1. The Importance of Cell Counting
<ol>
<li>Counting cells is a necessary but tedious step for in vitro cell culture.</li>
<li>Consistent cell concentrations ensure experimental reproducibility and accuracy.</li>
<li>Cell counts are important for monitoring cell health and proliferation rate, assessing immortalization or transformation, seeding cells for subsequent experiments, transfection or infection, and preparing for cell-based assays.</li>
<li>Despite the need for speed and accuracy in cell counting,, 71% of 400 researchers surveyed count cells using a hemocytometer. While hemocytometry is inexpensive, it is laborious and subject to user bias and misuse, which results in inaccurate counts.</li>
<li>Sources of error in hemocytometry include: uneven cell distribution in the sample, too many or too few cells in the sample, subjective decisions as to whether a given cell falls within the defined counting area, contamination of the hemocytometer, user-to-user variation, and variation of hemocytometer filling rate.</li>
<li>To alleviate the tedium associated with manual counting, 29% of researchers count cells using automated cell counting devices. However, for most researchers, the price associated with these large bench-top instruments is a barrier to their use.</li>
<li>The Scepter cell counter is an automated handheld device that offers the automation and accuracy of Coulter counting at a relatively low cost.</li>
<li>The system employs the Coulter principle of impedance-based particle detection in a miniaturized format using a combination of analog and digital hardware for sensing, signal processing, data storage, and graphical display.</li>
<li>The disposable sensor is engineered with a microfabricated, cell- sensing zone that enables discrimination by cell size and cell volume at sub-micron and sub-picoliter resolution.</li>
<li>Enhanced with precision liquid-handling channels and electronics, the Scepter cell counter graphically displays cell population statistics as a histogram.</li>
</ol>
2. Preparation of a Single-cell Suspension for Counting
<ol>
<li>To use the Scepter cell counter, begin by diluting a single cell suspension of cells, in phosphate buffered saline. A concentration of 10,000-500,000 cells per mL is optimal.</li>
<li>Transfer 100 &mu;L of diluted cells to a 1.5 mL microfuge tube.</li>
</ol>
3. Performing Cell Counts using the Scepter
<ol>
<li>Turn on the Scepter cell counter by pressing and holding the toggle on the back of the instrument and wait for the on-screen instructions to appear.</li>
<li>When prompted, attach a sensor to the end of the Scepter unit with the electrode sensing panel facing toward the front of the instrument.</li>
<li>Scepter sensors consist of a precision molded sampling chamber, with an electronic sensing zone and integrated cell-sensing electrodes. The sensors make it possible to discriminate cell size at sub-micron resolution, and cell volume at sub-picoliter resolution.</li>
<li>Consistent sensor engineering, ensures that equal sample volumes are counted and accurate size measurements are made.</li>
<li>Once the sensor is attached, perform each step of the counting process according to the on-screen instructions as follows:</li>
<li>When the screen reads "Hold down the plunger to begin," depress the plunger. The screen will then read "submerge the sensor." Submerge the sensor into the cell suspension solution and release the plunger to aspirate the cell suspension into the sensor.</li>
<li>The plunger acts as an electrical signal and will draw the sample plus dead volume into the sensor. The speed or accuracy of plunging does not matter as it does with pipetting. This process will take several seconds. When ready, the instrument will beep, and "Sample loaded" will be displayed on the screen.</li>
<li>Once the sample is loaded into the sensor it is drawn through the sensing zone to assess cell size and cell volume, then through the microchannels. Once 50 &mu;L of sample is counted the Sensor tells the instrument to stop counting.</li>
<li>After the count is complete, the instrument's screen will read "Count Complete. Please remove sensor and discard." Follow the prompt, and remove and discard the sensor.</li>
<li>The instrument will then display a histogram of cell size or diameter on its screen as well as the cell concentration per mL.</li>
</ol>
4. Data Analysis
<ol>
<li>After the count is complete and the histogram is displayed on the instrument, apply gates by pushing the toggle button. Select "auto gating" or "use last" to either automatically set the gate based on the histogram profile, or to use the gates from the previous count.</li>
<li>Regardless of the selection, the gates can be manually moved to fine-tune the data. Push the toggle button again to select the left gate and scroll the toggle to move the gate as desired.</li>
<li>Then press the toggle again to activate the right gate and scroll the toggle to place the right gate. Click again; a new cell concentration, average cell volume and diameter will be displayed.</li>
<li>The toggle can also be moved to either side to switch the display between date, time, mean cell size, and mean cell volume.</li>
<li>Up to 72 histograms can be stored on the instrument itself. To upload the data to a personal computer, attach the instrument via the USB cable port on the display screen.</li>
<li>On the PC, double click on the Scepter Software Application to launch the software then click on "connect". All counts will automatically port to the software.</li>
<li>The Scepter Application Software displays the histogram detailing the population distribution of the culture in either volume or diameter. Downloaded files can then be exported to Microsoft Excel, where the raw data will be displayed in bins of specified cell diameters or cell volumes.</li>
</ol>
5. Representative Results
<ol>
<li>Nineteen different cell lines were prepared for counting and theoretical starting concentrations were determined using a Coulter Counter. These cell suspensions were then further diluted to obtain cell suspensions at theoretical cell concentrations.</li>
<li>The Scepter cell counter was then used to determine cell concentrations. Figure 1 shows 5 dilutions that were measured for 5 representative cell lines along with the theoretical cell concentration.</li>
<li>The high degree of linearity as indicated by the R2 values, demonstrates that Scepter counting is a reliable method for the cell lines tested, across a wide linear operating range.</li>
<li>The overall relative accuracy of Scepter counting is demonstrated in Figure 2, in which data obtained from Scepter counting are compared with counts obtained from Coulter Counting.</li>
<li>The log of Scepter cell counts is plotted against the log of Coulter counts, and shows that, across all cell lines tested, Scepter counts match Coulter counts.</li>
<li>Counts of 19 cell lines were performed using a Z2 Coulter Counter, the Scepter cell counter, an automated vision-based counter such as Vi-Cell or Countless system, and a hemocytemeter. </li>
<li>Counts were performed according to the manufacturer's instructions, using the same cell starting suspension and identical dilutions. The results shown in Figure 3 demonstrate that cell concentrations measured by Scepter counting closely match theoretical cell concentrations with high linearity.</li>
<li>The percent coefficient of variation, or CV, was calculated for each dilution of each cell line for each counting system. Figure 4 shows the average %CVs over 19 cell lines with respect to cell concentration and counting method.</li>
<li>Within the operating range, the Scepter cell counter is more precise than vision based hemocytometer counting, displaying smaller standard deviations and smaller average coefficients of variation.</li>
<li>Scepter counting is ten times faster than hemocytometry and faster than other automated counters too. In Figure 5, SF9, MCF7, and HEK293 cells were counted using the methods described in this video.</li>
<li>The average time required to measure cell concentration was recorded. Note that hemocytometry was performed only once per sample.</li>
<li>Scepter makes measurements based on cell size. Thus any changes in cellular morphology can be captured in the histogram. Healthy cultures display a nice bell shaped Gaussian distribution, while unhealthy cells, lyse, and cause a shift in the histogram to the left (Figure 6, left panel).</li>
<li>Similarly, when a population of 50% dead cells were mixed with 50% live cells, histogram shifts were also observed (Figure 6, right panel).</li>
</ol>
<img src="/files/ftp_upload/2204/2204fig1.jpg" alt="Figure 1" />
<img src="/files/ftp_upload/2204/2204fig2.jpg" alt="Figure 2" />
<img src="/files/ftp_upload/2204/2204fig3.jpg" alt="Figure 3" />
<img src="/files/ftp_upload/2204/2204fig4.jpg" alt="Figure 4" />
<img src="/files/ftp_upload/2204/2204fig5.jpg" alt="Figure 5" />
<img src="/files/ftp_upload/2204/2204fig6.jpg" alt="Figure 6" />
<img src="/files/ftp_upload/2204/2204fig7.jpg" alt="Figure 7" />

<ol>
	<li>Barghshoon, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Cell Counting Survey. , (2009).</li><li>Tucker, K. G., Chalder, S., al-Rubeai, M., Thomas, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measurement+of+hybridoma+cell+number,+viability,+and+morphology+using+fully+automated+image+analysis.">Measurement of hybridoma cell number, viability, and morphology using fully automated image analysis.</a> Enzyme Microb Technol. 16 (1), 29-35 (1994).</li><li>Houwen, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Fifty years of hematology innovation: the Coulter principle. , (2003).</li></ol>

Kathleen Ongena, Chandreyee Das, Janet L. Smith, S&oacute;nia Gil, and Grace Johnston are employed by Millipore, which produces the instruments used in this article.

Comparing the performance of the Scepter cell counter to results from other counting methods, we conclude that this new handheld, automated cell counter delivers precise, fast, and reliable cell counts over a wide operating range. The functionality of Scepter counting is a result of the precision engineered technology embedded into the sensor tip and the sophisticated counting instrumentation based upon the Coulter principle. The performance quality suggests Scepter counting as method to increase the speed and improve the reproducibility of cell-based assays. The handheld format allows quick tedium free counting right at the cell culture hood.

<table border="1">
 <tbody>
 <tr>
 <td>Material Name</td>
 <td>Company</td>
 <td>Catalogue Number</td>
 <td>Comment</td>
 </tr>
 <tr>
 <td>Scepter 2.0 Handheld Automated Cell Counter with 40 &mu;m Scepter Sensors</td>
 <td>EMD Millipore</td>
 <td>PHCC20040</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Scepter 2.0 Handheld Automated Cell Counter with 60 &mu;m Scepter Sensors</td>
 <td>EMD Millipore</td>
 <td>PHCC20060</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Scepter Sensors, 60 &mu;m</td>
 <td>EMD Millipore</td>
 <td>PHCC60050</td>
 <td>50 Pack of sensors</td>
 </tr>
 <tr>
 <td>Scepter Sensors, 40 &mu;m</td>
 <td>EMD Millipore</td>
 <td>PHCC40050</td>
 <td>50 Pack of sensors</td>
 </tr>
 <tr>
 <td>Universal Power Adapter</td>
 <td>EMD Millipore</td>
 <td>PHCCPOWER</td>
 <td> </td>
 </tr>
 <tr>
 <td>Scepter O-Ring Kit</td>
 <td>EMD Millipore</td>
 <td>PHCCOCLIP</td>
 <td>2 O-rings and one filter cover</td>
 </tr>
 </tbody>
</table>

determining cell number during cell culture using the scepter cell counter

Counting cells is often a necessary but tedious step for in vitro cell culture. Consistent cell concentrations ensure experimental reproducibility and accuracy. Cell counts are important for monitoring cell health and proliferation rate, assessing immortalization or transformation, seeding cells for subsequent experiments, transfection or infection, and preparing for cell-based assays. It is important that cell counts be accurate, consistent, and fast, particularly for quantitative measurements of cellular responses.
Despite this need for speed and accuracy in cell counting, 71% of 400 researchers surveyed1 who count cells using a hemocytometer. While hemocytometry is inexpensive, it is laborious and subject to user bias and misuse, which results in inaccurate counts. Hemocytometers are made of special optical glass on which cell suspensions are loaded in specified volumes and counted under a microscope. Sources of errors in hemocytometry include: uneven cell distribution in the sample, too many or too few cells in the sample, subjective decisions as to whether a given cell falls within the defined counting area, contamination of the hemocytometer, user-to-user variation, and variation of hemocytometer filling rate2.
To alleviate the tedium associated with manual counting, 29% of researchers count cells using automated cell counting devices; these include vision-based counters, systems that detect cells using the Coulter principle, or flow cytometry1. For most researchers, the main barrier to using an automated system is the price associated with these large benchtop instruments1.
The Scepter cell counter is an automated handheld device that offers the automation and accuracy of Coulter counting at a relatively low cost. The system employs the Coulter principle of impedance-based particle detection3 in a miniaturized format using a combination of analog and digital hardware for sensing, signal processing, data storage, and graphical display. The disposable tip is engineered with a microfabricated, cell- sensing zone that enables discrimination by cell size and cell volume at sub-micron and sub-picoliter resolution. Enhanced with precision liquid-handling channels and electronics, the Scepter cell counter reports cell population statistics graphically displayed as a histogram.

Watch this Scientific Journal Video about Determining Cell Number During Cell Culture using the Scepter Cell Counter at JoVE.com

Determining Cell Number During Cell Culture using the Scepter Cell Counter

The Scepter Cell Counter is a handheld automated device that can be used to count cells, monitor cell diameter and volume, and be used to check the health and quality of cellular populations from one culture to the next.

Counting cells is often a necessary but tedious step for in vitro cell culture. Consistent cell concentrations ensure experimental reproducibility and ...

determining-cell-number-during-cell-culture-using-scepter-cell

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37.9K Views.  Millipore Inc. Il contatore di cellule Scettro è un dispositivo portatile automatizzato che può essere usato per contare le celle, monitorare diametro delle cellule e il volume, e possono essere usate per controllare la salute e la qualità di popolazioni cellulari da una cultura all'altra.

Video: Determining Cell Number During Cell Culture using the Scepter Cell Counter

Human ES cells: Starting Culture from Frozen Cells

Here we demonstrate how our lab begins a HuES human embryonic stem cell line culture from a frozen stock.  First, a one to two day old ten cm plate of  approximately one (to two) million irradiated mouse embryonic fibroblast feeder cells is rinsed with HuES media to remove residual serum and cell debris, and then HuES media added and left to equilibrate in the cell culture incubator.  A frozen vial of cells from long term liquid nitrogen storage or a -80C freezer is sourced and quickly submerged in a 37C water bath for quick thawing.  Cells in freezing media are then removed from the vial and placed in a large volume of HuES media.  The large volume of HuES media facilitates removal of excess serum and DMSO, which can cause HuES human embryonic stem cells to differentiate.  Cells are gently spun out of suspension, and then re-suspended in a small volume of fresh HuES media that is then used to seed the MEF plate.  It is considered important to seed the MEF plate by gently adding the HuES cells in a drop wise fashion to evenly disperse them throughout the plate.  The newly established HuES culture plate is returned to the incubator for 48 hrs before media is replaced, then is fed every 24 hours thereafter.

Here we demonstrate how our lab begins a HuES human embryonic stem cell line culture from a frozen stock.

Here we demonstrate how our lab begins a HuES human embryonic stem cell line culture from a frozen stock.  First, a one to two day old ten cm plate of  ...

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, ...

Lentiviral-mediated Knockdown During Ex Vivo Erythropoiesis of Human Hematopoietic Stem Cells

Erythropoiesis is a commonly used model system to study cell differentiation. During erythropoiesis, pluripotent adult human hematopoietic stem cells (HSCs) differentiate into oligopotent progenitors, committed precursors and mature red blood cells 1. This process is regulated for a large part at the level of gene expression, whereby specific transcription factors activate lineage-specific genes while concomitantly repressing genes that are specific to other cell types 2. Studies on transcription factors regulating erythropoiesis are often performed using human and murine cell lines that represent, to some extent, erythroid cells at given stages of differentiation 3-5. However transformed cell lines can only partially mimic erythroid cells and most importantly they do not allow one to comprehensibly study the dynamic changes that occur as cells progress through many stages towards their final erythroid fate. Therefore, a current challenge remains the development of a protocol to obtain relatively homogenous populations of primary HSCs and erythroid cells at various stages of differentiation in quantities that are sufficient to perform genomics and proteomics experiments.
Here we describe an ex vivo cell culture protocol to induce erythroid differentiation from human hematopoietic stem/progenitor cells that have been isolated from either cord blood, bone marrow, or adult peripheral blood mobilized with G-CSF (leukapheresis). This culture system, initially developed by the Douay laboratory 6, uses cytokines and co-culture on mesenchymal cells to mimic the bone marrow microenvironment. Using this ex vivo differentiation protocol, we observe a strong amplification of erythroid progenitors, an induction of differentiation exclusively towards the erythroid lineage and a complete maturation to the stage of enucleated red blood cells. Thus, this system provides an opportunity to study the molecular mechanism of transcriptional regulation as hematopoietic stem cells progress along the erythroid lineage. 
Studying erythropoiesis at the transcriptional level also requires the ability to over-express or knockdown specific factors in primary erythroid cells. For this purpose, we use a lentivirus-mediated gene delivery system that allows for the efficient infection of both dividing and non-dividing cells 7. Here we show that we are able to efficiently knockdown the transcription factor TAL1 in primary human erythroid cells. In addition, GFP expression demonstrates an efficiency of lentiviral infection close to 90%. Thus, our protocol provides a highly useful system for characterization of the regulatory network of transcription factors that control erythropoiesis.

Lentiviral-mediated Knockdown During Ex Vivo Erythropoiesis of Human Hematopoietic Stem Cells

An ex vivo protocol to generate mature human red blood cells from hematopoietic stem/progenitors is described. Additionally we describe an efficient lentiviral-delivery method to knockdown the transcription factor TAL1 in primary erythroid cells. The efficiency of lentivirus mediated gene delivery is demonstrated using GFP expressing viruses.

Erythropoiesis is a commonly used model system to study cell differentiation. During erythropoiesis, pluripotent adult human hematopoietic stem cells ...

Determination of Mammalian Cell Counts, Cell Size and Cell Health Using the Moxi Z Mini Automated Cell Counter

Particle and cell counting is used for a variety of applications including routine cell culture, hematological analysis, and industrial controls1-5. A critical breakthrough in cell/particle counting technologies was the development of the Coulter technique by Wallace Coulter over 50 years ago. The technique involves the application of an electric field across a micron-sized aperture and hydrodynamically focusing single particles through the aperture. The resulting occlusion of the aperture by the particles yields a measurable change in electric impedance that can be directly and precisely correlated to cell size/volume. The recognition of the approach as the benchmark in cell/particle counting stems from the extraordinary precision and accuracy of its particle sizing and counts, particularly as compared to manual and imaging based technologies (accuracies on the order of 98% for Coulter counters versus 75-80% for manual and vision-based systems). This can be attributed to the fact that, unlike imaging-based approaches to cell counting, the Coulter Technique makes a true three-dimensional (3-D) measurement of cells/particles which dramatically reduces count interference from debris and clustering by calculating precise volumetric information about the cells/particles. Overall this provides a means for enumerating and sizing cells in a more accurate, less tedious, less time-consuming, and less subjective means than other counting techniques6.
Despite the prominence of the Coulter technique in cell counting, its widespread use in routine biological studies has been prohibitive due to the cost and size of traditional instruments. Although a less expensive Coulter-based instrument has been produced, it has limitations as compared to its more expensive counterparts in the correction for "coincidence events" in which two or more cells pass through the aperture and are measured simultaneously. Another limitation with existing Coulter technologies is the lack of metrics on the overall health of cell samples. Consequently, additional techniques must often be used in conjunction with Coulter counting to assess cell viability. This extends experimental setup time and cost since the traditional methods of viability assessment require cell staining and/or use of expensive and cumbersome equipment such as a flow cytometer.
The Moxi Z mini automated cell counter, described here, is an ultra-small benchtop instrument that combines the accuracy of the Coulter Principle with a thin-film sensor technology to enable precise sizing and counting of particles ranging from 3-25 microns, depending on the cell counting cassette used. The M type cassette can be used to count particles from with average diameters of 4 - 25 microns (dynamic range 2 - 34 microns), and the Type S cassette can be used to count particles with and average diameter of 3 - 20 microns (dynamic range 2 - 26 microns). Since the system uses a volumetric measurement method, the 4-25 microns corresponds to a cell volume range of 34 - 8,180 fL and the 3 - 20 microns corresponds to a cell volume range of 14 - 4200 fL, which is relevant when non-spherical particles are being measured. To perform mammalian cell counts using the Moxi Z, the cells to be counted are first diluted with ORFLO or similar diluent. A cell counting cassette is inserted into the instrument, and the sample is loaded into the port of the cassette. Thousands of cells are pulled, single-file through a "Cell Sensing Zone" (CSZ) in the thin-film membrane over 8-15 seconds. Following the run, the instrument uses proprietary curve-fitting in conjunction with a proprietary software algorithm to provide coincidence event correction along with an assessment of overall culture health by determining the ratio of the number of cells in the population of interest to the total number of particles. The total particle counts include shrunken and broken down dead cells, as well as other debris and contaminants. The results are presented in histogram format with an automatic curve fit, with gates that can be adjusted manually as needed.
Ultimately, the Moxi Z enables counting with a precision and accuracy comparable to a Coulter Z2, the current gold standard, while providing additional culture health information. Furthermore it achieves these results in less time, with a smaller footprint, with significantly easier operation and maintenance, and at a fraction of the cost of comparable technologies.

The Moxi Z miniature automated cell counter is a novel instrument that combines the Coulter Principle with patented thin-film sensor technology and a proprietary software algorithm to perform sizing and counting of a broad size range of particles as well as to determine the overall health of monodisperse mammalian cell cultures. This protocol describes the use of this instrument for counting and assessing the health of cell cultures.

Particle and cell counting is used for a variety of applications including routine cell culture, hematological analysis, and industrial controls1-5. A ...

Cell Tracking Using Photoconvertible Proteins During Zebrafish Development

Embryogenesis is a dynamic process that is best studied by using techniques that allow the documentation of developmental changes in vivo. The use of genetically-encoded fluorescent proteins has proven a valuable strategy for elucidating dynamic morphogenetic processes as they occur in the intact organism. During the past decade, the development of photoactivatable and photoconvertible fluorescent proteins has opened the possibility to investigate the fate of discrete subpopulations of tagged proteins1. Unlike photoactivatable proteins, photoconvertible fluorescent proteins (PCFPs) are readily tracked and imaged in their native emission state prior to photoconversion, making it easier to identify and select regions by optical inspection. PCFPs, such as Kaede2, KikGR3, Dendra4 and EosFP5, can be shifted from green to red upon exposure to UV or blue light due to a His-Tyr-Gly tripeptide sequence which forms a green chromophore that can be photoconverted to a red one by a light-catalyzed &beta;-elimination and subsequent extension of a &pi;-conjugated system3. PCFPs and their monomeric variants are useful tools for tracking cells6-10 and studying protein dynamics11-14, respectively. During recent years, PCFPs have been expressed in different animal model, such as zebrafish6, chicken7,8 and mouse9,10 for cell fate tracking. Here we report a protocol for cell-specific photoconversion of PCFPs in the living zebrafish embryo and further tracking of photoconverted proteins at later developmental stages. This methodology allows studying, in a tissue-specific manner, cell biological events underlying morphogenesis in the zebrafish animal model.

Here, we present a method for the photoactivated switch of photoconvertible fluorescent proteins (PCFPs) in the living zebrafish embryo and further tracking of photoconverted protein at specific time points during development. This methodology allows monitoring of cell biological events underlying different developmental processes in a live vertebrate organism.

Embryogenesis is a dynamic process that is best studied by using techniques that allow the documentation of developmental changes in vivo. The use of ...

Blastomere Explants to Test for Cell Fate Commitment During Embryonic Development

Fate maps, constructed from lineage tracing all of the cells of an embryo, reveal which tissues descend from each cell of the embryo. Although fate maps are very useful for identifying the precursors of an organ and for elucidating the developmental path by which the descendant cells populate that organ in the normal embryo, they do not illustrate the full developmental potential of a precursor cell or identify the mechanisms by which its fate is determined. To test for cell fate commitment, one compares a cell's normal repertoire of descendants in the intact embryo (the fate map) with those expressed after an experimental manipulation. Is the cell's fate fixed (committed) regardless of the surrounding cellular environment, or is it influenced by external factors provided by its neighbors? Using the comprehensive fate maps of the Xenopus embryo, we describe how to identify, isolate and culture single cleavage stage precursors, called blastomeres. This approach allows one to assess whether these early cells are committed to the fate they acquire in their normal environment in the intact embryo, require interactions with their neighboring cells, or can be influenced to express alternate fates if exposed to other types of signals.

The fate of an individual embryonic cell can be influenced by inherited molecules and/or by signals from neighboring cells. Utilizing fate maps of the cleavage stage Xenopus embryo, single blastomeres can be identified for culture in isolation to assess the contributions of inherited molecules versus cell-cell interactions.

Fate maps, constructed from lineage tracing all of the cells of an embryo, reveal  which tissues descend from each cell of the embryo.  Although fate maps ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Comparison of Three Different Methods for Determining Cell Proliferation in Breast Cancer Cell Lines

Measuring cell proliferation can be performed by a number of different methods, each with varying levels of sensitivity, reproducibility and compatibility with high-throughput formatting. This protocol describes the use of three different methods for measuring cell proliferation in vitro including conventional hemocytometer counting chamber, a luminescence-based assay that utilizes the change in the metabolic activity of viable cells as a measure of the relative number of cells, and a multi-mode cell imager that measures cell number using a counting algorithm. Each method presents its own advantages and disadvantages for the measurement of cell proliferation, including time, cost and high-throughput compatibility. This protocol demonstrates that each method could accurately measure cell proliferation over time, and was sensitive to detect growth at differing cellular densities. Additionally, measurement of cell proliferation using a cell imager was able to provide further information such as morphology, confluence and allowed for a continual monitoring of cell proliferation over time. In conclusion, each method is capable of measuring cell proliferation, but the chosen method is user-dependent.

This protocol describes the use of three different methods for analyzing cell proliferation in breast cancer cell lines. This includes the use of conventional cell counting, luminescence-based cell viability, and cell counting through the use of a cell imager. Each offers advantages for the reproducible measurement of cell proliferation.

Measuring cell proliferation can be performed by a number of different methods, each with varying levels of sensitivity, reproducibility and compatibility ...

Live Cell Fluorescence Microscopy to Observe Essential Processes During Microbial Cell Growth

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of proteins which are essential for bacterial survival. A series of target-specific dyes can be used as probes to better understand these processes. Staining with lipophilic dyes enables the observation of membrane structure, visualization of lipid microdomains, and detection of membrane blebs. Use of fluorescent-d-amino acids (FDAAs) to probe the sites of peptidoglycan biosynthesis can indicate potential defects in cell wall biogenesis or cell growth patterning. Finally, nucleic acid stains can indicate possible defects in DNA replication or chromosome segregation. Cyanine DNA stains label living cells and are suitable for time-lapse microscopy enabling real-time observations of nucleoid morphology during cell growth. Protocols for cell labeling can be applied to protein depletion mutants to identify defects in membrane structure, cell wall biogenesis, or chromosome segregation. Furthermore, time-lapse microscopy can be used to monitor morphological changes as an essential protein is removed and can provide additional insights into protein function. For example, the depletion of essential cell division proteins results in filamentation or branching, whereas the depletion of cell growth proteins may cause cells to become shorter or rounder. Here, protocols for cell growth, target-specific labeling, and time-lapse microscopy are provided for the bacterial plant pathogen Agrobacterium tumefaciens. Together, target-specific dyes and time-lapse microscopy enable characterization of essential processes in A. tumefaciens. Finally, the protocols provided can be readily modified to probe essential processes in other bacteria.

Understanding the function of essential processes in bacteria is challenging. Fluorescence microscopy with target-specific dyes can provide key insights into microbial cell growth and cell cycle progression. Here, Agrobacterium tumefaciens is used as a model bacterium to highlight methods for live cell imaging for characterization of essential processes.

Core cellular processes such as DNA replication and segregation, protein synthesis, cell wall biosynthesis, and cell division rely on the function of ...

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

Pancreatic beta-cells respond to increasing blood glucose concentrations by secreting the hormone insulin. The dysfunction of beta-cells leads to hyperglycemia and severe, life-threatening consequences. Understanding how the beta-cells operate under physiological conditions and what genetic and environmental factors might cause their dysfunction could lead to better treatment options for diabetic patients. The ability to measure calcium levels in beta-cells serves as an important indicator of beta-cell function, as the influx of calcium ions triggers insulin release. Here we describe a protocol for monitoring the glucose-stimulated calcium influx in zebrafish beta-cells by using GCaMP6s, a genetically encoded sensor of calcium. The method allows monitoring the intracellular calcium dynamics with single-cell resolution in ex vivo mounted islets. The glucose-responsiveness of beta-cells within the same islet can be captured simultaneously under different glucose concentrations, which suggests the presence of functional heterogeneity among zebrafish beta-cells. Furthermore, the technique provides high temporal and spatial resolution, which reveals the oscillatory nature of the calcium influx upon glucose stimulation. Our approach opens the doors to use the zebrafish as a model to investigate the contribution of genetic and environmental factors to beta-cell function and dysfunction.

Beta-cell functionality is important for blood-glucose homeostasis, which is evaluated at single-cell resolution using a genetically encoded reporter for calcium influx.