. It is therefore of interest to directly measure mitochondrial calcium in living cells in situ during apoptosis. High-resolution fluorescent imaging of cells loaded with dual-excitation ratiometric and non-ratiometric synthetic calcium indicator dyes has been proven to be a reliable and versatile tool to study various aspects of intracellular calcium signaling. Measuring cytosolic calcium fluxes using these techniques is relatively straightforward. However, measuring intramitochondrial calcium levels in intact cells using synthetic calcium indicators such as rhod-2 and rhod-FF is more challenging. Synthetic indicators targeted to mitochondria have blunted responses to repetitive increases in mitochondrial calcium, and disrupt mitochondrial morphology3. Additionally, synthetic indicators tend to leak out of mitochondria over several hours which makes them unsuitable for long-term experiments. Thus, genetically encoded calcium indicators based upon green fluorescent protein (GFP)4 or aequorin5 targeted to mitochondria have greatly facilitated measurement of mitochondrial calcium dynamics. Here, we describe a simple method for real-time measurement of mitochondrial calcium fluxes in response to different stimuli. The method is based on fluorescence microscopy of 'ratiometric-pericam' which is selectively targeted to mitochondria. Ratiometric pericam is a calcium indicator based on a fusion of circularly permuted yellow fluorescent protein and calmodulin4. Binding of calcium to ratiometric pericam causes a shift of its excitation peak from 415 nm to 494 nm, while the emission spectrum, which peaks around 515 nm, remains unchanged. Ratiometric pericam binds a single calcium ion with a dissociation constant in vitro of ~1.7 &mu;M4. These properties of ratiometric pericam allow the quantification of rapid and long-term changes in mitochondrial calcium concentration. Furthermore, we describe adaptation of this methodology to a standard wide-field calcium imaging microscope with commonly available filter sets. Using two distinct agonists, the purinergic agonist ATP and apoptosis-inducing drug staurosporine, we demonstrate that this method is appropriate for monitoring changes in mitochondrial calcium concentration with a temporal resolution of seconds to hours. Furthermore, we also demonstrate that ratiometric pericam is also useful for measuring mitochondrial fission/fragmentation during apoptosis. Thus, ratiometric pericam is particularly well suited for continuous long-term measurement of mitochondrial calcium dynamics during apoptosis.

This protocol describes a method for real-time measurement of mitochondrial calcium fluxes by fluorescent imaging. The method takes advantage of a circularly permutated YFP-based dual-excitation ratiometric calcium sensor (ratiometric pericam-mt) selectively expressed in mitochondria.

Dynamic changes in intracellular calcium concentration in response to various stimuli regulates many cellular processes such as proliferation, ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

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 ...

Preparation of Pancreatic Acinar Cells for the Purpose of Calcium Imaging, Cell Injury Measurements, and Adenoviral Infection

The pancreatic acinar cell is the main parenchymal cell of the exocrine pancreas and plays a primary role in the secretion of pancreatic enzymes into the pancreatic duct. It is also the site for the initiation of pancreatitis. Here we describe how acinar cells are isolated from whole pancreas tissue and intracellular calcium signals are measured. In addition, we describe the techniques of transfecting these cells with adenoviral constructs, and subsequently measuring the leakage of lactate dehydrogenase, a marker of cell injury, during conditions that induce acinar cell injury in vitro. These techniques provide a powerful tool to characterize acinar cell physiology and pathology.

We describe a reproducible method of preparing mouse pancreatic acinar cells from a mouse for the purpose of examining acinar cell calcium signals and cellular injury with physiologically and pathologically relevant stimuli. A method for adenoviral infection of these cells is also provided.

The pancreatic acinar  cell is the main parenchymal cell of the exocrine pancreas and plays a primary  role in the secretion of pancreatic enzymes into ...

Monitoring Endoplasmic Reticulum Calcium Homeostasis Using a Gaussia Luciferase SERCaMP

The endoplasmic reticulum (ER) contains the highest level of intracellular calcium, with concentrations approximately 5,000-fold greater than cytoplasmic levels. Tight control over ER calcium is imperative for protein folding, modification and trafficking. Perturbations to ER calcium can result in the activation of the unfolded protein response, a three-prong ER stress response mechanism, and contribute to pathogenesis in a variety of diseases. The ability to monitor ER calcium alterations during disease onset and progression is important in principle, yet challenging in practice. Currently available methods for monitoring ER calcium, such as calcium-dependent fluorescent dyes and proteins, have provided insight into ER calcium dynamics in cells, however these tools are not well suited for in vivo studies. Our lab has demonstrated that a modification to the carboxy-terminus of Gaussia luciferase confers secretion of the reporter in response to ER calcium depletion. The methods for using a luciferase based, secreted ER calcium monitoring protein (SERCaMP) for in vitro and in vivo applications are described herein. This video highlights hepatic injections, pharmacological manipulation of GLuc-SERCaMP, blood collection and processing, and assay parameters for longitudinal monitoring of ER calcium.

Monitoring Endoplasmic Reticulum Calcium Homeostasis Using a Gaussia Luciferase SERCaMP

Endoplasmic reticulum calcium homeostasis is disrupted in diverse pathologies. A secreted ER calcium monitoring protein (SERCaMP) reporter can be used to detect disruptions in the ER calcium store. This protocol describes the use of a Gaussia luciferase SERCaMP to examine ER calcium homeostasis in vitro and in vivo.

The endoplasmic reticulum (ER) contains the highest level of intracellular calcium, with concentrations approximately 5,000-fold greater than cytoplasmic ...

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 ...

Concurrent EEG and Functional MRI Recording and Integration Analysis for Dynamic Cortical Activity Imaging

Electroencephalography (EEG) and functional magnetic resonance imaging (fMRI) are two of the fundamental noninvasive methods for identifying brain activity. Multimodal methods have sought to combine the high temporal resolution of EEG with the spatial precision of fMRI, but the complexity of this approach is currently in need of improvement. The protocol presented here describes the recently developed spatiotemporal fMRI-constrained EEG source imaging method, which seeks to rectify source biases and improve EEG-fMRI source localization through the dynamic recruitment of fMRI sub-regions. The process begins with the collection of multimodal data from concurrent EEG and fMRI scans, the generation of 3D cortical models, and independent EEG and fMRI processing. The processed fMRI activation maps are then split into multiple priors, according to their location and surrounding area. These are taken as priors in a two-level hierarchical Bayesian algorithm for EEG source localization. For each window of interest (defined by the operator), specific segments of the fMRI activation map will be identified as active to optimize a parameter known as model evidence. These will be used as soft constraints on the identified cortical activity, increasing the specificity of the multimodal imaging method by reducing cross-talk and avoiding erroneous activity in other conditionally active fMRI regions. The method generates cortical maps of activity and time-courses, which may be taken as final results, or used as a basis for further analyses (analyses of correlation, causation, etc.) While the method is somewhat limited by its modalities (it will not find EEG-invisible sources), it is broadly compatible with most major processing software, and is suitable for most neuroimaging studies.

An EEG-fMRI multimodal imaging method, known as the spatiotemporal fMRI-constrained EEG source imaging method, is described here. The presented method employs conditionally-active fMRI sub-maps, or priors, to guide EEG source localization in a manner that improves spatial specificity and limits erroneous results.

Calcium Imaging of Cortical Neurons using Fura-2 AM

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