The authors would like to thank Amanda Struckhoff for the initial help with the experiment. This work was supported by a grant from NIH 5RO1CA115706.

1. Preparation of Plate Coated with Collagen
<ol>
 <li>Dilute collagen to a final concentration of 10 &mu;g/ml in Opti-Mem media.</li>
 <li>Coat 6 well plates with collagen, by adding 1.5 ml from step 1 to each well. Incubate overnight at 4 &deg;C.</li>
 <li>On the following day, wash the plate 2 times with 1x Phosphate Buffered Saline (PBS) and store them in PBS.</li>
</ol>
2. Cell Preparation and Seeding on a 6-well Plate
Note: Use warm media and PBS to ensure optimal conditions for cell movement
<ol>
 <li>Grow MDA-MB-231 (an invasive breast cancer cell line) sparsely in Dulbecco's Modified Eagle Medium (DMEM) with 10% Fetal Bovine Serum (FBS). In this example, we use the MDA-MB-231; however, this method can be applied to any adherent cancerous and non cancerous cell lines.</li>
 <li>Wash cells twice with warm 1x PBS and collect by trypsinization. Check cells under a microscope to make sure that the cells are properly detached by trypsin treatment.</li>
 <li>Collect the cells by adding DMEM medium with 10% FBS, gently spin down the cells at 1000 g for 5 minutes. Resuspend the cell pellet gently in DMEM medium with 10% FBS.</li>
 <li>Break the cell pellet by pipetting up and down gently. It is important to break down cell clusters into individual cells as much as possible.</li>
 <li>Count the cells and dilute to a final concentration of 50,000 cells/ml in DMEM media with 10% FBS. Dispense 2 ml into each well.</li>
 <li>Incubate the plate at 37 &deg;C in a tissue culture incubator overnight, until starting the time-lapse microscopy.</li>
 <li>To ensure ample space for migration, create a wound using a pipette tip, wash the plate with 1X PBS to remove debris (formed as a result of wound) and add DMEM with 10% FBS. Check under the microscope for floating cells. If large numbers of cells are floating, repeat the washing procedure. Now, the plate is ready to be setup on the microscope for image acquisition.</li>
</ol>
3. Microscope Setup and Automated Image Acquisition
There are several imaging software packages that control the microscope for automated image acquisition. Here, we use Slidebook 5.0 imaging software and automatic acquisitions are performed on a Olympus microscope (Olympus IX81), coupled to an EM-Hamamatsu C9100 camera, which is excellent for live-cell applications where speed of acquisition and minimal phototoxicity is paramount. The system includes a stage top environmental control chamber (Neue Live Cell) and an automated XY stage controller (Prior Proscan). Images are acquired with a 10X UPLANFL Ph1/0.30 objective, in bright field mode, using a defined time interval. The following protocol describes the image acquisition.
<ol>
 <li>Turn on the microscope, camera, and live cell imaging apparatus. Position the plate over the microscopic stage. Cover the plate by the Neue Live Cell environmental control chamber, which is connected with a CO2 supply and temperature sensors. Set the thermostat at 37&deg;C and CO2 at 5%.</li>
 <li>SlideBook software allows fully automated objective selection, wavelength selection and easy switching between fluorescent and bright field techniques. In order to set up different parameters of focus control, open SlideBook software &gt;go to the menu bar &gt; select focus control, by clicking &gt; <img alt="Figure 0 Button 1" src="/files/ftp_upload/3585/3585fig0btn1.jpg" /> and select following parameters:
 <ol>
 <li>In the "Objective box," define the objective by selection from drop down window. Note that we are using a phase objective and have manually selected the appropriate phase ring in the condenser. If using a system with an automated condenser, this will be automatically set through the software program. Alternatively, if neither objective turret nor condenser are automated these should each be selected manually.</li>
 <li>In the "Filter Set" box:
 <ol>
 <li>Select settings to direct the light path to either the eyepiece or camera from the drop down box to see available filters ("Live" for eyepiece, "Widefield" for Camera in our system).</li>
 <li>Select the appropriate filter for brightfield illumination (in our set up, this option is labeled "DIC" which represents an open position for the brightfield light path).</li>
 <li>Select the light source by clicking on "Open Bright" to open the bright field shutter.</li>
 </ol>
 </li>
 </ol>
 </li>
 <li>To view the sample under the eye piece, select the bright field filter (position "6" in our system) on the turret. After initially viewing the sample through the eyepiece to find appropriate fields and bring the sample into focus (steps 3.2.1 to 3.2.2.3). Move the filter turret to change the light path to the camera for image capture (position "1" in our system).</li>
 <li>The automated stage allows the selection of different XY coordinates to capture multiple fields of interest during image acquisition. In SlideBook software, go to the focus control, select "XY", select different positions though the eye piece, then click "set point" lock in each location for image capture (Figure 1).</li>
 <li>Fine tune the focus of the camera image that is viewed on the screen. This may be accomplished by either using the manual controls on the microscope or the computer controls, which can adjust the focus in much finer increments.</li>
 <li>Using the controls in the "Focus Controls" window, adjust the z position of the stage. For best results, focus on flat cellular protrusions near the contact with the plate.</li>
 <li>To aid with focusing use the slider to adjust the exposure times to be in an appropriate range for focusing. If multiple XY positions have been selected, adjust the focus for each individual position: Under focus control &gt; XY&gt; visit point. Note: this is done to make sure, that fields are correctly focused for an optimal snapshot.</li>
 <li>Use the software to set up capture window parameters. In SlideBook, open the capture by selecting <img alt="Figure 0 Button 2" src="/files/ftp_upload/3585/3585fig0btn2.jpg" /> from the menu bar. See Figure 2. We then check the following parameters.
 <ol>
 <li>Check Capture type &gt; as time lapse.</li>
 <li>Check Filter set: Wide field: DIC (for bright field image).</li>
 <li>"Time lapse capture" is the number of time points that will be captured (it can vary, depending on the cell type). Duration is the total length of time for the experiment. Units of time [in milliseconds (ms), seconds (s), minutes (m) or hours (h)] can be selected from the dropdown menu.</li>
 <li>"Interval" is the delay between the beginning of one timepoint and the beginning of the next timepoint. As you type in two of these values, the third field will be calculated automatically.</li>
 <li>Multiple XY capture: select "multipoint list" for more than 1 XY position.</li>
 <li>Name the file. In the given example, we have named it '231'.</li>
 </ol>
 </li>
 <li>To avoid unexpected shut down of computer due to multiple images, use the "spool option" to save the file. It is optional, however it is highly recommended. For this purpose, under capture control&gt; select Advanced&gt; Spool&gt;Capture to memory and save to spool file after each time points. Browse the location to save the slide book file (Figure 3). The generated movie can be imported later from the designated location. Steps of microscope setup has been summarized in <a href="https://www.jove.com/files/ftp_upload/3585/Figure_3.1_merged.pdf" target="_blank">file 3.1</a>.</li>
</ol>
4. Image Processing and Analysis of Migration: Calculation of Speed and Displacement
In this example, image processing is done by SlideBook5, 6 5.0 software. There are various other programs such as ImageJ7 for the image processing and analysis.
<ol>
 <li>Import the SlideBook files generated from images of different fields at various time points. Go to menu bar&gt; under Image&gt; Import&gt;Slide book spool&gt;select the desired file (Figure 4a).</li>
 <li>Here our goal is to track the movement of individual cells. It can be done using automated tracking or individual tracking. In the example, we have used manual tracking, since this facilitates ruling out artifacts easily, which is more difficult with automated tracking.</li>
 <li>Open the SlideBook file, in which we have to perform tracking. Under menu: Select Mask &gt; Particle Tracking Protocol &gt; Manual Particle Tracking Protocol (Figure 4b). A window will appear with start and end time points (Figure 4c).</li>
 <li>Begin to track movement of individual cells by clicking the center of each cell (nucleus). Track approximately 10 cells from each movie and at least three different movies should be selected from different locations so that approximately 30 cells will be analyzed from one experiment. Each experiment should be repeated three times. To allow an unbiased approach, select cells from different locations such as some cells from front, middle, corners etc. Click "OK", once tracking is done.</li>
 <li>Random migration can be analyzed by measuring changes in speed, velocity or displacement etc. To analyze the data, under menu &gt; select mask &gt; particle tracking&gt;particle tracking protocol. Select tracking parameters as described, below: (Figure 5a):
 <ol>
 <li>Track with - Select the parameter that you wish to track from the dropdown list. In this example, we have used, Center of Area (the coordinates of the center of the Object).</li>
 <li>Click "track and continue". Once tracking is complete, this will automatically advance to the next step.</li>
 </ol>
 </li>
 <li>Choose desired statistics for each path, such as displacement and average speed (see Figure 5b):
 <ol>
 <li>Average Speed - the total distance traveled divided by the time required to travel that distance.</li>
 <li>Total Displacement - the total distance traveled by the object.</li>
 <li>Click "Calculate and Continue" if you wish to progress to the next step, or simply select "Calculate" if you wish to view your statistics without going to the next step. A file will be generated which can be saved as text file and can later be exported to Excel.</li>
 </ol>
 </li>
 <li>In order to track objects using automated tracking protocol, first a Mask has to be created.
 <ol>
 <li>Select Mask &gt; Segment Image&gt; a segment window would appear &gt;select until threshold is achieved such that separating one object from others&gt; click apply&gt; ok (see Figure 5c). Step 4.5.1 is the same.</li>
 <li>Remove Objects smaller than - Check this box to selectively remove objects smaller than a given size. Enter the size in the edit field and choose either microns or pixels as the unit. This is used to remove artifacts frequently caused by debris.</li>
 <li>Minimum movement - enter the minimum number of time points that must be tracked in order to determine a path.</li>
 <li>Maximum Movement - enter the maximum movement that you expect from any given object from one time point to the next.</li>
 <li>Steps to analyze the data are the same as manual tracking (see the steps 4.5.2 to 4.6.2).</li>
 </ol>
 </li>
</ol>
5. Representative Results
An example of a random migration analysis as quantified in terms of speed and displacement (Figure 6). After exporting the file to excel, data is plotted using a box and whiskers plot. Statistical analysis comparing total displacement and average speed between test and control is performed using Mann- Whitney test. The test sample shows an increase in average speed as compared to the control. The test sample shows an increase in total displacement as compared to the control.
Movie of control and test: Control MDA- MB-231 and test MDA-MB-231 cells were seeded on collagen overnight. Time- lapse video microscopy was performed by Olympus microscope, coupled to an EM-Hamamatsu camera. Images are taken at an interval of 1 hour for a total of 18 hours during random migration. Please see Movie 1 and Movie 2 of random migration for control and test samples.
Microscope setup:
<img alt="Figure 1" src="/files/ftp_upload/3585/3585fig1.jpg" /> 
Figure 1. Setting up multiple points for image capture.
<img alt="Figure 2" src="/files/ftp_upload/3585/3585fig2.jpg" /> 
Figure 2. Steps depicting capture control. <a href="https://www.jove.com/files/ftp_upload/3585/3585fig2large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 3" src="/files/ftp_upload/3585/3585fig3.jpg" /> 
Figure 3. How to save multiple images.
Image processing and analysis of migration.
<img alt="Figure 4a" src="/files/ftp_upload/3585/3585fig4a.jpg" /> 
Figure 4a. Steps to import image for data analysis.
<img alt="Figure 4b" src="/files/ftp_upload/3585/3585fig4b.jpg" /> 
Figure 4b. Steps to analyze data using particle tracking protocol.
<img alt="Figure 4c" src="/files/ftp_upload/3585/3585fig4c.jpg" /> 
Figure 4c. Window depicting path length of tracked particles.
Steps of data analysis.
<img alt="Figure 5a" src="/files/ftp_upload/3585/3585fig5a.jpg" /> 
Figure 5a. Steps depicting parameters of particle tracking protocol.
<img alt="Figure 5b" src="/files/ftp_upload/3585/3585fig5b.jpg" /> 
Figure 5b. Selection of path statistics to be analyzed.
<img alt="Figure 5c" src="/files/ftp_upload/3585/3585fig5c.jpg" /> 
Figure 5c. Steps to create mask for automated tracking protocol.
<img alt="Figure 6" src="/files/ftp_upload/3585/3585fig6.jpg" /> 
Figure 6. Average speed (Figure 6a) and displacement (Figure 6b) is plotted in a whiskers plot. The test sample shows an increase of speed and displacement as compare to control.
Movies
Movie 1. Migration of control cells. <a href="https://www.jove.com/files/ftp_upload/3585/3585movie1.avi" target="_blank">Click here to view the movie</a>.
Movie 2. Migration of test cells. <a href="https://www.jove.com/files/ftp_upload/3585/3585movie2.avi" target="_blank">Click here to view the movie</a>.
Control MDA- MB-231 and test MDA-MB-231 cells were seeded sparsely on collagen for overnight. Time- lapse video microscopy was performed by Olympus microscope, coupled to an EM-Hamamatsu camera. Images are taken at an interval of 1hour for 18 hours during random migration. Please see the movies of random migration for control and test.

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We have nothing to disclose.

The real time random migration assay enables accurate, sensitive, analysis of cell migration parameters such as speed and displacement. This method is not restricted to end point measurement value, hence it is more quantitative. Since, cells are monitored in normal DMEM medium with serum; it reduces the deleterious effect of longer incubation in serum free media. It has been shown that the migration of cells depends on the formatting and breaking cell contacts with the substratum8, thus, this method could be u...

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>DMEM</td>
 <td>Thermo Scientific</td>
 <td>SH30243.01</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>FBS</td>
 <td>Gemini Bio- Products</td>
 <td>100-106</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Opti-Mem</td>
 <td>Invitrogen</td>
 <td>31985-070</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Collagen</td>
 <td>BD</td>
 <td>354231</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Microscope with Live Cell chamber &amp; Automated XY stage controller</td>
 <td>Olympus </td>
 <td>Olympus 1X81</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Environmental control chamber</td>
 <td>Neue</td>
 <td>Neue Live Cell Chamber </td>
 <td>Ours has a custom built rectangular glass plate top for appropriate optics through the top of the chamber. The rectangular shape accommodates multi-well plates used in many of our experiments. </td>
 </tr>
 <tr>
 <td>Automated XY stage</td>
 <td>Prior </td>
 <td>Prior Proscan</td>
 <td>This allows precise return to multiply selected XY positions during time lapse microscopy.</td>
 </tr>
 <tr>
 <td>Camera</td>
 <td>&nbsp;</td>
 <td>Hamamatsu EM camera C9100 </td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Software</td>
 <td>Typically purchased through microscope vendor such as Olympus</td>
 <td>Slide Book 5 </td>
 <td>&nbsp;</td>
 </tr>
 </tbody>
</table>

quantitative analysis of random migration of cells using time-lapse video microscopy

Cell migration is a dynamic process, which is important for embryonic development, tissue repair, immune system function, and tumor invasion 1, 2. During directional migration, cells move rapidly in response to an extracellular chemotactic signal, or in response to intrinsic cues 3 provided by the basic motility machinery. Random migration occurs when a cell possesses low intrinsic directionality, allowing the cells to explore their local environment.
Cell migration is a complex process, in the initial response cell undergoes polarization and extends protrusions in the direction of migration 2. Traditional methods to measure migration such as the Boyden chamber migration assay is an easy method to measure chemotaxis in vitro, which allows measuring migration as an end point result. However, this approach neither allows measurement of individual migration parameters, nor does it allow to visualization of morphological changes that cell undergoes during migration.
Here, we present a method that allows us to monitor migrating cells in real time using video - time lapse microscopy. Since cell migration and invasion are hallmarks of cancer, this method will be applicable in studying cancer cell migration and invasion in vitro. Random migration of platelets has been considered as one of the parameters of platelet function 4, hence this method could also be helpful in studying platelet functions. This assay has the advantage of being rapid, reliable, reproducible, and does not require optimization of cell numbers. In order to maintain physiologically suitable conditions for cells, the microscope is equipped with CO2 supply and temperature thermostat. Cell movement is monitored by taking pictures using a camera fitted to the microscope at regular intervals. Cell migration can be calculated by measuring average speed and average displacement, which is calculated by Slidebook software.

Watch this Scientific Journal Video about Quantitative Analysis of Random Migration of Cells Using Time-lapse Video Microscopy at JoVE.com

Quantitative Analysis of Random Migration of Cells Using Time-lapse Video Microscopy

This method allows monitoring of cells in real time and quantitative measurements of different cell migration parameters such as speed, displacement, and velocity. Unlike the traditional methods, this real time approach is not based on endpoint quantitative migration measurements; instead it allows monitoring and calculating different parameters continuously.

Cell migration is a dynamic process, which is important for embryonic development, tissue repair, immune system function, and tumor invasion 1, 2. During ...

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16.7K Views.  LSU School of Medicine. This method allows monitoring of cells in real time and quantitative measurements of different cell migration parameters such as speed, displacement, and velocity. Unlike the traditional methods, this real time approach is not based on endpoint quantitative migration measurements; instead it allows monitoring and calculating different parameters continuously.

Video: Quantitative Analysis of Random Migration of Cells Using Time-lapse Video Microscopy

Monitoring Actin Disassembly with Time-lapse Microscopy

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 ...

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes. The transparency of the 
embryos, the genetic resources, and the relative ease of transformation are qualities that make C. elegans an excellent model for early embryogenesis. Laser-based 
confocal microscopy and fluorescently labeled tags allow researchers to follow specific cellular structures and proteins in the developing embryo. For example, 
one can follow specific organelles, such as lysosomes or mitochondria, using fluorescently labeled dyes. These dyes can be delivered to the early embryo by means 
of microinjection into the adult gonad. Also, the localization of specific proteins can be followed using fluorescent protein tags. Examples are presented here 
demonstrating the use of a fluorescent lysosomal dye as well as fluorescently tagged histone and ubiquitin proteins. The labeled histone is used to visualize 
the DNA and thus identify the stage of the cell cycle. GFP-tagged ubiquitin reveals the dynamics of ubiquitinated vesicles in the early embryo. Observations 
of labeled lysosomes and GFP:: ubiquitin can be used to determine if there is colocalization between ubiquitinated vesicles and lysosomes. A technique for 
the microinjection of the lysosomal dye is presented. Techniques for generating transgenenic strains are presented elsewhere (1, 2). For imaging, embryos 
are cut out of adult hermaphrodite nematodes and mounted onto 2% agarose pads followed by time-lapse microscopy on a standard laser scanning confocal 
microscope or a spinning disk confocal microscope. This methodology provides for the high resolution visualization of early embryogenesis.

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

This article describes a technique for the visualization of the early events of embryogenesis in the nematode Caenorhabditis elegans.

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes.  The transparency of the 
embryos, the genetic ...

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

Use of Time Lapse Microscopy to Visualize Anoxia-induced Suspended Animation in C. elegans Embryos

Caenorhabdits elegans has been used extensively in the study of stress resistance, which is facilitated by the transparency of the adult and embryo stages as well as by the availability of genetic mutants and transgenic strains expressing a myriad of fusion proteins1-4. In addition, dynamic processes such as cell division can be viewed using fluorescently labeled reporter proteins. The study of mitosis can be facilitated through the use of time-lapse experiments in various systems including intact organisms; thus the early C. elegans embryo is well suited for this study. Presented here is a technique by which in vivo imaging of sub-cellular structures in response to anoxic (99.999% N2; &lt;2 ppm O2) stress is possible using a simple gas flow through setup on a high-powered microscope. A microincubation chamber is used in conjunction with nitrogen gas flow through and a spinning disc confocal microscope to create a controlled environment in which animals can be imaged in vivo. Using GFP-tagged gamma tubulin and histone, the dynamics and arrest of cell division can be monitored before, during and after exposure to an oxygen-deprived environment. The results of this technique are high resolution, detailed videos and images of cellular structures within blastomeres of embryos exposed to oxygen deprivation.

Use of Time Lapse Microscopy to Visualize Anoxia-induced Suspended Animation in C. elegans Embryos

Described here is an in vivo technique to image sub-cellular structures in animals exposed to anoxia using a gas flow through microincubation chamber in conjunction with a spinning disc confocal microscope. This method is straightforward and flexible enough to suit a variety of experimental parameters and model systems.

Caenorhabdits elegans has been used extensively in the study of stress resistance, which is facilitated by the transparency of the adult and embryo stages ...

Quantitative Analysis of Autophagy using Advanced 3D Fluorescence Microscopy

Prostate cancer is the leading form of malignancies among men in the U.S. While surgery carries a significant risk of impotence and incontinence, traditional chemotherapeutic approaches have been largely unsuccessful. Hormone therapy is effective at early stage, but often fails with the eventual development of hormone-refractory tumors. We have been interested in developing therapeutics targeting specific metabolic deficiency of tumor cells. We recently showed that prostate tumor cells specifically lack an enzyme (argininosuccinate synthase, or ASS) involved in the synthesis of the amino acid arginine1. This condition causes the tumor cells to become dependent on exogenous arginine, and they undergo metabolic stress when free arginine is depleted by arginine deiminase (ADI)1,10. Indeed, we have shown that human prostate cancer cells CWR22Rv1 are effectively killed by ADI with caspase-independent apoptosis and aggressive autophagy (or macroautophagy)1,2,3. Autophagy is an evolutionarily-conserved process that allows cells to metabolize unwanted proteins by lysosomal breakdown during nutritional starvation4,5. Although the essential components of this pathway are well-characterized6,7,8,9, many aspects of the molecular mechanism are still unclear - in particular, what is the role of autophagy in the death-response of prostate cancer cells after ADI treatment? In order to address this question, we required an experimental method to measure the level and extent of autophagic response in cells - and since there are no known molecular markers that can accurately track this process, we chose to develop an imaging-based approach, using quantitative 3D fluorescence microscopy11,12.
Using CWR22Rv1 cells specifically-labeled with fluorescent probes for autophagosomes and lysosomes, we show that 3D image stacks acquired with either widefield deconvolution microscopy (and later, with super-resolution, structured-illumination microscopy) can clearly capture the early stages of autophagy induction. With commercially available digital image analysis applications, we can readily obtain statistical information about autophagosome and lysosome number, size, distribution, and degree of colocalization from any imaged cell. This information allows us to precisely track the progress of autophagy in living cells and enables our continued investigation into the role of autophagy in cancer chemotherapy.

Autophagy is a ubiquitous process that enables cells to degrade and recycle proteins and organelles. We apply advanced fluorescence microscopy to visualize and quantify the small, but essential, physical changes associated with the induction of autophagy, including the formation and distribution of autophagosomes and lysosomes, and their fusion into autolysosomes.

Prostate cancer is the leading form of malignancies among men in the U.S. While surgery carries a significant risk of impotence and incontinence, ...

Visualization of Craniofacial Development in the sox10: kaede Transgenic Zebrafish Line Using Time-lapse Confocal Microscopy

Vertebrate palatogenesis is a highly choreographed and complex developmental process, which involves migration of cranial neural crest (CNC) cells, convergence and extension of facial prominences, and maturation of the craniofacial skeleton. To study the contribution of the cranial neural crest to specific regions of the zebrafish palate a sox10: kaede transgenic zebrafish line was generated. Sox10 provides lineage restriction of the kaede reporter protein to the neural crest, thereby making the cell labeling a more precise process than traditional dye or reporter mRNA injection. Kaede is a photo-convertible protein that turns from green to red after photo activation and makes it possible to follow cells precisely. The sox10: kaede transgenic line was used to perform lineage analysis to delineate CNC cell populations that give rise to maxillary versus mandibular elements and illustrate homology of facial prominences to amniotes. This protocol describes the steps to generate a live time-lapse video of a sox10: kaede zebrafish embryo. Development of the ethmoid plate will serve as a practical example. This protocol can be applied to making a time-lapse confocal recording of any kaede or similar photoconvertible reporter protein in transgenic zebrafish. Furthermore, it can be used to capture not only normal, but also abnormal development of craniofacial structures in the zebrafish mutants.

Visualization of experimental data has become a key element in presenting results to the scientific community. Generation of live time-lapse recording of growing embryos contributes to better presentation and understanding of complex developmental processes. This protocol is a step-by-step guide to cell labeling via photoconversion of kaede protein in zebrafish.

Vertebrate palatogenesis is a highly choreographed and complex developmental process, which involves migration of cranial neural crest (CNC) cells, ...

Time-Lapse Video Microscopy for Assessment of EYFP-Parkin Aggregation as a Marker for Cellular Mitophagy

Time-lapse video microscopy can be defined as the real time imaging of living cells. This technique relies on the collection of images at different time points. Time intervals can be set through a computer interface that controls the microscope-integrated camera. This kind of microscopy requires both the ability to acquire very rapid events and the signal generated by the observed cellular structure during these events. After the images have been collected, a movie of the entire experiment is assembled to show the dynamic of the molecular events of interest. Time-lapse video microscopy has a broad range of applications in the biomedical research field and is a powerful and unique tool for following the dynamics of the cellular events in real time. Through this technique, we can assess cellular events such as migration, division, signal transduction, growth, and death. Moreover, using fluorescent molecular probes we are able to mark specific molecules, such as DNA, RNA or proteins and follow them through their molecular pathways and functions. Time-lapse video microscopy has multiple advantages, the major one being the ability to collect data at the single-cell level, that make it a unique technology for investigation in the field of cell biology. However, time-lapse video microscopy has limitations that can interfere with the acquisition of high quality images. Images can be compromised by both external factors; temperature fluctuations, vibrations, humidity and internal factors; pH, cell motility. Herein, we describe a protocol for the dynamic acquisition of a specific protein, Parkin, fused with the enhanced yellow fluorescent protein (EYFP) in order to track the selective removal of damaged mitochondria, using a time-lapse video microscopy approach.

Herein, we describe in detail a time-lapse video microscopy approach to measuring the temporal recruitment of EYFP-Parkin during the selective removal of damaged mitochondria. This dynamic process of EYFP-Parkin-dependent removal of damaged mitochondria can be used as an indicator of cellular health under different experimental conditions.

Time-lapse video microscopy can be defined as the real time imaging of living cells. This technique relies on the collection of images at different time ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

Lens-free Video Microscopy for the Dynamic and Quantitative Analysis of Adherent Cell Culture

Here, we demonstrate that lens-free video microscopy enables us to simultaneously capture the kinetics of thousands of cells directly inside the incubator and that it is possible to monitor and quantify single cells along several cell cycles. We describe the full protocol used to monitor and quantify a HeLa cell culture for 2.7 days. First, cell culture acquisition is performed with a lens-free video microscope, and then the data is analyzed following a four-step process: multi-wavelength holographic reconstruction, cell-tracking, cell segmentation and cell division detection algorithms. As a result, we show that it is possible to gather a dataset featuring more than 10,000 cell cycle tracks and more than 2 x 106 cell morphological measurements.

Lens-free video microscopy enables us to monitor cell cultures directly inside the incubator. Here we describe the full protocol used to acquire and analyze a 2.7 day long acquisition of cultured HeLa cells, leading to a dataset of 2.2 x 106 measurements of individual cell morphology and 10584 cell cycle tracks.

Here, we demonstrate that lens-free video microscopy enables us to simultaneously capture the kinetics of thousands of cells directly inside the incubator ...