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 ΔF/F0.
21 Related JoVE Articles!
Cytosolic Calcium Measurements in Renal Epithelial Cells by Flow Cytometry
Institutions: University of Witten/Herdecke, University of Witten/Herdecke.
A variety of cellular processes, both physiological and pathophysiological, require or are governed by calcium, including exocytosis, mitochondrial function, cell death, cell metabolism and cell migration to name but a few. Cytosolic calcium is normally maintained at low nanomolar concentrations; rather it is found in high micromolar to millimolar concentrations in the endoplasmic reticulum, mitochondrial matrix and the extracellular compartment. Upon stimulation, a transient increase in cytosolic calcium serves to signal downstream events. Detecting changes in cytosolic calcium is normally performed using a live cell imaging set up with calcium binding dyes that exhibit either an increase in fluorescence intensity or a shift in the emission wavelength upon calcium binding. However, a live cell imaging set up is not freely accessible to all researchers. Alternative detection methods have been optimized for immunological cells with flow cytometry and for non-immunological adherent cells with a fluorescence microplate reader. Here, we describe an optimized, simple method for detecting changes in epithelial cells with flow cytometry using a single wavelength calcium binding dye. Adherent renal proximal tubule epithelial cells, which are normally difficult to load with dyes, were loaded with a fluorescent cell permeable calcium binding dye in the presence of probenecid, brought into suspension and calcium signals were monitored before and after addition of thapsigargin, tunicamycin and ionomycin.
Cellular Biology, Issue 92, Kidney, FACS, second messenger, proximal tubule, calcium indicators, probenecid, endoplasmic reticulum, ionomycin
Whole-Cell Recording of Calcium Release-Activated Calcium (CRAC) Currents in Human T Lymphocytes
Institutions: University of California, Davis.
In T lymphocytes, depletion of Ca2+
from the intracellular Ca2+
store leads to activation of plasmalemmal Ca2+
channels, called Calcium Release-Activated Calcium (CRAC) channels. CRAC channels play important role in regulation of T cell proliferation and gene expression. Abnormal CRAC channel function in T cells has been linked to severe combined immunodeficiency and autoimmune diseases 1, 2
. Studying CRAC channel function in human T cells may uncover new molecular mechanisms regulating normal immune responses and unravel the causes of related human diseases. Electrophysiological recordings of membrane currents provide the most accurate assessment of functional channel properties and their regulation. Electrophysiological assessment of CRAC channel currents in Jurkat T cells, a human leukemia T cell line, was first performed more than 20 years ago 3
, however, CRAC current measurements in normal human T cells remains a challenging task. The difficulties in recording CRAC channel currents in normal T cells are compounded by the fact that blood-derived T lymphocytes are much smaller in size than Jurkat T cells and, therefore, the endogenous whole-cell CRAC currents are very low in amplitude. Here, we give a step-by-step procedure that we routinely use to record the Ca2+
currents via CRAC channels in resting human T cells isolated from the peripheral blood of healthy volunteers. The method described here was adopted from the procedures used for recording the CRAC currents in Jurkat T cells and activated human T cells 4-8
Immunology, Issue 46, human T lymphocytes, CRAC channels, CRAC currents, patch-clamp
Measuring the Induced Membrane Voltage with Di-8-ANEPPS
Institutions: University of Ljubljana.
Placement of a cell into an external electric field causes a local charge redistribution inside and outside of the cell in the vicinity of the cell membrane, resulting in a voltage across the membrane. This voltage, termed the induced membrane voltage (also induced transmembrane voltage, or induced transmembrane potential difference) and denoted by ΔΦ, exists only as long as the external field is present. If the resting voltage is present on the membrane, the induced voltage superimposes (adds) onto it. By using one of the potentiometric fluorescent dyes, such as di-8-ANEPPS, it is possible to observe the variations of ΔΦ on the cell membrane and to measure its value noninvasively. di-8-ANEPPS becomes strongly fluorescent when bound to the lipid bilayer of the cell membrane, with the change of the fluorescence intensity proportional to the change of ΔΦ. This video shows the protocol for measuring ΔΦ using di-8-ANEPPS and also demonstrates the influence of cell shape on the amplitude and spatial distribution of ΔΦ.
Cellular Biology, Issue 33, induced transmembrane voltage, induced membrane voltage, induced transmembrane potential, potentiometric dyes, di-8-ANEPPS, electroporation, electropermeabilization
Cell-based Calcium Assay for Medium to High Throughput Screening of TRP Channel Functions using FlexStation 3
Institutions: The University of Texas Health Science Center at Houston.
The Molecular Devices' FlexStation 3 is a benchtop multi-mode microplate reader capable of automated fluorescence measurement in multi-well plates. It is ideal for medium- to high-throughput screens in academic settings. It has an integrated fluid transfer module equipped with a multi-channel pipetter and the machine reads one column at a time to monitor fluorescence changes of a variety of fluorescent reagents. For example, FlexStation 3 has been used to study the function of Ca2+
-permeable ion channels and G-protein coupled receptors by measuring the changes of intracellular free Ca2+
levels. Transient receptor potential (TRP) channels are a large family of nonselective cation channels that play important roles in many physiological and pathophysiological functions. Most of the TRP channels are calcium permeable and induce calcium influx upon activation. In this video, we demonstrate the application of FlexStation 3 to study the pharmacological profile of the TRPA1 channel, a molecular sensor for numerous noxious stimuli. HEK293 cells transiently or stably expressing human TRPA1 channels, grown in 96-well plates, are loaded with a Ca2+
-sensitive fluorescent dye, Fluo-4, and real-time fluorescence changes in these cells are measured before and during the application of a TRPA1 agonist using the FLEX mode of the FlexStation 3. The effect of a putative TRPA1 antagonist was also examined. Data are transferred from the SoftMax Pro software to construct concentration-response relationships of TRPA1 activators and inhibitors.
Bioengineering, Issue 54, TRP channels, Calcium assay, FlexStation 3
Isolation of Human Atrial Myocytes for Simultaneous Measurements of Ca2+ Transients and Membrane Currents
Institutions: University of Duisburg-Essen , University of Heidelberg .
The study of electrophysiological properties of cardiac ion channels with the patch-clamp technique and the exploration of cardiac cellular Ca2+
handling abnormalities requires isolated cardiomyocytes. In addition, the possibility to investigate myocytes from patients using these techniques is an invaluable requirement to elucidate the molecular basis of cardiac diseases such as atrial fibrillation (AF).1
Here we describe a method for isolation of human atrial myocytes which are suitable for both patch-clamp studies and simultaneous measurements of intracellular Ca2+
concentrations. First, right atrial appendages obtained from patients undergoing open heart surgery are chopped into small tissue chunks ("chunk method") and washed in Ca2+
-free solution. Then the tissue chunks are digested in collagenase and protease containing solutions with 20 μM Ca2+
. Thereafter, the isolated myocytes are harvested by filtration and centrifugation of the tissue suspension. Finally, the Ca2+
concentration in the cell storage solution is adjusted stepwise to 0.2 mM. We briefly discuss the meaning of Ca2+
buffering during the isolation process and also provide representative recordings of action potentials and membrane currents, both together with simultaneous Ca2+
transient measurements, performed in these isolated myocytes.
Cellular Biology, Issue 77, Medicine, Molecular Biology, Physiology, Anatomy, Cardiology, Pharmacology, human atrial myocytes, cell isolation, collagenase, calcium transient, calcium current, patch-clamp, ion currents, isolation, cell culture, myocytes, cardiomyocytes, electrophysiology, patch clamp
Simultaneous pH Measurement in Endocytic and Cytosolic Compartments in Living Cells using Confocal Microscopy
Institutions: Université de Sherbrooke, Centre de Recherche Clinique Etienne-Le Bel.
Intracellular pH is tightly regulated and differences in pH between the cytoplasm and organelles have been reported1
. Regulation of cellular pH is crucial for homeostatic control of physiological processes that include: protein, DNA and RNA synthesis, vesicular trafficking, cell growth and cell division. Alterations in cellular pH homeostasis can lead to detrimental functional changes and promote progression of various diseases2
. Various methods are available for measuring intracellular pH but very few of these allow simultaneous measurement of pH in the cytoplasm and in organelles. Here, we describe in detail a rapid and accurate method for the simultaneous measurement of cytoplasmic and organellar pH by using confocal microscopy on living cells3
. This goal is achieved with the use of two pH-sensing ratiometric dyes that possess selective cellular compartment partitioning. For instance, SNARF-1 is compartmentalized inside the cytoplasm whereas HPTS is compartmentalized inside endosomal/lysosomal organelles. Although HPTS is commonly used as a cytoplasmic pH indicator, this dye can specifically label vesicles along the endosomal-lysosomal pathway after being taken up by pinocytosis3,4
. Using these pH-sensing probes, it is possible to simultaneously measure pH within the endocytic and cytoplasmic compartments. The optimal excitation wavelength of HPTS varies depending on the pH while for SNARF-1, it is the optimal emission wavelength that varies. Following loading with SNARF-1 and HPTS, cells are cultured in different pH-calibrated solutions to construct a pH standard curve for each probe. Cell imaging by confocal microscopy allows elimination of artifacts and background noise. Because of the spectral properties of HPTS, this probe is better suited for measurement of the mildly acidic endosomal compartment or to demonstrate alkalinization of the endosomal/lysosomal organelles. This method simplifies data analysis, improves accuracy of pH measurements and can be used to address fundamental questions related to pH modulation during cell responses to external challenges.
Biochemistry, Issue 86, Confocal microscopy, pH measurement, live cell imaging, ratiometric pH probes, fluorescence, intravesicular pH, cytosolic pH, endosomes, lysosomes
Measuring Intracellular Ca2+ Changes in Human Sperm using Four Techniques: Conventional Fluorometry, Stopped Flow Fluorometry, Flow Cytometry and Single Cell Imaging
Institutions: Instituto de Biotecnología-Universidad Nacional Autónoma de México, Edison State College.
Spermatozoa are male reproductive cells especially designed to reach, recognize and fuse with the egg. To perform these tasks, sperm cells must be prepared to face a constantly changing environment and to overcome several physical barriers. Being in essence transcriptionally and translationally silent, these motile cells rely profoundly on diverse signaling mechanisms to orient themselves and swim in a directed fashion, and to contend with challenging environmental conditions during their journey to find the egg. In particular, Ca2+
-mediated signaling is pivotal for several sperm functions: activation of motility, capacitation (a complex process that prepares sperm for the acrosome reaction) and the acrosome reaction (an exocytotic event that allows sperm-egg fusion). The use of fluorescent dyes to track intracellular fluctuations of this ion is of remarkable importance due to their ease of application, sensitivity, and versatility of detection. Using one single dye-loading protocol we utilize four different fluorometric techniques to monitor sperm Ca2+
dynamics. Each technique provides distinct information that enables spatial and/or temporal resolution, generating data both at single cell and cell population levels.
Cellular Biology, Issue 75, Medicine, Molecular Biology, Genetics, Biophysics, Anatomy, Physiology, Spermatozoa, Ion Channels, Cell Physiological Processes, Calcium Signaling, Reproductive Physiological Processes, fluorometry, Flow cytometry, stopped flow fluorometry, single-cell imaging, human sperm, sperm physiology, intracellular Ca2+, Ca2+ signaling, Ca2+ imaging, fluorescent dyes, imaging
Measurement of Vacuolar and Cytosolic pH In Vivo in Yeast Cell Suspensions
Institutions: SUNY Upstate Medical University.
Vacuolar and cytosolic pH are highly regulated in yeast cells and occupy a central role in overall pH homeostasis. We describe protocols for ratiometric measurement of pH in vivo
using pH-sensitive fluorophores localized to the vacuole or cytosol. Vacuolar pH is measured using BCECF, which localizes to the vacuole in yeast when introduced into cells in its acetoxymethyl ester form. Cytosolic pH is measured with a pH-sensitive GFP expressed under control of a yeast promoter, yeast pHluorin. Methods for measurement of fluorescence ratios in yeast cell suspensions in a fluorimeter are described. Through these protocols, single time point measurements of pH under different conditions or in different yeast mutants have been compared and changes in pH over time have been monitored. These methods have also been adapted to a fluorescence plate reader format for high-throughput experiments. Advantages of ratiometric pH measurements over other approaches currently in use, potential experimental problems and solutions, and prospects for future use of these techniques are also described.
Molecular Biology, Issue 74, Biochemistry, Microbiology, Cellular Biology, Biophysics, Physiology, Proteins, Vacuoles, Cytosol, Yeasts, Membrane Transport Proteins, Ion Pumps, Fluorometry, yeast, intracellular pH, vacuole, fluorescence, ratiometric, cells
Mitochondria-associated ER Membranes (MAMs) and Glycosphingolipid Enriched Microdomains (GEMs): Isolation from Mouse Brain
Institutions: St Jude Children's Research Hospital.
Intracellular organelles are highly dynamic structures with varying shape and composition, which are subjected to cell-specific intrinsic and extrinsic cues. Their membranes are often juxtaposed at defined contact sites, which become hubs for the exchange of signaling molecules and membrane components1,2,3,4
. The inter-organellar membrane microdomains that are formed between the endoplasmic reticulum (ER) and the mitochondria at the opening of the IP3-sensitive Ca2+
channel are known as the mitochondria associated-ER membranes or MAMs4,5,6
. The protein/lipid composition and biochemical properties of these membrane contact sites have been extensively studied particularly in relation to their role in regulating intracellular Ca2+ 4,5,6
. The ER serves as the primary store of intracellular Ca2+
, and in this capacity regulates a myriad of cellular processes downstream of Ca2+
signaling, including post-translational protein folding and protein maturation7. Mitochondria, on the other hand, maintain Ca2+
homeostasis, by buffering cytosolic Ca2+
concentration thereby preventing the initiation of apoptotic pathways downstream of Ca2+
. The dynamic nature of the MAMs makes them ideal sites to dissect basic cellular mechanisms, including Ca2+
signaling and regulation of mitochondrial Ca2+
concentration, lipid biosynthesis and transport, energy metabolism and cell survival 4,9,10,11,12
. Several protocols have been described for the purification of these microdomains from liver tissue and cultured cells13,14
Taking previously published methods into account, we have adapted a protocol for the isolation of mitochondria and MAMs from the adult mouse brain. To this procedure we have added an extra purification step, namely a Triton X100 extraction, which enables the isolation of the glycosphingolipid enriched microdomain (GEM) fraction of the MAMs. These GEM preparations share several protein components with caveolae and lipid rafts, derived from the plasma membrane or other intracellular membranes, and are proposed to function as gathering points for the clustering of receptor proteins and for protein–protein interactions4,15
Neuroscience, Issue 73, Genetics, Cellular Biology, Molecular Biology, Biochemistry, Membrane Microdomains, Endoplasmic Reticulum, Mitochondria, Intracellular Membranes, Glycosphingolipids, Gangliosides, Endoplasmic Reticulum Stress, Cell Biology, Neurosciences, MAMs, GEMs, Mitochondria, ER, membrane microdomains, subcellular fractionation, lipids, brain, mouse, isolation, animal model
Examination of Synaptic Vesicle Recycling Using FM Dyes During Evoked, Spontaneous, and Miniature Synaptic Activities
Institutions: University of Iowa Carver College of Medicine, University of Bath.
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.
Neuroscience, Issue 85, Presynaptic Terminals, Synaptic Vesicles, Microscopy, Biological Assay, Nervous System, Endocytosis, exocytosis, fluorescence imaging, FM dye, neuron, photobleaching
A Microplate Assay to Assess Chemical Effects on RBL-2H3 Mast Cell Degranulation: Effects of Triclosan without Use of an Organic Solvent
Institutions: University of Maine, Orono, University of Maine, Orono.
Mast cells play important roles in allergic disease and immune defense against parasites. Once activated (e.g.
by an allergen), they degranulate, a process that results in the exocytosis of allergic mediators. Modulation of mast cell degranulation by drugs and toxicants may have positive or adverse effects on human health. Mast cell function has been dissected in detail with the use of rat basophilic leukemia mast cells (RBL-2H3), a widely accepted model of human mucosal mast cells3-5
. Mast cell granule component and the allergic mediator β-hexosaminidase, which is released linearly in tandem with histamine from mast cells6
, can easily and reliably be measured through reaction with a fluorogenic substrate, yielding measurable fluorescence intensity in a microplate assay that is amenable to high-throughput studies1
. Originally published by Naal et al.1
, we have adapted this degranulation assay for the screening of drugs and toxicants and demonstrate its use here.
Triclosan is a broad-spectrum antibacterial agent that is present in many consumer products and has been found to be a therapeutic aid in human allergic skin disease7-11
, although the mechanism for this effect is unknown. Here we demonstrate an assay for the effect of triclosan on mast cell degranulation. We recently showed that triclosan strongly affects mast cell function2
. In an effort to avoid use of an organic solvent, triclosan is dissolved directly into aqueous buffer with heat and stirring, and resultant concentration is confirmed using UV-Vis spectrophotometry (using ε280
= 4,200 L/M/cm)12
. This protocol has the potential to be used with a variety of chemicals to determine their effects on mast cell degranulation, and more broadly, their allergic potential.
Immunology, Issue 81, mast cell, basophil, degranulation, RBL-2H3, triclosan, irgasan, antibacterial, β-hexosaminidase, allergy, Asthma, toxicants, ionophore, antigen, fluorescence, microplate, UV-Vis
Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber
Institutions: Robert Wood Johnson Medical School , Robert Wood Johnson Medical School , University of Missouri-Kansas City.
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+
. 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
(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
Cellular Biology, Issue 60, Mn quenching, 2-APB, Fura-2, Orai1, esophageal squamous cell carcinoma, skinned muscle fiber
Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles
Institutions: UMDNJ-Robert Wood Johnson Medical School, University of Missouri-Kansas City, Ohio State University .
Described here is a method to measure contractility of isolated skeletal muscles. Parameters such as muscle force, muscle power, contractile kinetics, fatigability, and recovery after fatigue can be obtained to assess specific aspects of the excitation-contraction coupling (ECC) process such as excitability, contractile machinery and Ca2+
handling ability. This method removes the nerve and blood supply and focuses on the isolated skeletal muscle itself. We routinely use this method to identify genetic components that alter the contractile property of skeletal muscle though modulating Ca2+
signaling pathways. Here, we describe a newly identified skeletal muscle phenotype, i.e.
, mechanic alternans, as an example of the various and rich information that can be obtained using the in vitro
muscle contractility assay. Combination of this assay with single cell assays, genetic approaches and biochemistry assays can provide important insights into the mechanisms of ECC in skeletal muscle.
Physiology, Issue 69, extensor digitorum longus, soleus, in vitro contractility, calcium signaling, muscle-tendon complex, mechanic alternans
Detection of Toxin Translocation into the Host Cytosol by Surface Plasmon Resonance
Institutions: University of Central Florida.
AB toxins consist of an enzymatic A subunit and a cell-binding B subunit1
. These toxins are secreted into the extracellular milieu, but they act upon targets within the eukaryotic cytosol. Some AB toxins travel by vesicle carriers from the cell surface to the endoplasmic reticulum (ER) before entering the cytosol2-4
. In the ER, the catalytic A chain dissociates from the rest of the toxin and moves through a protein-conducting channel to reach its cytosolic target5
. The translocated, cytosolic A chain is difficult to detect because toxin trafficking to the ER is an extremely inefficient process: most internalized toxin is routed to the lysosomes for degradation, so only a small fraction of surface-bound toxin reaches the Golgi apparatus and ER6-12
To monitor toxin translocation from the ER to the cytosol in cultured cells, we combined a subcellular fractionation protocol with the highly sensitive detection method of surface plasmon resonance (SPR)13-15
. The plasma membrane of toxin-treated cells is selectively permeabilized with digitonin, allowing collection of a cytosolic fraction which is subsequently perfused over an SPR sensor coated with an anti-toxin A chain antibody. The antibody-coated sensor can capture and detect pg/mL quantities of cytosolic toxin. With this protocol, it is possible to follow the kinetics of toxin entry into the cytosol and to characterize inhibitory effects on the translocation event. The concentration of cytosolic toxin can also be calculated from a standard curve generated with known quantities of A chain standards that have been perfused over the sensor. Our method represents a rapid, sensitive, and quantitative detection system that does not require radiolabeling or other modifications to the target toxin.
Immunology, Issue 59, Surface plasmon resonance, AB toxin, translocation, endoplasmic reticulum, cell culture, cholera toxin, pertussis toxin
Live Cell Calcium Imaging Combined with siRNA Mediated Gene Silencing Identifies Ca2+ Leak Channels in the ER Membrane and their Regulatory Mechanisms
Institutions: Saarland University, Saarland University.
In mammalian cells, the endoplasmic reticulum (ER) plays a key role in protein biogenesis as well as in calcium signalling1
. The heterotrimeric Sec61 complex in the ER membrane provides an aqueous path for newly-synthesized polypeptides into the lumen of the ER. Recent work from various laboratories suggested that this heterotrimeric complex may also form transient Ca2+
. The key observation for this notion was that release of nascent polypeptides from the ribosome and Sec61 complex by puromycin leads to transient release of Ca2+
from the ER. Furthermore, it had been observed in vitro
that the ER luminal protein BiP is involved in preventing ion permeability at the level of the Sec61 complex9,10
. We have established an experimental system that allows us to directly address the role of the Sec61 complex as potential Ca2+
leak channel and to characterize its putative regulatory mechanisms11-13
. This system combines siRNA mediated gene silencing and live cell Ca2+
. Cells are treated with siRNAs that are directed against the coding and untranslated region (UTR), respectively, of the SEC61A1
gene or a negative control siRNA. In complementation analysis, the cells are co-transfected with an IRES-GFP vector that allows the siRNA-resistant expression of the wildtype SEC61A1
gene. Then the cells are loaded with the ratiometric Ca2+
-indicator FURA-2 to monitor simultaneously changes in the cytosolic Ca2+
concentration in a number of cells via a fluorescence microscope. The continuous measurement of cytosolic Ca2+
also allows the evaluation of the impact of various agents, such as puromycin, small molecule inhibitors, and thapsigargin on Ca2+
leakage. This experimental system gives us the unique opportunities to i) evaluate the contribution of different ER membrane proteins to passive Ca2+
efflux from the ER in various cell types, ii) characterize the proteins and mechanisms that limit this passive Ca2+
efflux, and iii) study the effects of disease linked mutations in the relevant components.
Cell Biology, Issue 53, Cellular calcium homeostasis, calmodulin, complementation, endoplasmic reticulum, ER calcium leakage, gene silencing, IQ motif, mutant analysis, Sec61 complex
Measuring Spatial and Temporal Ca2+ Signals in Arabidopsis Plants
Institutions: Purdue University, Purdue University, Jiangsu Academy of Agricultural Sciences, Zhejiang University, Shanxi Academy of Agricultural Sciences, Chinese Academy of Sciences.
Developmental and environmental cues induce Ca2+
fluctuations in plant cells. Stimulus-specific spatial-temporal Ca2+
patterns are sensed by cellular Ca2+
binding proteins that initiate Ca2+
signaling cascades. However, we still know little about how stimulus specific Ca2+
signals are generated. The specificity of a Ca2+
signal may be attributed to the sophisticated regulation of the activities of Ca2+
channels and/or transporters in response to a given stimulus. To identify these cellular components and understand their functions, it is crucial to use systems that allow a sensitive and robust recording of Ca2+
signals at both the tissue and cellular levels. Genetically encoded Ca2+
indicators that are targeted to different cellular compartments have provided a platform for live cell confocal imaging of cellular Ca2+
signals. Here we describe instructions for the use of two Ca2+
detection systems: aequorin based FAS (film adhesive seedlings) luminescence Ca2+
imaging and case12 based live cell confocal fluorescence Ca2+
imaging. Luminescence imaging using the FAS system provides a simple, robust and sensitive detection of spatial and temporal Ca2+
signals at the tissue level, while live cell confocal imaging using Case12 provides simultaneous detection of cytosolic and nuclear Ca2+
signals at a high resolution.
Plant Biology, Issue 91, Aequorin, Case12, abiotic stress, heavy metal stress, copper ion, calcium imaging, Arabidopsis
Mechanical Stimulation-induced Calcium Wave Propagation in Cell Monolayers: The Example of Bovine Corneal Endothelial Cells
Institutions: KU Leuven.
Intercellular communication is essential for the coordination of physiological processes between cells in a variety of organs and tissues, including the brain, liver, retina, cochlea and vasculature. In experimental settings, intercellular Ca2+
-waves can be elicited by applying a mechanical stimulus to a single cell. This leads to the release of the intracellular signaling molecules IP3
that initiate the propagation of the Ca2+
-wave concentrically from the mechanically stimulated cell to the neighboring cells. The main molecular pathways that control intercellular Ca2+
-wave propagation are provided by gap junction channels through the direct transfer of IP3
and by hemichannels through the release of ATP. Identification and characterization of the properties and regulation of different connexin and pannexin isoforms as gap junction channels and hemichannels are allowed by the quantification of the spread of the intercellular Ca2+
-wave, siRNA, and the use of inhibitors of gap junction channels and hemichannels. Here, we describe a method to measure intercellular Ca2+
-wave in monolayers of primary corneal endothelial cells loaded with Fluo4-AM in response to a controlled and localized mechanical stimulus provoked by an acute, short-lasting deformation of the cell as a result of touching the cell membrane with a micromanipulator-controlled glass micropipette with a tip diameter of less than 1 μm. We also describe the isolation of primary bovine corneal endothelial cells and its use as model system to assess Cx43-hemichannel activity as the driven force for intercellular Ca2+
-waves through the release of ATP. Finally, we discuss the use, advantages, limitations and alternatives of this method in the context of gap junction channel and hemichannel research.
Cellular Biology, Issue 77, Molecular Biology, Medicine, Biomedical Engineering, Biophysics, Immunology, Ophthalmology, Gap Junctions, Connexins, Connexin 43, Calcium Signaling, Ca2+, Cell Communication, Paracrine Communication, Intercellular communication, calcium wave propagation, gap junctions, hemichannels, endothelial cells, cell signaling, cell, isolation, cell culture
Inducing Plasticity of Astrocytic Receptors by Manipulation of Neuronal Firing Rates
Institutions: University of California Riverside, University of California Riverside, University of California Riverside.
Close to two decades of research has established that astrocytes in situ
and in vivo
express numerous G protein-coupled receptors (GPCRs) that can be stimulated by neuronally-released transmitter. However, the ability of astrocytic receptors to exhibit plasticity in response to changes in neuronal activity has received little attention. Here we describe a model system that can be used to globally scale up or down astrocytic group I metabotropic glutamate receptors (mGluRs) in acute brain slices. Included are methods on how to prepare parasagittal hippocampal slices, construct chambers suitable for long-term slice incubation, bidirectionally manipulate neuronal action potential frequency, load astrocytes and astrocyte processes with fluorescent Ca2+
indicator, and measure changes in astrocytic Gq GPCR activity by recording spontaneous and evoked astrocyte Ca2+
events using confocal microscopy. In essence, a “calcium roadmap” is provided for how to measure plasticity of astrocytic Gq GPCRs. Applications of the technique for study of astrocytes are discussed. Having an understanding of how astrocytic receptor signaling is affected by changes in neuronal activity has important implications for both normal synaptic function as well as processes underlying neurological disorders and neurodegenerative disease.
Neuroscience, Issue 85, astrocyte, plasticity, mGluRs, neuronal Firing, electrophysiology, Gq GPCRs, Bolus-loading, calcium, microdomains, acute slices, Hippocampus, mouse
Analysis of Tubular Membrane Networks in Cardiac Myocytes from Atria and Ventricles
Institutions: Heart Research Center Goettingen, University Medical Center Goettingen, German Center for Cardiovascular Research (DZHK) partner site Goettingen, University of Maryland School of Medicine.
In cardiac myocytes a complex network of membrane tubules - the transverse-axial tubule system (TATS) - controls deep intracellular signaling functions. While the outer surface membrane and associated TATS membrane components appear to be continuous, there are substantial differences in lipid and protein content. In ventricular myocytes (VMs), certain TATS components are highly abundant contributing to rectilinear tubule networks and regular branching 3D architectures. It is thought that peripheral TATS components propagate action potentials from the cell surface to thousands of remote intracellular sarcoendoplasmic reticulum (SER) membrane contact domains, thereby activating intracellular Ca2+
release units (CRUs). In contrast to VMs, the organization and functional role of TATS membranes in atrial myocytes (AMs) is significantly different and much less understood. Taken together, quantitative structural characterization of TATS membrane networks in healthy and diseased myocytes is an essential prerequisite towards better understanding of functional plasticity and pathophysiological reorganization. Here, we present a strategic combination of protocols for direct quantitative analysis of TATS membrane networks in living VMs and AMs. For this, we accompany primary cell isolations of mouse VMs and/or AMs with critical quality control steps and direct membrane staining protocols for fluorescence imaging of TATS membranes. Using an optimized workflow for confocal or superresolution TATS image processing, binarized and skeletonized data are generated for quantitative analysis of the TATS network and its components. Unlike previously published indirect regional aggregate image analysis strategies, our protocols enable direct characterization of specific components and derive complex physiological properties of TATS membrane networks in living myocytes with high throughput and open access software tools. In summary, the combined protocol strategy can be readily applied for quantitative TATS network studies during physiological myocyte adaptation or disease changes, comparison of different cardiac or skeletal muscle cell types, phenotyping of transgenic models, and pharmacological or therapeutic interventions.
Bioengineering, Issue 92, cardiac myocyte, atria, ventricle, heart, primary cell isolation, fluorescence microscopy, membrane tubule, transverse-axial tubule system, image analysis, image processing, T-tubule, collagenase
Identification of protein complexes with quantitative proteomics in S. cerevisiae
Institutions: University of British Columbia - UBC, University of British Columbia - UBC.
Lipids are the building blocks of cellular membranes that function as barriers and in compartmentalization of cellular processes, and recently, as important intracellular signalling molecules. However, unlike proteins, lipids are small hydrophobic molecules that traffic primarily by poorly described nonvesicular routes, which are hypothesized to occur at membrane contact sites (MCSs). MCSs are regions where the endoplasmic reticulum (ER) makes direct physical contact with a partnering organelle, e.g., plasma membrane (PM). The ER portion of ER-PM MCSs is enriched in lipid-synthesizing enzymes, suggesting that lipid synthesis is directed to these sites and implying that MCSs are important for lipid traffic. Yeast is an ideal model to study ER-PM MCSs because of their abundance, with over 1000 contacts per cell, and their conserved nature in all eukaryotes. Uncovering the proteins that constitute MCSs is critical to understanding how lipids traffic is accomplished in cells, and how they act as signaling molecules. We have found that an ER called Scs2p localize to ER-PM MCSs and is important for their formation. We are focused on uncovering the molecular partners of Scs2p. Identification of protein complexes traditionally relies on first resolving purified protein samples by gel electrophoresis, followed by in-gel digestion of protein bands and analysis of peptides by mass spectrometry. This often limits the study to a small subset of proteins. Also, protein complexes are exposed to denaturing or non-physiological conditions during the procedure. To circumvent these problems, we have implemented a large-scale quantitative proteomics technique to extract unbiased and quantified data. We use stable isotope labeling with amino acids in cell culture (SILAC) to incorporate staple isotope nuclei in proteins in an untagged control strain. Equal volumes of tagged culture and untagged, SILAC-labeled culture are mixed together and lysed by grinding in liquid nitrogen. We then carry out an affinity purification procedure to pull down protein complexes. Finally, we precipitate the protein sample, which is ready for analysis by high-performance liquid chromatography/ tandem mass spectrometry. Most importantly, proteins in the control strain are labeled by the heavy isotope and will produce a mass/ charge shift that can be quantified against the unlabeled proteins in the bait strain. Therefore, contaminants, or unspecific binding can be easily eliminated. By using this approach, we have identified several novel proteins that localize to ER-PM MCSs. Here we present a detailed description of our approach.
Biochemistry, Issue 25, Quantitative proteomics, Stable isotope, Amino acid labeling, SILAC, Isotope-coded affinity tag, Isotope labeling, Quantitation, Saccharomyces cerevisiae, ER polarization
Calcium Imaging of Cortical Neurons using Fura-2 AM
Institutions: Stanford University , Stanford University School of Medicine.
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.
Neuroscience, Issue 23, calcium imaging, calcium channels, calcium, neurons, excitable cells, time-lapse, Fura-2, Calcium indicator,intracellular calcium