This work is funded by the Wellcome Trust, The Medical Research Council, The Royal Society, Birmingham Science City and The Infertility Research Trust. We would like to acknowledge the expert assistance of Cairn Research in constructing and integrating the imaging rigs.

Sperm from healthy fertile males, with a normal semen analysis, are normally prepared for imaging as follows.
<ol>
<li> Semen samples are stored at 37&deg;C for no more than 30 min. Cells are isolated from the seminal plasma by swim up into supplemented Earle's balanced salt solution (sEBSS; mM: 1.8 CaCl2.2H2O, 5.37 KCl, 0.81 MgSO4.7H2O, 26.2 NaHCO3, 1.0 NaH2PO4.2H2O, 116.4 NaCl, 55.6 D-glucose, 2.73 Na pyruvate, 41.8 Na lactate) supplemented with 0.3% charcoal de-lipidated/fatty acid free Fraction V BSA (quality of the BSA is crucial for successful capacitation of sperm). 1 ml of sEBBS is pipetted into each of a series of 5 ml tubes and gently underlayered with 0.3 ml of semen. After incubation for 1 hour (37&deg;C; 6% CO2) the top 0.7 ml is gently removed from each tube and pooled. 10 &mu;l of the sperm suspension is diluted with 90 &mu;l of 1% (v/v) formalin to immobilize the cells, then sperm are counted in a Neubauer chamber. Cell density in the suspension is then adjusted (with sEBSS) to 6 million cells/ml.</li>
<li> The sample is then divided into aliquots of 200 &mu;l in loosely-capped tubes and incubated (37&deg;C; 6% CO2) in for 5-6 h to allow capacitation.</li>
<li> Coverslips (22x50 mm) have previously been treated with poly-D-lysine. 10 &mu;l of poly-D-lysine solution (10% w/v) is applied as a number of small drops to the centre of the coverslip. The poly-D-lysine is then allowed to air dry. This can be on a heated stage and should be to complete dryness. A coverslip is attached with vacuum grease to an enclosed, purpose-built, perfusable, polycarbonate imaging chamber (dimensions 35 mm x 20 mm x 5 mm; capacity &asymp; 180 &mu;l) similar to the Warner RC20 chamber .The poly-D-lysine-coated coverslip forms the base of the chamber when the cells are viewed on an inverted microscope and a 12 mm diameter circular coverslip forms the upper surface of the chamber, allowing transmission of light.</li>
<li> Oregon Green BAPTA1-AM (OGB) or Calcium Green 1-AM are used for labeling cells. OGB is prepared by dissolving in DMSO. Pluronic acid F127 (a detergent) is included in the DMSO to prevent 'clumping' of the dye. This can be prepared in the laboratory (100 mg Pluronic in 0.5 ml DMSO), immediately before use, or can be purchased pre-prepared. 20 &mu;l is added to a 50 &mu;g aliquot of OGB (2.5 mg/ml). The vial can then be stored frozen and thawed for later use.</li>
<li> After capacitation of the sperm (5-6 h in sEBBS, 37&deg;C; 6% CO2) a tube is selected for imaging and 1.2 &mu;l of OGB solution is added, giving a final concentration of 10 &mu;M. The tube is then incubated for a further 40 min, after which the cell suspension is gently injected into the inflow port of the imaging chamber with a p1000 (blue) pipette tip. The chamber is placed in the incubator, poly-D-lysine-coated coverslip side down, for 20 min. Cells tend to swim along the surfaces of the chamber and will adhere to the poly-D-lysine-coated area. </li>
<li> The chamber is now mounted on the microscope, connected to the perfusion apparatus and perfused at approx 0.5 ml/min. A roller pump feeds saline into the chamber and overflow from the exit port is removed by a suction pump with trap. The preparation is left in the dark for at least 10 min whilst loose cells and extracellular dye are removed by perfusion of the chamber. Temperature stability is crucially important because small fluctuations may not only affect cell Ca2+ homeostasis, but may also affect Kd of the dye. Experiments are normally performed at 25&deg;C, achieved by maintaining the temperature of the dark room at this level. We have found that maintaining room temperature at the required level provides greater temperature and focus stability than using a temperature controlled stage or a thermostatically controlled heater on the saline inflow. </li>
<li> Cells are viewed under phase contrast (40x or 60x oil-immersion) objective and an area for imaging is selected. The portion of the poly-D-lysine coated area that is near the inlet port is preferred, where saline flow is laminar and consistent. It is also important to use an area where cell density is appropriate (excessive density affects the quality and integrity of data through pickup of signal from adjacent cells) and where cells are adequately attached but flagellar activity is discernible. A phase contrast image is saved. The microscope is then switched to fluorescence mode (excitation 480 nm; emission 540 nm) and exposure time is reduced to the minimum required to obtain a clear fluorescence image (usually 50-200 ms).</li>
<li> The experiment is started by activating the time-series image acquisition software (IQ). Typically images are acquired at 0.1 Hz and illumination is restricted to the periods of image acquisition only using a software controlled shutter. Much greater image acquisition rates can be used but allowance must be made for problems of bleaching and generation of artifacts due to photo-damage to the cells (see discussion). IQ (and most other acquisition software platforms) will allow real-time graphing of fluorescence during the experiment.</li>
<li> After a control period of 3-5 minutes duration manipulation of saline constituents (such as [Ca2+]o) and application of drugs is achieved by replacement of saline in the perfusion header. Time of addition of each stimulus is noted so that time of arrival in the imaging chamber can be calculated, providing sufficient accuracy for assessment of the slow responses described here (sampled at 0.1 Hz). For more rapid responses greater accuracy can be achieved by adding stimuli directly to the saline flow through a second inflow port as provided in the Warner RC20 chamber. </li>
<li> Images are analyzed offline using IQ. Regions of interest (ROIs) are drawn round each cell (or part of cell) and also a cell free area is selected (for automatic background subtraction by the software. The image series is then checked to remove cells which died or underwent acrosome reaction (which appears as a large and rapid increase in fluorescence followed by loss of cytoplasmic dye) during the experiment and those that showed sufficient movement to cause artifacts when analyzed as a time series.</li>
<li>A time-fluorescence intensity series is then obtained for each ROI and these data are analyzed offline in Excel. Initially fluorescence is normalized to the mean value obtained during the control period, using the equation R = [ (F Frest) / Frest] x 100%,where R is % change in fluorescence compared to the control period,, F is Fluorescence intensity at a time t and Frest is the mean of at least 30 determinations of F obtained during the control period.</li>
</ol>
Representative Results
Figure 1 shows a series of images from an experiment in which the cells were stimulated with 3 &mu;M progesterone. The upper row and movie 1 show the fluorescence images in grey-scale and the same mages are shown below (and in movie 2) in pseudocolor (lookup table '16_colors' in Image J), where 'cool' colors indicate low fluorescent intensity and 'warm' colors indicate high fluorescence intensity. Cells can be viewed in pseudocolor during the experiment, using lookup tables in IQ, but using grey scale is generally more informative for assessing .whether the cells are well-labeled. The first two images were collected at 0 s and 100 s after commencing recording. Fluorescence should be stable and tends to be strongest in the post acrosomal area and sperm neck. Progesterone entered the imaging chamber at approximately 175 s and the 3rd and 4th images (180 s and 220 s) shows a rapid increase in [Ca2+]i (fluorescence) originating in posterior head and neck region of the cell. Subsequent images are at 290, 360, 740, 990 and 1440 s, showing the decay of the initial [Ca2+]i transient and development of a secondary sustained elevation. Figure 2 shows a spreadsheet analysis, converting raw fluorescence values for each ROI to normalized fluorescence (% change in fluorescence compared to the control period) Figure 3 shows time-normalized fluorescence plots for 3 cells showing 'typical' responses to 3 &mu;M progesterone.
<img src="/files/ftp_upload/1996/1996fig1.jpg" alt="Figure 1" /> Figure 1. Example data from a cell stimulated with progesterone. A series of fluorescence images in grey scale (upper row) and pseudocolor (lower row) are shown. Time points are 0, 180, 220, 290, 360 and 990s. 3 &mu;M progesterone entered the imaging chamber at 175 s. ROIs are shown in the first panel. 
<img src="/files/ftp_upload/1996/1996fig2.jpg" alt="Figure 2" /> Figure 2. Analysis of single cell fluorescence data in Excel. Figures in black are time series of fluorescence data , arranged vertically. Figures in red (below) are normalized data obtained using the equation given in the text. Graph shows normalized fluorescence-time plot for cell 64 in this experiment which, upon stimulation, shows a transient rise in fluorescence followed by a slow rise and series of small oscillations. 
<img src="/files/ftp_upload/1996/1996fig3.jpg" alt="Figure 3" /> Figure 3. Fluorescence-time traces for 3 individual cells, expressed as % change with respect to control value.. 3 &mu;M progesterone was added at the time marked by the arrow. Dashed line shows mean control fluorescence.

<ol>
	<li>Haughland, R. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Handbook of Fluorescent Probes and Research Products. , (2002).</li><li>Nishigaki, T., Wood, C. D., Shiba, K., Baba, S. A., Darszon, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stroboscopic+illumination+using+light-emitting+diodes+reduces+phototoxicity+in+fluorescence+cell+imaging.">Stroboscopic illumination using light-emitting diodes reduces phototoxicity in fluorescence cell imaging.</a> Biotechniques. 41, 191-197 (2006).</li></ol>

No conflicts of interest declared.

When carried out successfully, this technique allows recording of the distribution and kinetics of spontaneous and induced Ca2+ signals in human sperm. Responses can be obtained from a large number of cells (up to 200) which is important since human sperm can show a lot of variation in their spontaneous and stimulated Ca2+ signals. Successful labeling and survival of the cells during recording are highly dependent on sample quality. Poor samples give poor labeling , poor responses and cells may die during recording. Human sperm are very sensitive to photo-damage and we therefore use the minimal necessary illumination (both intensity and exposure time) in order to maximize cell survival and minimize artifacts due to cell death. A high-sensitivity camera (permitting use of low-intensity illumination) is a great benefit since the camera will pick up weak fluorescence. Back-illuminated, EM-CCD cameras are particularly well-suited, though very expensive. LED illumination (instead of using a xenon or mercury lamp with excitation filter also improves cell survival (Nishigaki et al, 2006). We do not use Fura-2, despite the obvious advantages of ratio-metric imaging, because Fura requires excitation with UV light and because it is necessary to take two images for each ratio. For data obtained with single wavelength, visible light dyes, normalization of the fluorescence intensity largely compensates for difference in dye loading between cells, but not for dye distribution within an individual cell, and this must be borne in mind. Though single wavelength dyes can, in principle, be calibrated, the accuracy of the technique is poor and we do not attempt to do this. Whereas the ratio data obtained with Fura-2 (or the emission ratio dye indo), even without calibration, give an acceptably faithful representation of relative changes in [Ca2+]i, the fluorescence intensity of the single wavelength dyes is far from linearly related to [Ca2+]i. Over the most useful part of the saturation curve the relationship for single wavelength dyes usually approximates to logarithmic. We currently use Oregon Green BAPTA-1 (Figure 4) because it is sensitive to small changes in [Ca2+]i close to resting level s (50-100 nM). When [Ca2+]i rises above 1 &mu;M responses will be under-represented in terms of fluorescence change and the dye may saturate. An option for improvement of the technique without resorting to UV excitation is to double load cells with a dye such as Fluo-3 and Fura red. Both can be excited at 488 nM but simultaneous their responses are very different. Recording of emission at 540 and 650 nM provides a ratio which can provide a better method for accurate monitoring of [Ca2+]i (Haughland, 2002) and may be applicable to sperm (Nisigaki et al, 2006). <img src="/files/ftp_upload/1996/1996fig4.jpg" alt="Figure 4" /> Figure 4. Relationship between free [Ca2+] (nM) and fluorescence emission at peak wavelength (approx 525 nm) for Oregon Green BAPTA-1 and Oregon Green BAPTA-2. OGB1 gives a higher fluorescence at resting [Ca2+] but saturates at lower levels and thus has a smaller useable range. Data for these plots were obtained from the Molecular Probes handbook (Haughland, 2002).

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		<th>Material Name</th>
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<td>Low light level Camera</td>
<td>&nbsp;</td>
<td>QImaging</td>
<td>Rolera XR</td>
<td>&nbsp;</td>
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<td>Oregon Green BAPTA-1</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>06807</td>
<td>&nbsp;</td>
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<td>&nbsp;</td>
<td>Andor</td>
<td>IQ</td>
<td>&nbsp;</td>
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<td>Imaging Software</td>
<td>&nbsp;</td>
<td>National Institues of Health</td>
<td>Image J</td>
<td>Public domain software</td>
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<td>LED illumination system</td>
<td>&nbsp;</td>
<td>Cairn Research</td>
<td>Opto LED</td>
<td>&nbsp;</td>
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<td>Pluronic F-127 (20% solution in DMSO)</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
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techniques for imaging ca2+ signaling in human sperm

Fluorescence microscopy of cells loaded with fluorescent, Ca2+-sensitive dyes is used for measurement of spatial and temporal aspects of Ca2+ signaling in live cells. Here we describe the method used in our laboratories for loading suspensions of human sperm with Ca2+-reporting dyes and measuring the fluorescence signal during physiological stimulation. Motile cells are isolated by direct swim-up and incubated under capacitating conditions for 0-24 h, depending upon the experiment. The cell-permeant AM (acetoxy methyl ester) ester form of the Ca2+-reporting dye is then added to a cell aliquot and a period of 1 h is allowed for loading of the dye into the cytoplasm. We use visible wavelength dyes to minimize photo-damage to the cells, but this means that ratiometric recording is not possible. Advantages and disadvantages of this approach are discussed. During the loading period cells are introduced into an imaging chamber and allowed to adhere to a poly-D-lysine coated coverslip. At the end of the loading period excess dye and loose cells are removed by connection of the chamber to the perfusion apparatus. The chamber is perfused continuously, stimuli and modified salines are then added to the perfusion header. Experiments are recorded by time-lapse acquisition of fluorescence images and analyzed in detail offline, by manually drawing regions of interest. Data are normalized to pre-stimulus levels such that, for each cell (or part of a cell), a graph showing the Ca2+ response as % change in fluorescence is obtained.

Watch this Scientific Journal Video about Techniques for Imaging Ca2+ Signaling in Human Sperm at JoVE.com

Techniques for Imaging Ca2+ Signaling in Human Sperm

Stimulus-evoked [Ca2+]i signals of individual human sperm are assessed. Motile cells are loaded with Ca2+-sensitive fluorescent dye (AM-ester method) and immobilised in a perfusable chamber. Cells are imaged by time-lapse fluorescence microscopy and stimulated via the perfusing medium. Responses of single cells (or regions) are analysed offline using Excel.

Techniques for Imaging Ca2+ Signaling in Human Sperm

Fluorescence microscopy of cells loaded with fluorescent, Ca2+-sensitive dyes is used for measurement of spatial and temporal aspects of Ca2+ signaling in ...

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17.0K Views.  University of Birmingham. Stimulus-evoked [Ca2+]i signals of individual human sperm are assessed. Motile cells are loaded with Ca2+-sensitive fluorescent dye (AM-ester method) and immobilised in a perfusable chamber. Cells are imaged by time-lapse fluorescence microscopy and stimulated via the perfusing medium. Responses of single cells (or regions) are analysed offline using Excel.

Video: Techniques for Imaging Ca2+ Signaling in Human Sperm

In vivo Ca2+- Imaging of Mushroom Body Neurons During Olfactory Learning in the Honey Bee

The in vivo and semi-in vivo preparation for Calcium imaging has been developed in our lab by Joerges, K&uuml;ttner and Galizia over ten years ago, to measure odor evoked activity in the antennal lobe1. From then on, it has been continuously refined and applied to different neuropiles in the bee brain. Here, we describe the preparation currently used in the lab to measure activity in mushroom body neurons using a dextran coupled calcium-sensitive dye (Fura-2). We retrogradely stain mushroom body neurons by injecting dye into their axons or soma region. We focus on reducing the invasiveness, to achieve a preparation in which it is still possible to train the bee using PER conditioning. We are able to monitor and quantify the behavioral response by recording electro-myograms from the muscle which controls the PER (M17)2.
After the physiological experiment the imaged structures are investigated in greater detail using confocal scanning microscopy to address the identity of the neurons.

In vivo Ca2+- Imaging of Mushroom Body Neurons During Olfactory Learning in the Honey Bee

Bees can be conditioned in an appetitive olfactory learning paradigm (PER-conditioning). Using odors as stimuli, we established a method in which behavior is recorded while simultaneously Calcium Imaging is used to measure odor evoked activity in mushroom body neurons in vivo.

The in vivo and semi-in vivo preparation for Calcium imaging has been developed in our lab by Joerges, K&uuml;ttner and Galizia over ten years ago, to ...

Fluorescence in situ hybridization (FISH) Protocol in Human Sperm

Aneuploidies are the most frequent chromosomal abnormalities in humans. Most of these abnormalities result from meiotic errors during the gametogenic process in the parents. In human males, these errors can lead to the production of spermatozoa with numerical chromosome abnormalities which represent an increased risk of transmitting these anomalies to the offspring.
For this reason, the technique of fluorescence in situ hybridization (FISH) on sperm nuclei has become a protocol widely incorporated in the context of clinical diagnosis. This practice provides an estimate of the frequencies of numerical chromosome abnormalities in the gametes of the patients that seek for genetic reproductive advice.
To date, the chromosomes most frequently included in sperm FISH analysis are chromosomes X, Y, 13, 18 and 21.
This video-article describes, step by step, how to process and fix a human semen sample, how to decondense and denature the sperm chromatin, how to proceed to obtain sperm FISH preparations, and how to visualize the results at the microscope. Special remarks of the most relevant steps are given to achieve the best results.

Fluorescence in situ hybridization FISH Protocol in Human Sperm

This video-article describes, step by step, how to process a semen sample to achieve good-quality fluorescence in situ hybridization on human spermatozoa. Preparations obtained can be used for aneuploidy screening in the context of clinical diagnosis.

Aneuploidies are the most frequent chromosomal abnormalities in humans. Most of these abnormalities result from meiotic errors during the gametogenic ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

Visualization of Vascular Ca2+ Signaling Triggered by Paracrine Derived ROS

Oxidative stress has been implicated in a number of pathologic conditions including ischemia/reperfusion damage and sepsis. The concept of oxidative stress refers to the aberrant formation of ROS (reactive oxygen species), which include O2&bull;-, H2O2, and hydroxyl radicals. Reactive oxygen species influences a multitude of cellular processes including signal transduction, cell proliferation and cell death1-6. ROS have the potential to damage vascular and organ cells directly, and can initiate secondary chemical reactions and genetic alterations that ultimately result in an amplification of the initial ROS-mediated tissue damage. A key component of the amplification cascade that exacerbates irreversible tissue damage is the recruitment and activation of circulating inflammatory cells. During inflammation, inflammatory cells produce cytokines such as tumor necrosis factor-&alpha; (TNF&alpha;) and IL-1 that activate endothelial cells (EC) and epithelial cells and further augment the inflammatory response7. Vascular endothelial dysfunction is an established feature of acute inflammation. Macrophages contribute to endothelial dysfunction during inflammation by mechanisms that remain unclear. Activation of macrophages results in the extracellular release of O2&bull;- and various pro-inflammatory cytokines, which triggers pathologic signaling in adjacent cells8. NADPH oxidases are the major and primary source of ROS in most of the cell types. Recently, it is shown by us and others9,10 that ROS produced by NADPH oxidases induce the mitochondrial ROS production during many pathophysiological conditions. Hence measuring the mitochondrial ROS production is equally important in addition to measuring cytosolic ROS. Macrophages produce ROS by the flavoprotein enzyme NADPH oxidase which plays a primary role in inflammation. Once activated, phagocytic NADPH oxidase produces copious amounts of O2&bull;- that are important in the host defense mechanism11,12. Although paracrine-derived O2&bull;- plays an important role in the pathogenesis of vascular diseases, visualization of paracrine ROS-induced intracellular signaling including Ca2+ mobilization is still hypothesis. We have developed a model in which activated macrophages are used as a source of O2&bull;- to transduce a signal to adjacent endothelial cells. Using this model we demonstrate that macrophage-derived O2&bull;- lead to calcium signaling in adjacent endothelial cells.

Visualization of Vascular Ca2+ Signaling Triggered by Paracrine Derived ROS

An efficient method to gain insights into visualizing the paracrine-derived ROS induction of endothelial Ca2+ signaling is described. This method takes advantage of measuring paracrine derived ROS triggered Ca2+ mobilization in vascular endothelial cells in a co-culture model.

Oxidative stress has been implicated in a number of pathologic conditions including ischemia/reperfusion damage and sepsis. The concept of oxidative ...

Isolation of Human Atrial Myocytes for Simultaneous Measurements of Ca2+ Transients and Membrane Currents

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 &mu;M 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+ and 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.

Isolation of Human Atrial Myocytes for Simultaneous Measurements of Ca2+ Transients and Membrane Currents

We describe the isolation of human atrial myocytes which can be used for intracellular Ca2+ measurements in combination with electrophysiological patch-clamp studies.

The study of  electrophysiological properties of cardiac ion channels with the patch-clamp  technique and the exploration of cardiac cellular Ca2+ ...

Measuring Intracellular Ca2+ Changes in Human Sperm using Four Techniques: Conventional Fluorometry, Stopped Flow Fluorometry, Flow Cytometry and Single Cell Imaging

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.

Measuring Intracellular Ca2+ Changes in Human Sperm using Four Techniques: Conventional Fluorometry, Stopped Flow Fluorometry, Flow Cytometry and Single Cell Imaging

Intracellular Ca2+ dynamics are very important in sperm physiology and Ca2+-sensitive fluorescent dyes constitute a versatile tool to study them. Population experiments (fluorometry and stopped flow fluorometry) and single cell experiments (flow cytometry and single cell imaging) are used to track spatio-temporal [Ca2+] changes in human sperm cells.

Spermatozoa  are male reproductive cells especially designed to reach, recognize and fuse  with the egg. To perform these tasks, sperm cells must be ...

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, ...

Two-photon Imaging of Intracellular Ca2+ Handling and Nitric Oxide Production in Endothelial and Smooth Muscle Cells of an Isolated Rat Aorta

Calcium is a very important regulator of many physiological processes in vascular tissues. Most endothelial and smooth muscle functions highly depend on changes in intracellular calcium ([Ca2+]i) and nitric oxide (NO). In order to understand how [Ca2+]i, NO and downstream molecules are handled by a blood vessel in response to vasoconstrictors and vasodilators, we developed a novel technique that applies calcium-labeling (or NO-labeling) dyes with two photon microscopy to measure calcium handling (or NO production) in isolated blood vessels. Described here is a detailed step-by-step procedure that demonstrates how to isolate an aorta from a rat, label calcium or NO within the endothelial or smooth muscle cells, and image calcium transients (or NO production) using a two photon microscope following physiological or pharmacological stimuli. The benefits of using the method are multi-fold: 1) it is possible to simultaneously measure calcium transients in both endothelial cells and smooth muscle cells in response to different stimuli; 2) it allows one to image endothelial cells and smooth muscle cells in their native setting; 3) this method is very sensitive to intracellular calcium or NO changes and generates high resolution images for precise measurements; and 4) described approach can be applied to the measurement of other molecules, such as reactive oxygen species. In summary, application of two photon laser emission microscopy to monitor calcium transients and NO production in the endothelial and smooth muscle cells of an isolated blood vessel has provided high quality quantitative data and promoted our understanding of the mechanisms regulating vascular function.

Two-photon Imaging of Intracellular Ca2+ Handling and Nitric Oxide Production in Endothelial and Smooth Muscle Cells of an Isolated Rat Aorta

Vascular cell functiondepends on activity of intracellular messengers. Described here is an ex vivo two photon imaging method that allows the measurement of intracellular calcium and nitric oxide levels in response to physiological and pharmacological stimuli in individual endothelial and smooth muscle cells of an isolated aorta.

Calcium is a very important regulator of many physiological processes in vascular tissues. Most endothelial and smooth muscle functions highly depend on ...

Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes

Chromosome segregation in human oocytes is error prone, resulting in aneuploidy, which is the leading genetic cause of miscarriage and birth defects. The study of chromosome behavior in oocytes from model organisms holds much promise to uncover the molecular basis of the susceptibility of human oocytes to aneuploidy. Drosophila melanogaster is amenable to genetic manipulation, with over 100 years of research, community, and technique development. Visualizing chromosome behavior and spindle assembly in Drosophila oocytes has particular challenges, however, due primarily to the presence of membranes surrounding the oocyte that are impenetrable to antibodies. We describe here protocols for the collection, preparation, and imaging of meiosis I spindle assembly and chromosome behavior in Drosophila oocytes, which allow the molecular dissection of chromosome segregation in this important model organism.

Techniques for Imaging Prometaphase and Metaphase of Meiosis I in Fixed Drosophila Oocytes

We present protocols for the collection, preparation, and imaging of mature Drosophila oocytes. These methods allow the visualization of chromosome behavior and spindle assembly and function during meiosis.

Chromosome segregation in human oocytes is error prone, resulting in aneuploidy, which is the leading genetic cause of miscarriage and birth defects. The ...

Discrimination and Characterization of Heterocellular Populations Using Quantitative Imaging Techniques

Cellular processes are complex and result from the interplay between multiple cell types and their environment. Existing cell biology techniques often do not allow for accurate interpretation of this interplay. Using a quantitative imaging-based approach, we present a high-content protocol for characterizing the dynamic phenotypic responses (i.e. morphology changes, proliferation, apoptosis) of heterogeneous cell populations to changes in environmental stimuli. We highlight our ability to distinguish between cell types based upon either fluorescence intensity or inherent morphology features depending on the application. This platform allows for a more comprehensive characterization of subpopulation response to perturbation while utilizing shorter time, smaller amounts of reagents, and lower likelihood of error than traditional cell biology assays. However, in some cases, cell populations may be difficult to identify and quantitate based on complex cellular features and will require additional troubleshooting; we highlight some of these circumstances in the protocol. We demonstrate this application using response to drug in a cancer model; however, it can easily be applied more broadly to other physiological processes. This protocol allows one to identify subpopulations within a co-culture system and characterize the particular response of each to external stimuli.

We have developed a novel protocol for studying heterocellular population dynamics in response to perturbations. This manuscript describes an imaging-based platform that produces quantitative datasets for simultaneous characterization of multiple cellular phenotypes of heterocellular populations in a robust manner.

Cellular processes are complex and result from the interplay between multiple cell types and their environment. Existing cell biology techniques often do ...