This work was supported by the AFM-Telethon to E.V. (#21545) and by the Ospedale San Raffaele (OSR) Seed Grant to S.Z. (ZAMBRA5X1000). Dr. Jean-Yves Tinevez from the Image Analysis Hub of the Institut Pasteur is acknowledged for publicly sharing his &#34;Simple Tracker&#34; MATLAB routines.

All procedures involving animal subjects were approved by the San Raffaele Institutional Animal Care and Use Committee.
1. Dissection of mouse hindlimb muscles
<ol>
	<li>Sterilize tweezers and scissors (both straight and curved) by autoclaving.</li>
	<li>Prepare and filter (0.22 &#181;m) all the media (blocking medium, digestion medium, proliferating medium, and differentiating medium) before starting the experiment (see Table of Materials).</li>
	<li>Put 5 mL of phosphate-buffered saline (PBS) in a 35 mm Petri dish for muscle collection.</li>
	<li>Sacrifice the mouse by cervical dislocation or by using CO2 and sterilize the skin with ethanol. Remove the skin from the hindlimbs and upper limbs by using scissors and tweezers, to facilitate the isolation of muscles. 
	NOTE: Muscles can be isolated from neonatal or adult mice. Neonatal mice have an increased number of satellite cells compared to adult mice. However, the total number of satellite cells that can be isolated from a single adult mouse is sufficient to perform the experiment described below.</li>
	<li>Isolate tibialis, soleus, extensor digitorum longus (EDL), gastrocnemius, quadriceps, and triceps, and put them in the Petri dish containing PBS. Leave the dish on ice. Carefully remove tendons and fat with sterilized scissors and tweezers.</li>
</ol>
2. Isolation of primary myoblasts
NOTE: All the procedures for cell culture are done in sterile conditions.
<ol>
	<li>Remove the muscles from the PBS. Cut and mince the muscles, by using sterile curved scissors, until a uniform mass is obtained. Put the muscle pieces into a 50 mL tube.</li>
	<li>Add 10 mL of the digestion medium to the muscle pieces and incubate at 37 &#176;C for 30 min under strong agitation (250 min-1) in a water bath, to enzymatically digest the muscles.</li>
	<li>Add 10 mL of Dulbecco&#8217;s modified Eagle&#8217;s medium (DMEM) containing 10% fetal bovine serum (FBS), 1% glutamine, 1% penicillin/streptomycin, and 1% gentamicin (blocking medium) to stop the digestion.</li>
	<li>Centrifuge at 40 x g for 5 min and collect the supernatant in a 50 mL tube. Save the pellet on ice. 
	NOTE: After the centrifugation, the supernatant contains some satellite cells, blood cells, endothelial cells, and fibroblasts, while the pellet contains pieces of undigested muscle and connective tissue. To increase the number of isolated cells, the pellet must be subjected to additional steps of digestion as described below.</li>
	<li>Centrifuge the supernatant at 650 x g for 5 min. Discard the supernatant and resuspend the pellet in 1 mL of DMEM containing 10% FBS. Keep the resuspended pellet on ice.</li>
	<li>Repeat steps 2.2&#8211;2.5 2x for the rest of the pellet obtained in step 2.4 and add the subsequently resuspended pellets to the first resuspended pellet conserved on ice.</li>
	<li>Pass the resuspended pellets through a 70 &#181;m filter and then through a 40 &#181;m filter. Add 15 mL of DMEM with 10% FBS, and centrifuge at 650 x g for 5 min.</li>
	<li>Resuspend the cell pellet in 3 mL of red blood cell lysis buffer to deplete the red blood cells. Incubate it for 5 min at room temperature. Add 40 mL of PBS to stop the lysis of the red blood cells, and centrifuge at 650 x g for 5 min. Remove the supernatant and resuspend the pellet in 5 mL of DMEM with 10% FBS.</li>
	<li>As preplating step, plate the cells in 20 mL of proliferation medium in a 150 mm uncoated Petri dish. After 1 h of incubation in a cell culture incubator (37 &#176;C, 5% CO2), collect the medium. 
	NOTE: Fibroblasts, attaching to the plate, are then discarded, while satellite cells remain in suspension.</li>
	<li>Repeat step 2.9 3x and, at the last preplating step, collect the medium which contains the satellite cells in suspension.</li>
	<li>Centrifuge at 650 x g for 5 min.</li>
	<li>While the cells are centrifuging, coat two 150 mm Petri dishes with collagen (10 mL of a 0.1% solution in 0.1 M acetic acid). To do so, put the collagen on the plate, incubate for 5 min, and remove it. Let the plate dry for 30 min.</li>
	<li>Discard the supernatant obtained in step 2.11 and resuspend the cell pellet in 40 mL of proliferation medium (Table of Materials). Plate the cells on two collagen-coated 150 mm Petri dishes (20 mL per dish) and keep the cells in a cell culture incubator (37 &#176;C, 5% CO2, 5% O2) in proliferation medium for 2&#8211;3 days with 1 &#181;g/mL of doxycycline to induce H2B-GFP expression. 
	NOTE: This step allows the proliferation of myoblasts to obtain a higher number of cells for further assays. Cell density is maintained very low to avoid a spontaneous differentiation of myoblasts into myotubes.</li>
</ol>
3. Myoblast proliferation and differentiation assays
NOTE: When myoblast density is high, some cells initiate elongation, and then, it is necessary to split the cells. Generally, it is possible to split cells 2x&#8211;3x without affecting their phenotype.
<ol>
	<li>Discard the medium, wash the cells with 5 mL of PBS, add 2 mL of trypsin (1x), and incubate the plate at 37 &#176;C in a cell incubator for 5 min. Check under the microscope and, when round cells detach, add 5 mL of DMEM with 10% FBS to inactivate the trypsin. Count the number of cells (usually it is possible to obtain about 1 x 106 cells/mouse). 
	NOTE: Incubation with trypsin for 5 min is usually enough to detach the proliferating myoblasts.</li>
	<li>Subject the cells to one of the assays described below.
	<ol>
		<li>Proliferation assay 
		<ol>
			<li>Plate 50,000 cells/well in 1 mL/well of proliferation medium in a collagen-coated 12-well plate and incubate the plated cells in a cell culture incubator (37 &#176;C, 5% CO2). Fix and count the cells at 24, 48, or 72 h. 
			NOTE: The medium is replaced every 48 h to avoid nutrient exhaustion.</li>
		</ol>
		</li>
		<li>Differentiation assay 
		<ol>
			<li>Coat a 12-well plate with 350 &#181;L of differentiation medium containing basement membrane matrix (1:100). Incubate the plate at 37 &#176;C for 30 min and remove the medium.</li>
			<li>Plate 200,000 cells/well in the differentiation medium in the matrix-coated plate. Incubate the cells in a cell culture incubator (37 &#176;C, 5% CO2, 5% O2) until they adhere to the plate (2 h are generally enough for the cells to adhere and to start differentiation).</li>
			<li>To assess differentiation, perform staining for myosin heavy chain and nuclei as previously described11. Alternatively, perform live-imaging analyses as described below.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
4. Live-imaging of myoblast differentiation and nuclear tracking
<ol>
	<li>For live cell imaging, put the cells, obtained in section 3.2.2 and plated in a 12-well plate, under a confocal microscope, equipped with an incubation system to maintain the cells at 37 &#176;C in 5% CO2.</li>
	<li>Use a 20x dry objective (0.7 NA) to get fields of view with hundreds of cells, enough to monitor myofiber formation. H2B-GFP cells can be imaged with a low-intensity 488 nm Argon laser. 
	NOTE: An open pinhole ensures that the image depth (~10 &#181;m) contains the cell thickness so that cells are kept in focus throughout the experiment. Using a confocal microscope is advantageous since it improves the signal-to-noise ratio of the images, although wide-field microscopy could also be used. Myofiber formation can be monitored through the transmission channel.</li>
	<li>Acquire images of 16-bit and 1,024 x 1,024 pixels per frame every 6 min for 16 h. For each acquired position, generate a multiframe.tif file (see example in the Supplementary Files). 
	NOTE: In the present protocol, commercial software associated with the microscope (Table of Materials) has been used; use the appropriate software to achieve the same goal when using other microscopes.</li>
	<li>Download the software provided as a .zip supplementary file. 
	NOTE: The routines are scripts that must be run in MATLAB. The software is an adaptation of a published software12 optimized for the tracking of the nuclei during myotube formation. This software is divided into two parts: the first one identifies the nuclei in each frame by segmenting them, while the second one generates &#8220;tracks&#8221; (trajectories) of nuclei movement. The complete code for nuclear segmentation and tracking and a step-by-step guide to getting started are provided in the Supplementary Files.</li>
	<li>Extract the zip file and save it in a desired path on the personal computer (PC) in use. Perform all the below passages on a PC with the necessary software already installed. Note that the .zip file is composed of three folders as mentioned below. 
	NOTE: Segmentation Routines contains the functions to be run for the segmentation. Tracking Routines, instead, contains the functions required for the tracking. Example Segmentation Tracking provides the basic scripts and files to get acquainted with the system. The software used for segmentation and tracking of the nuclei runs properly in MATLAB, R2015a or later versions.</li>
	<li>To segment the nuclei from the .tif file, name the file, for example, NameFile.tif.
	<ol>
		<li>Open the Example Segmentation Tracking folder and click on DoSegmentation.m. This will open the command window in MATLAB.</li>
		<li>Check the Current Folder window on the left to see all the elements in the Example of Segmentation Tracking folder. Double-click on DoSegmentation.m. 
		NOTE: Detailed information on the modifications that have to be done in the script can be found in the StepbyStep.txt file. It is mandatory to modify the script so that the variable filenameinput is NameFile.tif and folderinput is the folder where the .tif file is contained.</li>
		<li>Run the script (by clicking on the green Run button). The result of the segmentation will be saved as a file.mat (SegmentedNameFile.mat), while the segmentation of the nuclei can be visualized in the output figure. 
		NOTE: Parameters for the segmentation, described in the script, can be modified if needed. The quality of the segmentation can be checked using the script CheckSegmentation.m (see the StepbyStep.txt file). Based on this, if the quality of the segmentation is poor (only a few nuclei detected), adjust the parameters of the segmentation, clearly indicated in the DoSegmentation.m script, and repeat the procedure.</li>
	</ol>
	</li>
	<li>To generate the tracks (i.e., the trajectories of each of the nuclei in time), using the nuclear positions obtained in the segmentation, run the script GenerateTracks.m in the same working folder. Be sure to add the proper segmentation file as an input, obtained before (see the StepbyStep.txt file for further information). 
	NOTE: As an output, the user will obtain a file TrackedNameFile.mat, with a matrix for the x-coordinates and another matrix for the y-coordinates of the generated tracks.</li>
	<li>Evaluate the quality of the tracks using CheckTracking.m (see the StepbyStep.txt file for further information). Use the output figure of this last passage to help visualize each nuclei position in time. Check on the left for green crosses that indicate each segmented nucleus. To select one specific nucleus, use the lower scrollbar. Note that the selected nucleus will be circled in red, while on the right, the trajectory of the selected cell can be followed in time (by using the upper scrollbar).</li>
</ol>

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A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cellular+dynamics+in+the+muscle+satellite+cell+niche.">Cellular dynamics in the muscle satellite cell niche.</a> EMBO reports. 14 (12), 1062-1072 (2013).</li><li>Mauro, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Satellite+cell+of+skeletal+muscle+fibers.">Satellite cell of skeletal muscle fibers.</a> The Journal of Biophysical and Biochemical Cytology. 9, 493-495 (1961).</li><li>Motohashi, N., Asakura, Y., Asakura, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation,+culture,+and+transplantation+of+muscle+satellite+cells.">Isolation, culture, and transplantation of muscle satellite cells.</a> Journal of Visualized Experiments. 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The authors have nothing to disclose.

Muscle fibers (myofibers) are multinucleated cells that are formed by the fusion of muscle precursors cells (myoblasts) derived from muscle stem cells (satellite cells) that undergo proliferation and differentiation2,3,4. To assess myoblast differentiation, the common procedure consists of culturing myoblasts in differentiating medium and fixing the cells at different time points to perform immunofluorescence staining for MyHC, ...

Skeletal muscle is the largest tissue in the human body, totaling 35%-40% of body mass1. Satellite cells are muscle stem cells, anatomically characterized by their position (juxtaposed to the plasma membrane, underneath the basal lamina of muscle fibers), that give rise to proliferating myoblasts (myogenic progenitor cells), which eventually differentiate and integrate into existing myofibers and/or fuse to form new myofibers2,3,4. Their discovery and the progress in the study of their biology has led to significant insights into muscle development and regeneration.
Protocols to isolate and differentiate myoblasts into myotubes have been developed many years ago and are still widely used to study skeletal muscle differentiation5,6,7. However, most of these methods represent static procedures that rely on the analysis of fixed cells and, consequently, do not allow scientists to fully explore highly dynamic processes, such as myoblast fusion and myofiber maturation. The most striking example is nuclear positioning, which is tightly regulated, with nuclei initially in the center of the myofiber and, then, located at the periphery after myofiber maturation8,9. Live imaging is the most appropriate technique to obtain further insights into such a peculiar phenomenon.
Here, we describe a method that enables scientists to record myoblast differentiation and myotube formation by time-lapse microscopy and to perform quantitative analyses from the automatic tracking of myoblast nuclei. This method provides a high-throughput quantitative characterization of nuclear dynamics and myoblast behavior during differentiation and fusion. The protocol is divided into four different parts, namely (1) the collection of muscles from the hindlimbs of mice, (2) the isolation of primary myoblasts that consists in mechanical and enzymatic digestion, (3) myoblast proliferation and differentiation, and (4) live imaging to track nuclei within the first 16 h of myoblast differentiation.
In the following procedure, myoblasts are isolated from H2B-GFP mice and treated with 1 &#181;g/mL of doxycycline to induce H2B-GFP expression, as previously described10. Alternatively, it is possible to isolate the myoblasts from other transgenic mice that express a fluorescent protein in the nucleus or to transfect the cells isolated from wild-type mice to express a fluorescent protein in the nucleus, as described by Pimentel et al.9.

<table><tbody><tr><td>chicken embryos extract</td><td>Seralab</td><td>CE-650-J</td><td></td></tr><tr><td>collagenase</td><td>Sigma</td><td>C9263-1G</td><td>125units/mg</td></tr><tr><td>collagen from calf skin</td><td>Sigma</td><td>C8919</td><td></td></tr><tr><td>dispase</td><td>Gibco</td><td>17105-041</td><td>1.78 units/mg</td></tr><tr><td>doxyciclin</td><td>Sigma</td><td>D1515</td><td></td></tr><tr><td>DMEM</td><td>Sigma</td><td>D5671</td><td></td></tr><tr><td>fetal bovine serum</td><td>Life technologies</td><td>10270106</td><td></td></tr><tr><td>gentamicin</td><td>Sigma</td><td>G1397</td><td></td></tr><tr><td>Horse serum</td><td>Invitrogen</td><td>16050-098</td><td></td></tr><tr><td>Hoechst</td><td>life technologies</td><td>33342</td><td></td></tr><tr><td>IMDM</td><td>Sigma</td><td>I3390</td><td></td></tr><tr><td>L-glutamine</td><td>Sigma</td><td>G7513</td><td></td></tr><tr><td>Matrigel</td><td>Corning</td><td>356231</td><td></td></tr><tr><td>penicillin-streptomycin</td><td>Sigma</td><td>P0781</td><td></td></tr><tr><td>red blood cells lysis medium</td><td>Biolegend</td><td>420301</td><td></td></tr><tr><td>&lt;strong&gt;&lt;u&gt;Digestion medium&lt;/u&gt;&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>collagenase</td><td>40 mg</td><td></td><td></td></tr><tr><td>dispase</td><td>70 mg</td><td></td><td></td></tr><tr><td>PBS</td><td>20 ml</td><td></td><td></td></tr><tr><td>filtered 0.22um</td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;&lt;u&gt;Blocking medium&lt;/u&gt;&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>DMEM</td><td></td><td></td><td></td></tr><tr><td>Fetal bovine serum</td><td>10%</td><td></td><td></td></tr><tr><td>L-glutammine</td><td>1%</td><td></td><td></td></tr><tr><td>penicillin-streptomycin</td><td>1%</td><td></td><td></td></tr><tr><td>gentamicin</td><td>1 &amp;permil;</td><td></td><td></td></tr><tr><td>filtered 0.22um</td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;&lt;u&gt;proliferation medium&lt;/u&gt;&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>IMDM</td><td></td><td></td><td></td></tr><tr><td>Fetal bovine serum</td><td>20%</td><td></td><td></td></tr><tr><td>L-glutammine</td><td>1%</td><td></td><td></td></tr><tr><td>penicillin-streptomycin</td><td>1%</td><td></td><td></td></tr><tr><td>gentamicin</td><td>1 &amp;permil;</td><td></td><td></td></tr><tr><td>chichen embryo extract</td><td>3%</td><td></td><td></td></tr><tr><td>filtered 0.22um</td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;&lt;u&gt;differentiation medium&lt;/u&gt;&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>IMDM</td><td></td><td></td><td></td></tr><tr><td>Horse serum</td><td>2%</td><td></td><td></td></tr><tr><td>L-glutammine</td><td>1%</td><td></td><td></td></tr><tr><td>penicillin-streptomycin</td><td>1%</td><td></td><td></td></tr><tr><td>gentamicin</td><td>1 &amp;permil;</td><td></td><td></td></tr><tr><td>chichen embryo extract</td><td>1%</td><td></td><td></td></tr><tr><td>filtered 0.22um</td><td></td><td></td><td></td></tr></tbody></table>

exploiting live imaging to track nuclei during myoblast differentiation and fusion

Nuclear positioning within cells is important for multiple cellular processes in development and regeneration. The most intriguing example of nuclear positioning occurs during skeletal muscle differentiation. Muscle fibers (myofibers) are multinucleated cells formed by the fusion of muscle precursor cells (myoblasts) derived from muscle stem cells (satellite cells) that undergo proliferation and differentiation. Correct nuclear positioning within myofibers is required for the proper muscle regeneration and function. The common procedure to assess myoblast differentiation and myofiber formation relies on fixed cells analyzed by immunofluorescence, which impedes the study of nuclear movement and cell behavior over time. Here, we describe a method for the analysis of myoblast differentiation and myofiber formation by live cell imaging. We provide a software for automated nuclear tracking to obtain a high-throughput quantitative characterization of nuclear dynamics and myoblast behavior (i.e., the trajectory) during differentiation and fusion.

To automatically follow nuclear movement during myoblast differentiation in live imaging, the nuclei should preferentially be fluorescently labeled. It is important to note that using DNA-intercalating molecules is not feasible because these molecules interfere with the proliferation and differentiation of primary myoblasts13. As an example, proliferation and differentiation have been analyzed in primary myoblasts cultured with or without Hoechst (<strong class="xf...

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Exploiting Live Imaging to Track Nuclei During Myoblast Differentiation and Fusion

Skeletal muscle differentiation is a highly dynamic process, which particularly relies on nuclear positioning. Here, we describe a method to track nuclei movements by live cell imaging during myoblast differentiation and myotube formation and to perform a quantitative characterization of nuclei dynamics by extracting information from automatic tracking.

Nuclear positioning within cells is important for multiple cellular processes in development and regeneration. The most intriguing example of nuclear ...

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8.0K Views.  Istituto di Ricovero e Cura a Carattere Scientifico (IRCCS) San Raffaele Scientific Institute. Skeletal muscle differentiation is a highly dynamic process, which particularly relies on nuclear positioning. Here, we describe a method to track nuclei movements by live cell imaging during myoblast differentiation and myotube formation and to perform a quantitative characterization of nuclei dynamics by extracting information from automatic tracking.

Video: Exploiting Live Imaging to Track Nuclei During Myoblast Differentiation and Fusion

In Vitro Differentiation of Mature Myofibers for Live Imaging

Skeletal muscles are composed of myofibers, the biggest cells in the mammalian body and one of the few syncytia. How the complex and evolutionarily conserved structures that compose it are assembled remains under investigation. Their size and physiological features often constrain manipulation and imaging applications. The culture of immortalized cell lines is widely used, but it can only replicate the early steps of differentiation.
Here, we describe a protocol that enables easy genetic manipulation of myofibers originating from primary mouse myoblasts. After one week of differentiation, the myofibers display contractility, aligned sarcomeres and triads, as well as peripheral nuclei. The entire differentiation process can be followed by live imaging or immunofluorescence. This system combines the advantages of the existing ex vivo and in vitro protocols. The possibility of easy and efficient transfection as well as the ease of access to all differentiation stages broadens the potential applications. Myofibers can subsequently be used not only to address relevant developmental and cell biology questions, but also to reproduce muscle disease phenotypes for clinical applications.

In Vitro Differentiation of Mature Myofibers for Live Imaging

Muscle cells are among the most complex eukaryotic cells. We present a protocol for the in vitro differentiation of highly mature myofibers that allows for genetic manipulation and clear imaging during all developmental stages.

Skeletal muscles are composed of myofibers, the biggest cells in the mammalian body and one of the few syncytia. How the complex and evolutionarily ...

Developmental Biology

Visualization of Cell Cycle Variations and Determination of Nucleation in Postnatal Cardiomyocytes

Cardiomyocytes are prone to variations of the cell cycle, such as endoreduplication (continuing rounds of DNA synthesis without karyokinesis and cytokinesis) and acytokinetic mitosis (karyokinesis but no cytokinesis). Such atypical cell cycle variations result in polyploid and multinucleated cells rather than in cell division. Therefore, to determine cardiac turnover and regeneration, it is of crucial importance to correctly identify cardiomyocyte nuclei, the number of nuclei per cell, and their cell cycle status. This is especially true for the use of nuclear markers for identifying cell cycle activity, such as thymidine analogues Ki-67, PCNA, or pHH3. Here, we present methods for recognizing cardiomyocytes and their nuclearity and for determining their cell cycle activity. We use two published transgenic systems: the Myh6-H2B-mCh transgenic mouse line, for the unequivocal identification of cardiomyocyte nuclei, and the CAG-eGFP-anillin mouse line, for distinguishing cell division from cell cycle variations. Combined together, these two systems ease the study of cardiac regeneration and plasticity.

To distinguish cell division from cell cycle variations in cardiomyocytes, we present protocols using two transgenic mouse lines: Myh6-H2B-mCh transgenic mice, for the unequivocal identification of cardiomyocyte nuclei, and CAG-eGFP-anillin mice, for distinguishing cell division from cell cycle variations.

Cardiomyocytes are prone to variations of the cell cycle, such as endoreduplication (continuing rounds of DNA synthesis without karyokinesis and ...

Molecular Imaging to Target Transplanted Muscle Progenitor Cells

Duchenne muscular dystrophy (DMD) is a severe genetic neuromuscular disorder that affects 1 in 3,500 boys, and is characterized by progressive muscle degeneration1, 2. In patients, the ability of resident muscle satellite cells (SCs) to regenerate damaged myofibers becomes increasingly inefficient4. Therefore, transplantation of muscle progenitor cells (MPCs)/myoblasts from healthy subjects is a promising therapeutic approach to DMD. A major limitation to the use of stem cell therapy, however, is a lack of reliable imaging technologies for long-term monitoring of implanted cells, and for evaluating its effectiveness. Here, we describe a non-invasive, real-time approach to evaluate the success of myoblast transplantation. This method takes advantage of a unified fusion reporter gene composed of genes (firefly luciferase [fluc], monomeric red fluorescent protein [mrfp] and sr39 thymidine kinase [sr39tk]) whose expression can be imaged with different imaging modalities9, 10. A variety of imaging modalities, including positron emission tomography (PET), single-photon emission computed tomography (SPECT), magnetic resonance imaging (MRI), optical imaging, and high frequency 3D-ultrasound are now available, each with unique advantages and limitations11. Bioluminescence imaging (BLI) studies, for example, have the advantage of being relatively low cost and high-throughput. It is for this reason that, in this study, we make use of the firefly luciferase (fluc) reporter gene sequence contained within the fusion gene and bioluminescence imaging (BLI) for the short-term localization of viable C2C12 myoblasts following implantation into a mouse model of DMD (muscular dystrophy on the X chromosome [mdx] mouse)12-14. Importantly, BLI provides us with a means to examine the kinetics of labeled MPCs post-implantation, and will be useful to track cells repeatedly over time and following migration. Our reporter gene approach further allows us to merge multiple imaging modalities in a single living subject; given the tomographic nature, fine spatial resolution and ability to scale up to larger animals and humans10,11, PET will form the basis of future work that we suggest may facilitate rapid translation of methods developed in cells to preclinical models and to clinical applications.

A non-invasive means to evaluate the success of myoblast transplantation is described. The method takes advantage of a unified fusion reporter gene composed of genes whose expression can be imaged with different imaging modalities. Here, we make use of a fluc reporter gene sequence to target cells via bioluminescence imaging.

Duchenne muscular dystrophy (DMD) is a severe genetic neuromuscular disorder that affects 1 in 3,500 boys, and is characterized by progressive muscle ...

Medicine

Isolation of Human Myoblasts, Assessment of Myogenic Differentiation, and Store-operated Calcium Entry Measurement

Satellite cells (SC) are muscle stem cells located between the plasma membrane of muscle fibers and the surrounding basal lamina. They are essential for muscle regeneration. Upon injury, which occurs frequently in skeletal muscles, SCs are activated. They proliferate as myoblasts and differentiate to repair muscle lesions. Among many events that take place during muscle differentiation, cytosolic Ca2+ signals are of great importance. These Ca2+ signals arise from Ca2+ release from internal Ca2+ stores, as well as from Ca2+ entry from the extracellular space, particularly the store-operated Ca2+ entry (SOCE). This paper describes a methodology used to obtain a pure population of human myoblasts from muscle samples collected after orthopedic surgery. The tissue is mechanically and enzymatically digested, and the cells are amplified and then sorted by flow cytometry according to the presence of specific membrane markers. Once obtained, human myoblasts are expanded and committed to differentiate by removing growth factors from the culture medium. The expression levels of specific transcription factors and in vitro immunofluorescence are used to assess the myogenic differentiation process in control conditions and after silencing proteins involved in Ca2+ signaling. Finally, we detail the use of Fura-2 as a ratiometric Ca2+ probe that provides reliable and reproducible measurements of SOCE.

Here, we describe a methodology to obtain a pure population of human myoblasts from adult muscle tissue. These cells are used to study in vitro skeletal muscle differentiation and, in particular, to study proteins involved in Ca2+ signaling.

Satellite cells (SC) are muscle stem cells located between the plasma membrane of muscle fibers and the surrounding basal lamina. They are essential for ...

High-resolution Live Imaging of Cell Behavior in the Developing Neuroepithelium

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central nervous system (CNS). Although much is known about the molecular mechanisms that pattern the spinal cord and elicit neuronal differentiation1, 2, we lack a deep understanding of these early events at the level of cell behavior. It is thus critical to study the behavior of neural progenitors in real time as they undergo neurogenesis.
In the past, real-time imaging of early embryonic tissue has been limited by cell/tissue viability in culture as well as the phototoxic effects of fluorescent imaging. Here we present a novel assay for imaging such tissue for long periods of time, utilizing a novel ex vivo slice culture protocol and wide-field fluorescence microscopy (Fig. 1). This approach achieves long-term time-lapse monitoring of chick embryonic spinal cord progenitor cells with high spatial and temporal resolution.
This assay may be modified to image a range of embryonic tissues3, 4 In addition to the observation of cellular and sub-cellular behaviors, the development of novel and highly sensitive reporters for gene activity (for example, Notch signaling5) makes this assay a powerful tool with which to understand how signaling regulates cell behavior during embryonic development.

Imaging embryonic tissue in real-time is challenging over long periods of time. Here we present an assay for monitoring cellular and sub-cellular changes in chick spinal cord for long periods with high spatial and temporal resolution. This technique can be adapted for other regions of the nervous system and developing embryo.

The embryonic spinal cord consists of cycling neural progenitor cells that give rise to a large percentage of the neuronal and glial cells of the central ...

Neuroscience

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Utilizing Combined Methodologies to Define the Role of Plasma Membrane Delivery During Axon Branching and Neuronal Morphogenesis

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular guidance cues like netrin-1 and results in dramatic increases to the surface area of the axonal plasma membrane. Netrin-1-dependent axon branching likely involves temporal and spatial control of plasma membrane expansion, the components of which are supplied through exocytic vesicle fusion. These fusion events are preceded by formation of SNARE complexes, comprising a v-SNARE, such as VAMP2 (vesicle-associated membrane protein 2), and plasma membrane t-SNAREs, syntaxin-1 and SNAP25 (synaptosomal-associated protein 25). Detailed herein isa multi-pronged approach used to examine the role of SNARE mediated exocytosis in axon branching. The strength of the combined approach is data acquisition at a range of spatial and temporal resolutions, spanning from the dynamics of single vesicle fusion events in individual neurons to SNARE complex formation and axon branching in populations of cultured neurons. This protocol takes advantage of established biochemical approaches to assay levels of endogenous SNARE complexes and Total Internal Reflection Fluorescence (TIRF) microscopy of cortical neurons expressing VAMP2 tagged with a pH-sensitive GFP (VAMP2-pHlourin) to identify netrin-1 dependent changes in exocytic activity in individual neurons. To elucidate the timing of netrin-1-dependent branching, time-lapse differential interference contrast (DIC) microscopy of single neurons over the order of hours is utilized. Fixed cell immunofluorescence paired with botulinum neurotoxins that cleave SNARE machinery and block exocytosis demonstrates that netrin-1 dependent axon branching requires SNARE-mediated exocytic activity.

Light microscopy techniques coupled with biochemical assays elucidate the involvement of SNARE-mediated exocytosis in netrin-dependent axon branching. This combination of techniques permits identification of molecular mechanisms controlling axon branching and cell shape change.

During neural development, growing axons extend to multiple synaptic partners by elaborating axonal branches. Axon branching is promoted by extracellular ...

Live Imaging Followed by Single Cell Tracking to Monitor Cell Biology and the Lineage Progression of Multiple Neural Populations

Understanding the mechanisms that control critical biological events of neural cell populations, such as proliferation, differentiation, or cell fate decisions, will be crucial to design therapeutic strategies for many diseases affecting the nervous system. Current methods to track cell populations rely on their final outcomes in still images and they generally fail to provide sufficient temporal resolution to identify behavioral features in single cells. Moreover, variations in cell death, behavioral heterogeneity within a cell population, dilution, spreading, or the low efficiency of the markers used to analyze cells are all important handicaps that will lead to incomplete or incorrect read-outs of the results. Conversely, performing live imaging and single cell tracking under appropriate conditions represents a powerful tool to monitor each of these events. Here, a time-lapse video-microscopy protocol, followed by post-processing, is described to track neural populations with single cell resolution, employing specific software. The methods described enable researchers to address essential questions regarding the cell biology and lineage progression of distinct neural populations.

A robust protocol to monitor neural populations by time-lapse video-microscopy followed by software-based post-processing is described. This method represents a powerful tool to identify biological events in a selected population during live imaging experiments.

Understanding the mechanisms that control critical biological events of neural cell populations, such as proliferation, differentiation, or cell fate ...

Exploiting Live Imaging to Track Nuclei During Myoblast Differentiation and Fusion

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Rozdziały w tym wideo

In Vitro Differentiation of Mature Myofibers for Live Imaging

Visualization of Cell Cycle Variations and Determination of Nucleation in Postnatal Cardiomyocytes

Molecular Imaging to Target Transplanted Muscle Progenitor Cells

Isolation of Human Myoblasts, Assessment of Myogenic Differentiation, and Store-operated Calcium Entry Measurement

High-resolution Live Imaging of Cell Behavior in the Developing Neuroepithelium

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Time-lapse Imaging of Mitosis After siRNA Transfection

Live Imaging of Drosophila Larval Neuroblasts

Utilizing Combined Methodologies to Define the Role of Plasma Membrane Delivery During Axon Branching and Neuronal Morphogenesis

Live Imaging Followed by Single Cell Tracking to Monitor Cell Biology and the Lineage Progression of Multiple Neural Populations