This research was made possible by a National Science Foundation Career award (MCB-0746066, Characterization of Transcriptional Activators that Regulate Emergent Behavior) to R.D.W.
We are grateful to L. J. Shimkus, B. S. Goldman, G. Suen, M. Singer, L. G. Welch, K. A. Murphy, and H. G. Taylor for helpful discussions and comments on the manuscript.

Supplies needed:
<ul>
 <li>Klett meter</li>
 <li>Pipette and tips</li>
 <li>2.5 ml microcentrifuge tubes</li>
 <li>Microcentrifuge</li>
 <li>CTTYE media:
 <ul>
 <li>1.0% Casitone (Difco Laboratories), 0.5% yeast extract (Difco Laboratories),</li>
 <li>10.0 mM Tris-HCl (pH 8.0), 1.0 mM KH2PO4, 8.0 mM MgSO4</li>
 </ul>
 </li>
 <li>TPM media:
 <ul>
 <li>10.0 mM Tris-HCl (pH 8.0), 1.0 mM KH2PO4, 8.0 mM MgSO4</li>
 </ul>
 </li>
 <li>Agarose</li>
 <li>Water bath set to 55&deg;C</li>
 <li>Glass microscope slides</li>
 <li>Large glass coverslips</li>
 <li>2 x 2 cm, 0.5-mm-thick silicone rubber gasket (Grace Bio-Lab Inc.)</li>
 <li>Parafilm (or labeling tape)</li>
 <li>Forceps</li>
 <li>Small binder clips</li>
 <li>Micro-sampling pipette (Fisher)</li>
 <li>100 &mu;l glass disposable tip (Fisher)</li>
 <li>Kimwipes</li>
</ul>
Part 1: Cell Preparation
Start by creating a sterile environment.
<ul>
 <li>Clean workspace, don gloves, and light burner.</li>
</ul>
<ol>
 <li>Start an overnight culture of cells by inoculating into a flask containing CTTYE broth and incubated in the dark at 32&deg;C with vigorous swirling.</li>
 <li>Once the culture reached a density of 5 x 108 cells/ml, pipette 1 ml cells into a 2.5 ml microcentrifuge tube.</li>
 <li>Pellet cells by centrifugation for 2 min at 16,000 x g (or max speed).</li>
 <li>Decant and discard supernatant.</li>
 <li>Wash cell pellet with 1 ml TPM (salt-balanced, nutrient-free media).
 <ul>
 <li>re-suspend and vortex</li>
 </ul>
 </li>
 <li>Re-pellet cells by spinning for 2 min at 16,000 x g (or max speed).</li>
 <li>Decant and discard supernatant. 
 
 CRITICAL STEP 
 
 </li>
 <li>Completely re-suspend pellet in 100 &mu;l TPM media using a combination of pipetting and vortexing. 
 
 IMPORTANT: This step ensures that there are no clumps of cells left in the tube. This may take a while.</li>
 <li>Set the cells aside at room temperature while preparing the tracking assay plates.</li>
</ol>
Part 2: Agar Preparation
<ol>
 <li>Prepare 50 ml of both TPM (assay media) and CTTYE (nutritive disk) media.</li>
 <li>Add 0.5 g agarose to each (agarose is less diffractive than agar).</li>
 <li>Autoclave to sterilize.</li>
 <li>After sterilization, maintain the media at 55&deg;C in an incubator (or water bath) while constructing the tracking assay plate complexes.</li>
</ol>
Part 3: Nutritive Disk Construction
<ol>
 <li>Place a sterile 0.5-mm-thick silicone rubber gasket on top of a flame-sterilized glass microscope slide, forming a small well.</li>
 <li>Pipette ~300 &mu;l of the 55&deg;C CTTYE media/agarose into the well created by the gasket on the slide and containing the nutritive disk. The CTTYE media/agarose should mound up.</li>
 <li>Place a flame sterilized slide (with no gasket) on top of the CTTYE media/agarose. 
 
 IMPORTANT: This slide must be set down at an angle to prevent bubbles from forming.</li>
 <li>Once the slide is in place, clamp the complex together with mini binder clips - one on each side.</li>
 <li>Place the clipped complex at 4&deg;C and allow the CTTYE media/agarose to set. This usually takes about 5 min.</li>
</ol>
Part 4: Tracking Assay Preparation - Set Up Plate Complexes
Assemble the components, prepare the slide complexes, place the nutritive disk, pour the TPM media/agarose, separate and dry plate complexes, plate cells, and assemble tracking assay plate complexes.
Assemble plate complex components
<ol>
 <li>For each complex, flame sterilized a glass microscope slide</li>
 <li>Place an autoclaved gasket on top of the slide and set aside (sterilized side up). This will form the bottom of the assay.</li>
 <li>Flame sterilize a glass cover slip and place it on top of a second glass microscope slide wrapped with labeling tape (or Parafilm) and set aside (sterilized side up). This is used as a support slide to keep the coverslip from cracking. This will form the top of the assay.</li>
 <li>Flame sterilize a third glass microscope slide and set aside (sterilized side up). This will be used to flatten the agarose. 
 
 IMPORTANT: Make sure the gaskets form a seal with the glass by pressing it down with forceps, otherwise the media/agarose may dry out.</li>
</ol>
Place nutritive disk
IMPORTANT: Steps 5 through 10 must be done to one slide complex at a time; otherwise the media/agarose could start to solidify resulting in poor movie quality.
<ol start="5">
 <li>Remove the nutritive disk complex form 4&deg;C (step 5 from part 3) and pry apart the slides exposing the now solidified CTTYE/agarose.</li>
 <li>Remove a 1 mm diameter 'nutritive disk' from the CTTYE media/agarose well described above using a micro-sampling pipette with a 100 &mu;l glass disposable tip and place it in the well created by the gasket on the cover slip (step 3 above).</li>
</ol>
Pour plates
CRITICAL STEP
<ol start="7">
 <li>Pipette ~300 &mu;l of the 55&deg;C TPM media/agarose into the well created by the gasket on the cover slip and containing the nutritive disk. The TPM media/agarose should mound up. 
 
 CRITICAL STEP</li>
 <li>Place the flame sterilized slide with no gasket (from step 3) on top of the TPM media/agarose. 
 
 IMPORTANT: This slide must be set down at an angle to prevent bubbles from forming. 
 
 </li>
 <li>Once the slide (from step 5) is in place, clamp the complex together with mini binder clips - one on each side.</li>
 <li>Place the clipped complex at 4&deg;C and allow the media/agarose to set. This usually takes about 5 min.</li>
</ol>
Separate and dry plates
IMPORTANT: To prevent the media/agarose from drying out, steps 11 through 20 should only be performed on one slide complex at a time. 
 
 IMPORTANT: For best results, steps 11 through 13 should be performed at 4&deg;C.
<ol start="11">
 <li>Once the media/agarose has set, remove the binder clips and squeeze the end of the complex to loosen the tape-wrapped slide.</li>
 <li>Remove the tape-wrapped slide and place it on the bench for further use. 
 
 CRITICAL STEP</li>
 <li>Using forceps as a wedge, separate the cover slip/gasket/media/agarose complex from the support slide (with no gasket) and discard support slide. 
 
 IMPORTANT: Do not use a prying motion to separate the cover slip/gasket complex. This could result in the cover slip breaking and/or the media/agarose sticking to the support slide.</li>
 <li>Place the cover slip/gasket/media/agarose complex on the tape-wrapped slide (gasket side up) and remove from 4&deg;C. Place this complex next to a burner to allow all visible moisture evaporate from the newly exposed media/agarose surface - no more than 1 min. 
 
 IMPORTANT: Do not let the media/agarose dry for too long as this could impact the M. xanthus swarm behavior.</li>
</ol>
Plate cells
CRITICAL STEP
<ol>
 <li>Once the excess moisture has evaporated, pipette 0.5 &mu;l of the concentrated cells (step 8 from part 1) onto the media/agarose approximately 1 mm away from the nutrient disk. 
 
 IMPORTANT: It is extremely important to make sure that the cells are deposited by moving the pipette straight down and then straight up. This ensures that the swarm will be circular. 
 
 IMPORTANT: It is extremely important not to touch the pipette tip to the media/agarose. This will make a depression on the surface and change the behavior of the M. xanthus swarm. 
 
 TIP: Depress the pipette tip before approaching the media/agarose. This will allow a drop of cells to appear on the bottom of the pipette tip and make it easier to deposit the cells.</li>
 <li>Once the cells are deposited, place the complex next to the burner to allow the cell spot to dry - no more than 20 sec.</li>
</ol>
Assemble assay
<ol>
 <li>Once the cell spot has dried, align the slide/gasket complex (from step 1) with the gasket on the cover slip/gasket/media/agarose/cell complex (from step 13) and gently press together to form a seal. 
 
 CRITICAL STEP</li>
 <li>Clean the surfaces of the slide and cover slip with a Kimwipe to remove the residue left by the Parafilm. The completed assay should resemble Figure 1.</li>
 <li>Place the completed slide complex on the heated stage maintained at 32&deg;C (slide down, cover slip up) immediately after wipe down (step 15) to prevent condensation form forming. If condensation has formed, let slide complex sit on the heated stage for several seconds before starting the image acquisition software.</li>
 <li>Choose the appropriate objective (2X, 4X, or 10X).</li>
</ol>
Part 5: Movie Preparation
CRITICAL STEP
<ol>
 <li>Turn on the camera and microscope, and check the light levels (make sure the light is not too intense). Start up the computer and open the image acquisition program.</li>
 <li>Click the live image window and focus the microscope. The image should appear similar to the one seen in Figure 3. Notice the uniform agar surface.</li>
 <li>Start acquiring images. 
 
 CRITICAL STEP</li>
 <li>Check the focus regularly during the first hour, as the media/agar tends to settle causing the focus to drift.</li>
 <li>Clean work area.</li>
 <li>Once image acquisition is complete, transfer acquired images for storage and break down the slide complex by soaking in 90% ethanol overnight.</li>
 <li>Autoclave gaskets for reuse.</li>
</ol>
<img src="/files/ftp_upload/2038/2038fig1.jpg" alt="Figure 1" /> 
 Figure 1. Cartoon illustration of the TM plate complex. (A) shows the basic TM plate complex in exploded view and cross section. (B) shows the use of larger gaskets.
<img src="/files/ftp_upload/2038/2038fig2.jpg" alt="Figure 2" /> 
 Figure 2. Microscope cluster. Each microscope node (inset) consists of a Nikon E400 microscope, objectives, a heated stage, an Insight camera, and a notebook computer. Each node is networked together and linked to a master controller computer. Two of the nodes are set up with fluorescence capabilities that consist of the EXFO light source and two Uniblitz shutters.
<img src="/files/ftp_upload/2038/2038fig3.jpg" alt="Figure 3" /> 
 Figure 3. A 20X image of tracking assay apparatus at time = 0. Scale bar, 1 mm.
<img src="/files/ftp_upload/2038/2038fig4.jpg" alt="Figure 4" /> 
 Figure 4. Adaptability of the TM plate complex. (A) a 100X image of M. xanthus gliding motility on CTTYE in 1.0% agar. (B) a 20X image of P. aeruginosa twitching motility. (C) a 20X image S. marcescens swarming motility. Both (B) and (C) were assayed on LB in 1.0% agar. (D) a 40X image of M. smegmatis sliding motility on LB in 0.5% agar. This image was captured using the alternative assay configuration seen in Fig 1B.
Video 1. A time-lapse video of an swarm subjected to the tracking assay.
Video 2. A time-lapse video of an M. xanthus swarm where 1% of the cells are fluorescently labeled. Alternating phase-contrast and fluorescent images were captured and overlaid to elucidate the position of fluorescent cells within the swarm. This video was captured on CTTYE in 1.0% agar.
Video 3. A time-lapse video of M. xanthus gliding motility. This video was captured on CTTYE in 1.0% agar.
Video 4. A time-lapse video of P. aeruginosa twitching motility. This video was captured on LB in 1.0% agar.
Video 5. A time-lapse video of S. marcescens swarming motility. This video was captured on LB in 1.0% agar.
Video 6. A time-lapse video of M. smegmatis sliding motility. This video was captured on LB in 0.5% agar using the alternative assay configuration seen in Fig 1B.

<ol>
	<li>Taylor, R. G., Welch, R. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemotaxis+as+an+emergent+property+of+a+swarm.">Chemotaxis as an emergent property of a swarm.</a> J Bacteriol. 190, 6811-6816 (2008).</li><li>Curtis, P. D., Taylor, R. G., Welch, R. D., Shimkets, L. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatial+Organization+of+Myxococcus+xanthus+During+Fruiting+Body+Formation.">Spatial Organization of Myxococcus xanthus During Fruiting Body Formation.</a> J Bacteriol. 189, 9126-9130 (2007).</li><li>Mignot, T., Merlie, J. P., Zusman, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulated+pole-to-pole+oscillations+of+a+bacterial+gliding+motility+protein.">Regulated pole-to-pole oscillations of a bacterial gliding motility protein.</a> Science. 310, 855-857 (2005).</li><li>Stoodley, P., Hall-Stoodley, L., Lappin-Scott, H. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detachment+surface+migration,+and+other+dynamic+behavior+in+bacterial+biofilms+revealed+by+digital+time-lapse+imaging.">Detachment surface migration, and other dynamic behavior in bacterial biofilms revealed by digital time-lapse imaging.</a> Methods Enzymol. 337, 306-319 (2001).</li><li>O'Toole, G., Kaplan, H. B., Kolter, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Biofilm+formation+as+microbial+development.">Biofilm formation as microbial development.</a> Annu Rev Microbiol. 54, 49-79 (2000).</li><li>O'Toole, G. A., Kolter, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Flagellar+and+twitching+motility+are+necessary+for+Pseudomonas+aeruginosa+biofilm+development.">Flagellar and twitching motility are necessary for Pseudomonas aeruginosa biofilm development.</a> Mol Microbiol. 30, 295-304 (1998).</li><li>Dalton, H. M., Poulsen, L. K., Halasz, P., Angles, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Substratum-induced+morphological+changes+in+a+marine+bacterium+and+their+relevance+to+biofilm+structure.">Substratum-induced morphological changes in a marine bacterium and their relevance to biofilm structure.</a> J Bacteriol. 176, 6900-6906 (1994).</li><li>Dworkin, M., Eide, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Myxococcus+xanthus+does+not+respond+chemotactically+to+moderate+concentration+gradients.">Myxococcus xanthus does not respond chemotactically to moderate concentration gradients.</a> J Bacteriol. 154, 437-442 (1983).</li><li>Dworkin, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tactic+behavior+of+Myxococcus+xanthus.">Tactic behavior of Myxococcus xanthus.</a> J Bacteriol. 154, 452-459 (1983).</li><li>Wu, S. S., Kaiser, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+of+expression+of+the+pilA+gene+in+Myxococcus+xanthus.">Regulation of expression of the pilA gene in Myxococcus xanthus.</a> J Bacteriol. 179, 7748-7758 (1997).</li><li>Bustamante, V. H., Mart&#237;nez-Flores, I., Vlamakis, H. C., Zusman, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+the+Frz+signal+transduction+system+of+Myxococcus+xanthus+shows+the+importance+of+the+conserved+C-terminal+region+of+the+cytoplasmic+chemoreceptor+FrzCD+in+sensing+signals.">Analysis of the Frz signal transduction system of Myxococcus xanthus shows the importance of the conserved C-terminal region of the cytoplasmic chemoreceptor FrzCD in sensing signals.</a> Mol Microbiol. 53, 1501-1513 (2004).</li></ol>

No conflicts of interest declared.

Time-lapse microcinematography (TM) has become a standard approach to studying prokaryotic motility 2-7. Traditionally, TM is performed by using filter paper wicks, thin agar pads, or agar slabs as substrates 8-11. These methods are adequate and cost effective when used to generate image sequences for general illustrations of bacterial movement. However, if image sequences must result in the generation of reproducible and quantitatively rigorous data, these methods are time consuming and somewhat unreliable. For example, variations in these techniques caused by human error could lead to a wide array of inaccuracies, from irregularities in the agar surface that could dramatically affect the behavior of the bacteria being studied to differences in the focal plane from one side of the assay substrate to the other. To solve these problems, we have designed a TM plate complex that is of sufficiently consistent quality to yield reproducible results by employing silicone gaskets that are both inexpensive and reusable (Fig. 1). In addition, the plate complex is highly modifiable and has proven to stay hydrated and sufficiently oxygenated for more than a week over a variety of media types and agar concentrations.
To facilitate the generation of time-lapse movies, 8 Nikon E400 microscopes were outfitted with 2X, 4X, and 10X objectives each. In addition, an Insight camera (Diagnostic Instruments, Inc.), and a heated stage (20/20 Technologies) was acquired for each microscope. Each camera was linked to a notebook computer on which the highly modifiable image acquisition software SPOT (Diagnostic Instruments, Inc.) had been installed. All the various components were assembled into nodes, each consisting of a microscope, three objectives, an Insight camera, a heated stage, and a controlling computer. Each of the eight nodes were networked together into a cluster and linked to a storage computer which is used to compile, analyze, and store the time-lapse videos generated by the cluster (Fig. 2). The storage computer was outfitted with a 1 terabyte RAID system, which is needed to store the vast amount of data generated by the cluster.
Once complete, the plate complex is placed on the heated stage of the microscope and tracking assay is initiated. Images are acquired at preset intervals that are determined based on the motility rate of the cells. For example, individual M. xanthus cells move at a rate of approximately one cell length per minute. If images are acquired of an M. xanthus swarm at that same rate (one image per minute), the resulting time-lapse video will appear smooth during playback. Once the image acquisition is complete, the images are compiled into a sequential matrix and played back at a sufficient rate that they appear to be moving (Video. 1). This time-lapse video can now be analyzed using a variety software packages.
Variations on Assay. The TM plate complex is highly modifiable and can be used to visualize many different microorganisms under a variety of conditions. Swarm visualization can be performed using phase-contrast microscopy (which records the behavior of the swarm as a single entity), fluorescence microscopy (which records the behavior of individual fluorescent cells that have been diluted in a non-fluorescent population), or a combination of both (which overlays alternating sequential phase-contrast and fluorescent images) (Video 2). The plate complex has been used to successfully generate time-lapse videos of several prokaryotic behaviors including M. xanthus gliding motility, Pseudomonas aeruginosa twitching motility, Seratia marcescens swarming motility, and the novel Mycobacterium smegmatis sliding motility (Fig. 4 and Videos 3-6).

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>1.0% Casitone</td>
<td>&nbsp;</td>
<td>Difco Laboratories</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>0.5% yeast extract</td>
<td>&nbsp;</td>
<td>Difco Laboratories</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Micro-sampling pipette</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>100 &mu;l glass disposable tip</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>2 x 2 cm, 0.5-mm-thick silicone rubber gasket</td>
<td>&nbsp;</td>
<td>Grace Bio-Lab Inc.</td>
<td>&nbsp;</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

recording multicellular behavior in myxococcus xanthus biofilms using time-lapse microcinematography

A swarm of the &delta;-proteobacterium Myxococcus xanthus contains millions of cells that act as a collective, coordinating movement through a series of signals to create complex, dynamic patterns as a response to environmental cues. These patterns are self-organizing and emergent; they cannot be predicted by observing the behavior of the individual cells. Using a time-lapse microcinematography tracking assay, we identified a distinct emergent pattern in M. xanthus called chemotaxis, defined as the directed movement of a swarm up a nutrient gradient toward its source 1.
In order to efficiently characterize chemotaxis via time-lapse microcinematography, we developed a highly modifiable plate complex (Figure 1) and constructed a cluster of 8 microscopes (Figure 2), each capable of capturing time-lapse videos. The assay is rigorous enough to allow consistent replication of quantifiable data, and the resulting videos allow us to observe and track subtle changes in swarm behavior. Once captured, the videos are transferred to an analysis/storage computer with enough memory to process and store thousands of videos. The flexibility of this setup has proven useful to several members of the M. xanthus community.

Watch this Scientific Journal Video about Recording Multicellular Behavior in Myxococcus xanthus Biofilms using Time-lapse Microcinematography at JoVE.com

Recording Multicellular Behavior in Myxococcus xanthus Biofilms using Time-lapse Microcinematography

To study Myxococcus xanthus swarm behavior, we have designed a time-lapse microcinematography protocol that can be modified for different assays. It employs standard growth conditions adapted for microscopy, and yields reproducible results by the use of inexpensive, reusable silicone gaskets. We have used this method to quantify multicellular chemotaxis.

Recording Multicellular Behavior in Myxococcus xanthus Biofilms using Time-lapse Microcinematography

A swarm of the &delta;-proteobacterium Myxococcus xanthus contains millions of cells that act as a collective, coordinating movement through a series of ...

recording-multicellular-behavior-myxococcus-xanthus-biofilms-using

Nanyang Technological University

Research

JoVE Journal

Biology

12.2K Views.  University of South Carolina (USC). To study Myxococcus xanthus swarm behavior, we have designed a time-lapse microcinematography protocol that can be modified for different assays. It employs standard growth conditions adapted for microscopy, and yields reproducible results by the use of inexpensive, reusable silicone gaskets. We have used this method to quantify multicellular chemotaxis.

Video: Recording Multicellular Behavior in Myxococcus xanthus Biofilms using Time-lapse Microcinematography

Monitoring Actin Disassembly with Time-lapse Microscopy

Okay, so what I'd like to do today is to show you how to construct a flow chamber, a fusion chamber for imaging on an upright microscope. What then I then do is I create two spaces using film. So what I do is I take the para film, I cut it into small strips.Next I'm going to, this will basically form the channels for the, the walls, and I dry them with a tilted air. I put the cover slip on top of the para, And here's the finished. Then I heat it on a heat block to melt the param and basically create a seal.So what this is, is the, the cover glass here, and then on top of the cover glass is two layers of phim as you see right here. And then on top of this, again, there's a, it's a 22 by two two mil meter cover slip. This one right here, which I pressed against the param to fall the seal.Okay, so the liquid goes in in one direction here and I can suck it out with a filter paper here in this direction. And I'll show you how to do that in a bit. So what I like to do is I like to put it in a humidified chamber, which is very simple.Take two kind of elevated strips and have a filter of paper underneath. And to keep it noisy as I, I put the cover, cover slip on the flow chamber on. Now the way you introduce a solution to the chamber is that you just put the pipet tip at the entrance of the chamber and you just split.Now what I do is I incubate the chamber with the lid closed for 10 minutes. What I usually do to suck the liquid out from the other side is to use it, get a filter paper and cut it into thin strips like this. And I take one of these thin strips and I simultaneously apply the solution into the entrance while putting the filter paper at the exit to suck the exchange, the solution.So you'll see the liquid flowing through the solution. And then I wait for five minutes for the mid close. Now what I'm ready to do is to actually put that the actin it and the actin, I actually have to polymerize, I polymerize inside the flow chamber and it's just the same procedures before.So because like this, and then now that the action is polymerized, I wash out all the un excess un polymerized action using a large volume of just buffer. So here I have about a hundred microliters and it's a few reaction chamber volumes worth of buffer. Just, you know, when the filter paper becomes saturated, I have to change the direction.Okay?And so I'm ready to do imaging with this. So perfusion chamber with actin attached to it. Now I take the slide, mount it under the microscope, and since I'm imaging with an oil objective, I'm gonna put some immersion oil on the top surface of the cover, grass touch, such so assess solution or protein or anything on acting inside.So what I do is I can actually exchange the solution side while I'm doing the time lapse imaging. As such.

Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo. Here we describe a method to mount zebrafish embryos for long-term imaging and demonstrate how to automate the capture of time-lapse images using a confocal microscope. We also describe a method to create controlled, precise damage to individual branches of peripheral sensory axons in zebrafish using the focused power of a femtosecond laser mounted on a two-photon microscope. The parameters for successful two-photon axotomy must be optimized for each microscope. We will demonstrate two-photon axotomy on both a custom built two-photon microscope and a Zeiss 510 confocal/two-photon to provide two examples.

Zebrafish trigeminal sensory neurons can be visualized in a transgenic line expressing GFP driven by a sensory neuron specific promoter 1. We have adapted this zebrafish trigeminal model to directly observe sensory axon regeneration in living zebrafish embryos. Embryos are anesthetized with tricaine and positioned within a drop of agarose as it solidifies. Immobilized embryos are sealed within an imaging chamber filled with phenylthiourea (PTU) Ringers. We have found that embryos can be continuously imaged in these chambers for 12-48 hours. A single confocal image is then captured to determine the desired site of axotomy. The region of interest is located on the two-photon microscope by imaging the sensory axons under low, non-damaging power. After zooming in on the desired site of axotomy, the power is increased and a single scan of that defined region is sufficient to sever the axon. Multiple location time-lapse imaging is then set up on a confocal microscope to directly observe axonal recovery from injury.

Here we describe a method for mounting zebrafish embryos for long-term imaging, two-photon imaging and tissue-damage techniques, and time-lapse confocal imaging.

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo.  ...

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 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.

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 ...

Quantitative Measurement of Invadopodia-mediated Extracellular Matrix Proteolysis in Single and Multicellular Contexts

Cellular invasion into local tissues is a process important in development and homeostasis. Malregulated invasion and subsequent cell movement is characteristic of multiple pathological processes, including inflammation, cardiovascular disease and tumor cell metastasis1. Focalized proteolytic degradation of extracellular matrix (ECM) components in the epithelial or endothelial basement membrane is a critical step in initiating cellular invasion. In tumor cells, extensive in vitro analysis has determined that ECM degradation is accomplished by ventral actin-rich membrane protrusive structures termed invadopodia2,3. Invadopodia form in close apposition to the ECM, where they moderate ECM breakdown through the action of matrix metalloproteinases (MMPs). The ability of tumor cells to form invadopodia directly correlates with the ability to invade into local stroma and associated vascular components3. 
Visualization of invadopodia-mediated ECM degradation of cells by fluorescent microscopy using dye-labeled matrix proteins coated onto glass coverslips has emerged as the most prevalent technique for evaluating the degree of matrix proteolysis and cellular invasive potential4,5. Here we describe a version of the standard method for generating fluorescently-labeled glass coverslips utilizing a commercially available Oregon Green-488 gelatin conjugate. This method is easily scaled to rapidly produce large numbers of coated coverslips. We show some of the common microscopic artifacts that are often encountered during this procedure and how these can be avoided. Finally, we describe standardized methods using readily available computer software to allow quantification of labeled gelatin matrix degradation mediated by individual cells and by entire cellular populations. The described procedures provide the ability to accurately and reproducibly monitor invadopodia activity, and can also serve as a platform for evaluating the efficacy of modulating protein expression or testing of anti-invasive compounds on extracellular matrix degradation in single and multicellular settings.

We describe the prototypical method for producing microscope coverslips coated with fluorescent gelatin for visualizing invadopodia-mediated matrix degradation. Computational techniques using available software are presented for quantifying the resultant levels of matrix proteolysis by single cells within a mixed population and for multicellular groups encompassing entire microscopic fields.

Cellular invasion into local tissues is a process important in development and homeostasis.  Malregulated invasion and subsequent cell movement is ...

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 ...

Acquiring Fluorescence Time-lapse Movies of Budding Yeast and Analyzing Single-cell Dynamics using GRAFTS

Fluorescence time-lapse microscopy has become a powerful tool in the study of many biological processes at the single-cell level. In particular, movies depicting the temporal dependence of gene expression provide insight into the dynamics of its regulation; however, there are many technical challenges to obtaining and analyzing fluorescence movies of single cells. We describe here a simple protocol using a commercially available microfluidic culture device to generate such data, and a MATLAB-based, graphical user interface (GUI) -based software package to quantify the fluorescence images. The software segments and tracks cells, enables the user to visually curate errors in the data, and automatically assigns lineage and division times. The GUI further analyzes the time series to produce whole cell traces as well as their first and second time derivatives. While the software was designed for S. cerevisiae, its modularity and versatility should allow it to serve as a platform for studying other cell types with few modifications.

We present a simple protocol to obtain fluorescence microscopy movies of growing yeast cells, and a GUI-based software package to extract single-cell time series data. The analysis includes automated lineage and division time assignment integrated with visual inspection and manual curation of tracked data.

Fluorescence time-lapse microscopy has become a powerful tool in the study of many biological processes at the single-cell level. In particular, movies ...

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, ...

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the function and regulation of protein folding was studied in vitro, in isolated tissue culture cells and in unicellular organisms. Recent studies have uncovered links between protein homeostasis (proteostasis), metabolism, development, aging, and temperature-sensing. These findings have led to the development of new tools for monitoring protein folding in the model metazoan organism Caenorhabditis elegans. In our laboratory, we combine behavioral assays, imaging and biochemical approaches using temperature-sensitive or naturally occurring metastable proteins as sensors of the folding environment to monitor protein misfolding. Behavioral assays that are associated with the misfolding of a specific protein provide a simple and powerful readout for protein folding, allowing for the fast screening of genes and conditions that modulate folding. Likewise, such misfolding can be associated with protein mislocalization in the cell. Monitoring protein localization can, therefore, highlight changes in cellular folding capacity occurring in different tissues, at various stages of development and in the face of changing conditions. Finally, using biochemical tools ex vivo, we can directly monitor protein stability and conformation. Thus, by combining behavioral assays, imaging and biochemical techniques, we are able to monitor protein misfolding at the resolution of the organism, the cell, and the protein, respectively.

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

To study the relationship between protein homeostasis, stress and aging, we monitored changes in protein folding by following protein dysfunction, protein localization in the cell and protein stability at the organismal, cellular and protein levels, using the genetically tractable metazoan Caenorhabditis elegans as a model system.

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the ...