Mikrofabrizierten Post-Array-Detektoren (mPADs): Ein Ansatz zur Mechanical Forces Isolate

Hi, I'm Nathan Snia Decky and with me Today is Wes Ligan and Robbie Desai and Mike Yang. And we are from Chris Chen's lab here at the University of Pennsylvania. In this video we'll present our approach to measure cellular attraction forces using a micro fabricated array of posts.

Traction forces are generated through myosin and acton interactions and play an important role in our physiology. During development, they enable cells to move from one location to the next in order to form the early structures of our tissue. And they also help in the healing process cause they're necessary for the closure of wounds or the migration and crawling of leukocytes through our body.

But unfortunately these same forces can be detrimental to our health in the case of cancer metastasis or vascular growth towards a tumor. So there are very few methods which one can actually measure these traction forces in a quantitative way, and especially at the micro and nanoscale of individual cells. Most of the common methods to study cells in vitro have been with glass bottom dishes or polystyrene dishes, but these are very rigid substrates and they're not, they do not allow us to actually physically measure the traction forces and the defamations that a cell causes.

And so it's been very difficult to understand the underlying mechanisms of traction forces without some kind of physical readout. And so in Chris Chen's lab we have a technique that allows us to overcome these limitations. Our method is based on a vertical array of flexible cantilevers, the stiffness and size scale of which are such that individual cells will spread across multiple cantilevers and deflect them in the process.

The dimensions that we use most frequently are three micron in diameter, diameter cylindrical posts, 10 microns tall with a nine micron center to center spacing and a rectangular array. However, these dimensions can be varied to accommodate a variety of cellular studies. We begin with a rigid silicon master that you can see here and then translate those into flexible PDMS cantilevers that we see here.

Using this technique we can measure the deflections of individual cantilevers under a microscope and back out the cellular traction forces with the magnitude and direction required to generate such deflections. We've turned these substrates M pads for micro post array detectors and in this video we'll show you how to fabricate the M pads, beginning with the silicone master through the PDMS replication, and finally to the computation of modulation and cellular contractility. Hi, here we are in our labs clean room.

This is a dust free and light sensitive room that allows us to make the SU eight posts that will serve as the silicon master. We first will start with a silicon wafer and with this we'll coat it with SU eight photo resist SU eight's a negative resist and it's light sensitive. So when we shine light through this photo mask, it will cause light to pattern the micro posts and then form a three dimensional structure, which will then be the SU eight posts that we then cast in PDMS.

So after we've done all our processing, the wafer will look like this, which is where we have the micro pillar, the micro post structures already fabricated into the wafer. So to make the masters, we first start with a bare silicon wafer and this has been cleaned with ozone treatment for seven minutes and we've also dehydrated this wafer for 30 minutes at 120 degrees Celsius. We're gonna spin SU eight 2002 onto this to form a very thin base layer for the posts.

So because we're gonna be spinning SU eight, I'm gonna wear a gas mask, Reload the wafer into this holder here, dust off any dust that might be on there, place it on the spinner, we're gonna check to make sure it's centered And then we'll proceed to pour the SU eight 2002 on top of this. And then we'll go ahead and spin the wafer. We then place these onto the hot plate, the soft bake.

We do this for one minute afterwards. We then take it and transfer it to the 95 degree hot plate. And we do this for two minutes.

And when it's finished, we'll then take this now and go off to do the flood exposure in the mask aligner. Okay, so we've just spun SU eight, the first layer of SU eight on this wafer, and now we need to flood expose the entire wafer to create that first base layer for the micro posts. So we need to give this 70 millijoules of exposure.

So we need to then measure the actual intensity of the light in our mask aligner here. So we load the meter into the weight into the mask aligner. We then expose it with light.

This is UV light here. We load the silicon wafer onto the mask aligner, place it into the mask aligner. And we don't have a mask this time, so that way we can expose the entire film.

Let's put the shutter on and let's go ahead and dial up one minute of exposure and we're gonna go ahead and expose. So after one minute the system is done, we can then take the wafer out. And now we need to do the post-exposure bake.

So we've just exposed this wafer and we're going to now add heat to cause the cross-linking reaction for this first coating of SU eight. So we place it on a 65 degree hot plate for one minute and then we transfer to a 95 degree hot plate for two minutes. After the two minutes process, we then turn off the hot plate and allow the the wafer to cool back to room temperature, which takes about 30 minutes.

So we've spun our second layer of SU eight onto this wafer. And now we need to actually expose this through the photo mask. This time we're gonna be using a UV filter to block out that deep UV light.

This is a Hoya optical glass that you can get, but this time we need to again measure the intensity. So we're gonna use our light meter or we're gonna use the the filter on top to measure the intensity that's been filtered out. And then we're gonna load the mask into the mask aligner.

This switch creates vacuum that holds the mask in place. We'll then place it into the mask aligner. We will then place the filter on top and then load our wafer in.

We wanna make sure there's good contact, so we raise it up into contact and then we will then start the exposure After exposure, we'll take our wafer out of the mask aligner. So now we have areas that have been selectively exposed. We'll take this to the oven, to the hot Plates to do the post-exposure bake on it.

So we've just done The po, the final post-exposure bake, and now we're going to develop this wafer so we can actually get the three dimensional structures. What we're gonna do is we're gonna wash this, develop this in P-G-M-E-A, which will wash away the unexposed SU eight. So we first start in one dish of P-G-M-E-A and we sit the wafer in here for two minutes with mild agitation.

So I'm gonna let this go for two minutes. So we're gonna transfer to a second dish of P-G-M-E-A. This will remove away that saturated P-G-M-E-A with SU eight.

After 10 seconds we'll bring it into IPA. This washes away the P-G-M-E-A. We put it in here for 20 seconds, again with some mild agitation.

And then right now we have the freestanding post structures. So we're gonna put this into hexane, which has a low surface tension. We're gonna wash for five seconds in one and then two hexanes, and then we're ready to blow dry it.

So we've just developed the wafer and now we're going to place it on a hot plate and slowly ramp the temperature up to one 50. And what happens is this higher temperature further cross links the SU eight to make it more structurally rigid and we leave it on this hot plate overnight. So anywhere from 14 to 20 hours and afterwards we will then turn off the hot plate and allow it to cool back to room temperature again.

So what we need to do is we're gonna dice this wafer down to a smaller size. We basically want to cut this silicon wafer and create a small chip of just this inner pattern here. And it's gonna look something like this structure there where they're gonna use epoxy.

And we're gonna glue that chip onto a glass slide like so. And this provides a structural backing. So to dice a wafer, it's sort of an art, but what you can do is you basically use a diamond scribe and you start at just one edge.

And because of the crystal planes, you just put a small notch Now that has created a fracture, which I can then tap with this tool, We can peel this away and we can then do more cuts here until we get a small structure like this. And then you quickly place the silicon chip now onto the glass and mount it into place. And you let that set.

And so after you're done, you'll have a, you'll have a device that's similar to this. So we've mounted the silicon master onto this glass slide and now we're gonna give it a coating of floy, which will help in releasing this structure from PDMS later on. So we're gonna place the chip into this desiccate and then we're going to take a few droplets of this tri Chloro Siling place a few drops into the desiccate.

We do this inside the hood because this is, this has a byproduct of hydro hydrochloric gas, which is pretty harsh. Go ahead and close up the lid, attach this to vacuum and then we go ahead and let this sit from the vacuum on and we let this deposit overnight to get a nice coating over the entire silicon wafer. Hi, I'm Wesley Ligan.

I'm a graduate student in Christopher Chen's lab. At this point, Nate has showed you how to create a silicone master for our ED substrates. However, before we can use this tool to measure cellular contractility is necessary to first replicate the rigid photo structures using an elastomeric silicone polymer.

Our first task is to generate a negative template of the silicone master we fabricated earlier. We do this using a silicon elastomer called PDMS or Polymethyl Sloane to our PDMS polymer. We now add a one to 10 ratio of curing agent.

Once we have the solution weighed out, it's necessary to thoroughly mix the PDMS polymer and the cross leaking agent. This mixing obviously introduces a lot of air bubbles into our solution. So at this point, it's necessary to de cover it and degas it in the vacuum desiccate.

Once the PDMS is fully degas and all of the bubbles have risen to the surface and popped, we can remove it from the vacuum desiccate. We can now use this mixture of PDMS and cross-linking agent to generate a negative mold of our silicon master. And to do this, we use a pie tin and place our mounted silicon master into the center of it.

We then carefully pour the DGAs polymer solution into the dish over the top of the silicon wafer trying to prevent any additional bubbles from forming. Typically we try to pour the polymer approximately a quarter inch thick. You can see that this process inevitably introduces a few air bubbles into the mixture.

So now we set this dish aside for approximately 30 minutes to allow the bubbles to rise to the surface and pop. We can see that after letting it sit, there are still a few air bubbles in the surface of the PDMS. However, because these are at the top surface, we can now use a quick burst of nitrogen to pop any remaining bubbles.

In order to fully crosslink the PDMS and CROSSLINKER solution and generate a rigid negative template of our silicon master, we bake it at 110 degrees Celsius for 20 minutes. We can see that there is a thin layer of PDMS on top of the silicone substrate. And before we can peel the two apart, it's necessary to first cut away this layer.

Then we can graft this top layer and gently peel it away from the silicon substrate. At this point, we can invert and very slowly peel the negative silicon substrate, the negative PDMS substrate away from the silicone master. Okay, we do this by holding one end down and gently peeling away the PDMS substrate.

You can see the front advance as the substrate peels away from the silicone master. Now that we've generated A-P-D-M-S negative mold of our silicone master, we can cut this into four individual negative molds to make the final pad substrates. It's important at this point not to touch the top of the negative substrate with forceps or your gloves to avoid damaging the surface.

So at this point, we've generated four individual negative templates from the silicone master consisting of three micron diameter wells, nine microns deep. In order to regenerate the raised cantilever surfaces, we can cast again on these negative molds from a PDI PDMS utilizing PDMS. However, before we do that, it's necessary to first render these negative molds non-adhesive by coating with a fluoros.

We do this in a hundred watt plasma chamber at full power for one minute. After allowing the chamber to pump down return on the plasma, this generates an ion gas field which etches the PDMS top surface and renders it more adhesive to the fluoro Cy Lane molecules. After removing the negative Molds from the vacuum desiccate, you can see here what we are left with is an array of three micron by 10 micron deep wells in a non-adhesive PDMS layer.

In order to regenerate the vertical cantilevers, which we use for the mpad substrates, it's necessary to cast again off of this using our same PDMS mixture and that will leave us with a positive freestanding array of elastomeric cantilevers, which identically mirror the silicon master we fabricated earlier. The first step in this procedure is to transfer our non-adhesive negative molds onto a a mounting jig we made out of PDMS strips and a glass microscope slide. The reason for this rig will become clear in the next step.

At this point, we can deposit a small drop of our same 10 to one mixture of PDMS to crosslinker already Degas onto the top surface of each of our negative molds. Only a small amount of PDMS is needed so that it does not run onto the edge of the negative molds. At this point.

In order to more fully spread the PDMS across the top surface of the molds, we can grab one using forceps, invert it, and bring it into contact with the adjacent mold carefully spreading around the PDMS to fully cover the entire substrate. Be very careful in this step not to introduce further bubbles into the top PDMS solution can then repeat with the other two wells, other two molds. Now that we have a more uniform PDMS coating across the top surface, we can place this jig into a vacuum desiccate to further remove any bubbles that may be trapped into the PDMS film on top of our negative molds.

This process typically takes about 30 minutes. While while our negative molds are degassing, we can clean the 22 by 22 millimeter glass substrates, which will become the final mounting place for the pads. The first step in this process is to blow off any dust particles using compressed nitrogen.

We'll expose the glass cover slips to gas plasma, which will ash the surface and make it more adhesive to the PMS, which we'll stick to it in the next step. After permitting the top layer of PDMS to DGAs for approximately 30 minutes and plasma activating the glass cover slips, we can then remove our template and bring the top surface of the glass cover slips, which has been exposed to the gas plasma into contact with the PDMS layer on top of the negative molds. Typically it helps to bring one edge of the glass cover slip into contact with the PDMS first and then allow the rest of the glass cover slip to settle in a SH so that the PDMS spreads out in a sheetlike fashion.

This helps reduce any bubbles introduced into the PDMS film. At this point, we can gently press on the top surface of the glass cover slip with our forceps to push out any excess PDMS as well as nudge the cover slip up against the PDMS cross in the center of the template. This will prevent the cover slip from moving around while the PDMS is curing.

We repeat this for the other substrates. At this point, we can transfer the entire template into the oven and bake at 110 degrees Celsius for 22 hours. After allowing the entire assembly to cool to room temperature, we can then remove our substrates, invert them, and carefully peel away the negative mold from the glass cover slip.

It's important to do this slowly and uniformly in order to avoid crushing the posts or tearing them from the glass substrate. We can see that after peeling away the negative mold, we are left with an array of, of cantilevers that precisely mimics the Silicon Master only that now they're in, they're constructed from a flexible PDMS elastomer. Okay, so at this point we've walked you through how we generate the Silicon Master of our Emad substrates.

We've then, we've used a replica molding process to generate an exact replica using A-P-D-M-S elastomeric polymer so that the, the cantilevers are now flexible to allow the cell to spread and contract and deflect the cantilevers in the process. At this point, our substrates are ready for processing for cell culture. Now that we've seen how to fabricate and replicate molds to make the eds, we're going to do a few more steps in order to render the EDS functional so that we can culture ourselves and subsequently analyze traction forces.

But before we do that, we need to do three things. We need to render the tips of the posts adhesive to cells. Secondly, we need to fluorescently label the PDMS so as to aid with visualization and analysis of post deflections.

And thirdly, we need to render the sidewalls of the posts as well as the base of the posts non-adhesive to to cells so that traction forces are exerted only at the post tips. So we'll see how these steps proceed via micro contact printing based strategy in the subsequent minutes. But first, the next step is to clean excess P-D-S-P-D-M-S off the eds.

So like I mentioned, the first step is to take the EDS substrate, which West just showed you how to generate and trim em off. The excess PDMS that resulted from curing the glass cover slip against the negative mold and using a sharp razor blade trim. The excess PDMS, you do this by, by putting the razor blade vertically down onto the substrate at the edge of the pillars and then scraping away.

And you can use one finger to hold the cover slip in place so that the cover slip does not move. So what we end up with is an an M pad substrate that has been cleaned of debris and so it is easily accessible. So for the next step, we have to micro contact, print the MPAs to render the tips of the posts adhesive.

In order to do this, we first need to generate stamps, and this process is very easy. We simply mix PDMS in instead of a 10 to one ratio, a one to 30 to one ratio. We pour in a Petri dish so that the PDMS height is approximately one centimeter.

Let it degas. That is let the bubbles rise to the top of the PDMS and cure the cross-linking reaction in the oven at 60 degrees for one hour after removing the dish from the oven and letting it cool, I'm now ready to cut out the stamps. So again, with the same kind of razor blade, simply cut by pressing vertically downward to into the PDMS until You make contact with the bottom of the Petri dish.

Now the orientation of the stamp is something that we want to pay attention to. The stamp surface is going to be the side of the PDMS that was cast against the bottom of the Petri dish so that it remains flat. So in this large square of PDMS that I've cut out, the stamp surface is now face up.

Now this stamp is much bigger than the M pads, so I'm going to trim it down to the size of a single M pad again by cutting with the razor blade, because it is very difficult to tell the correct orientation of the stamp by eye, I'm going to put a little notch in what will be the backside of the stamp that is the non-face side just like this. And I can then remove the stamp from the larger slab. And I now have A-P-D-M-S stamp for the next step.

We want to clean the PDMS stamps. Since it's very possible that dust from the atmosphere has settled onto the PDMS stamps as I was cutting them. This is a process in which we put the desired number of PDMS stamps into this crystallization dish and fill the crystallization dish dish with 100%ethanol.

So just add enough ethanol to cover the face of the stamps, but the exact amount of ethanol added is not important. And then place the crystallization disc in a ator and sonicate the stamps for five minutes. After the five minutes is up, we will finish sterilizing the stamps and do the remainder of the micro contact printing that is functionalization of the M pads in a sterile tissue culture hood.

So now we are in the tissue culture hood where we can finish sterilizing the stamps. To do this, we take the stamps from the crystallization dish, dip them in a dish full of ethanol to rinse off any remaining debris, and finally dip them in a dish of water to rinse off the ethanol. We then with a stream of nitrogen, dry the stamps until there are no longer visible drops remaining.

So the next step, we have to ink the stamps with a protein solution. The protein solution that we're using is 50 micrograms per mil of fibrin in DEI water. Simply take some of the protein solution up in the pipette And place drops of the protein solution onto the stamp face.

You want to do this by placing little droplets on the edge of the stamp face. First, repeat this for all of the stamps that you will be using and I should have done this in advance. So the next step after you've placed dots on the edges of all the stamps is to connect those droplets and fill in the protein solution on the stamp face.

The reason we did this is because protein solution in the droplets and sos on underneath the droplets and renders the stamp surface hydrophilic, making it easier to spread the extracellular matrix protein. So again, repeat this for all of the stamps that you're using now that we spread the protein solution on the face of the stamps, the last step in inking the stamps is to make sure that the edges are covered. To do this, we simply angle a pipette and run it along stamp face so that you make contact with both the corner of the PDMS stamp and the protein solution.

And again, repeat this for all edges of all stamps that you are inking. Now that we finish inking the stamps, we set that stamps, we leave the stamps in the tissue culture hood for one hour in order to facilitate complete absorption of the protein onto the stamp face. So now that we've waited one hour for proteins to its store, absorb onto the stamp surface, now it's time to rinse them with water and blow dry them with nitrogen.

To do this, we take sterile water and pour the sterile water into the dish containing the stamps. Careful not to pour the water directly onto the stamp face and you can use a pipette if need be. The exact amount of water that you had is not important so long as you bring the level of the water above the protein solution.

After this, with sterile tweezers, take one stamp, dip it into a new dish of water and gently blow it dry with nitrogen. Again, you'll know it's dry when you see, when you no longer see droplets of protein solution on the stamp. And repeat this for all stamps that you are processing one at a time.

So now that we have rinsed the stamps dry, it is time to activate the mpad substrates by treating them with UV ozone. For seven minutes, I remove the Petri dish lid from the top, from the top of the dish. Because you, because polystyrene is UV absorbent, I then place the dish into the UV ozone treater, close the drawer, set the time for seven minutes and let it go.

So now we are back in the tissue culture hood and it is time to stamp the mpad substrates, which were just treated with UV ozone. To do this, lift our rinsed and dried stamps with forceps. You can handle the stamps by the sides, but do not touch the face of the stamp.

You want to adjust the stamp in the forceps so that the protein side is accessible. You then invert the forceps, keeping in mind that the protein side is based down and slowly bring the stamp into contact with the eds. The one to 30 stamps are pretty sticky, so you may have to use a second pair of tweezers in order to release the stamp from the forceps.

You can drop the stamp down from a height of about two millimeters to about eight millimeters onto the emeds. Then tilt the dish of the emeds up and press down gently with forceps in order to create contact between the stamp and the eds. You will know when contact is made in one of two ways.

The first is to look at the ED surface through the side of the stamp. This may be hard to see on camera, but by eye I can clearly see that the stamp is making contact with the surface by looking at the surface through the side of the stamp. The second way to ensure contact is to with the p, is to bring the Petri dish out of the tissue culture hood with the lid on and view it under an inverted microscope.

By visualizing interference patterns with the microscope, you should be able to tell when, if and when the stamp is in contact with the eds. Once you have determined contact, the stamp only needs to remain in contact with this, with the ed substrate for a couple seconds. So it is now time to remove the stamp.

To do this, fold Eds down with one pair of tweezers and in one smooth motion, peel the stamp off. Be sure to keep track of which eds you have stamped and which stamps you have previous you have already used. And repeat this procedure once per EMPAs for every substrate that you wish to use.

Now that we stamp the EMPAs, the next step is to sterilize them and then incubate them in a lipophilic dye called dye eye. Dai is a lipophilic molecule that is fluorescently tagged, so it diffuses into the PDMS over the course of timed that allows us to visualize the post deflections under the microscope. So what I'm going to do is take each individual empa substrate one at a time and immerse it in 100%ethanol, 70%ethanol DI water DI water DI water DI water.

And finally, a five microgram per mil solution of dii. The DII solution is fluorescent, so to minimize its exposure to ambient light and minimize its photo bleaching, we cover the dishes with aluminum foil while they incubate in the tissue culture hood for one hour. Now that we've waited one hour for the for the EM pads to incubate in the DAI solution, we can remove the aluminum foil and rinse out the DAI solution.

We rinse out the by doing three sequential immersions in sterilized water and finally, we immerse into a solution of 0.2%onic F1 27. That's 0.2%weight per volume During all immersion steps, it's important to minimize the time that the EM pads are exposed to the air. This is to prevent damage to the ED substrates and specifically prevent two posts from collapsing onto each other.

The 0.2%onic F1 27 is a tri block copolymer that is composed of hydrophilic and hydrophobic domains. The hydrophilic domain is known to resist protein absorption and therefore cell adhesion while the hydrophobic domain binds to the temper to the to the hydrophobic PDMS. Thus, when PDMS is soaked in F1 27, the F1 27 binds to the PDMS and repel cell adhesion.

After one hour of incubation in chronic F1 27 in an aluminum foil covered Petri dish, we can rinse out the F1 27 by sequential immersions in sterile water, sterile water and PBS. After the immersion in PBS and the product F1 27 has been rinsed out, we can then immerse substrates the eds into cell culture solution of choice. The cell culture solution that I just immersed the EMPAs in can contain serum and or other growth factors and or supplements such as penicillin and streptomycin.

All of these ingredients are completely compatible with the EMPAs treatments. Now the next step, once the eds have been immersed in cell culture solution is to seed the cells of choice onto the eds. For this ex particular experimental setup, I will be seeding bovine pulmonary aortic endothelial cells into three different dishes.

The reason why I'm using three different dishes is because I will treat two of the dishes with known tactility inhibitors to inhibit the deflections of the posts as we'll be able to assay for at a later time. So to seed the cells, I simply take my cell suspension, draw up the appropriate amount into the pipette and pipette dropwise into each dish. I add enough cells so that when I do the assay to measure traction forces, the cells exist as single cell cells on the eds.

The amount that you add will depend on the spread area as well as the growth rate of your particular cells. We typically seed between 1500 and 5, 000 cells per centimeter squared of substrate. After you have finished seeding the cells or editing the the cell suspension, dropwise to the culture rock the culture gently back and forth several times in orthogonal directions to mix the cell suspension into the culture dish.

Repeat this process for all substrates that you are using. Immediately after seeding the cells, place them in a 37 degree incubator at an appropriate level of carbon dioxide to buffer the pH and the cell culture media. Two hour, approximately two hours after seeding cells, it is appropriate to rinse the cells by transferring the substrates from one dish of media to a fresh dish of the same cell culture media.

The reason for this rinse is that most cells will have begun spreading and will have attained nearly their final spread area by two hours. By rinsing the cell, the the substrates, you are removing any floating cells from the cell culture medium. That way only cells that have attached to the EDS and are beginning spreading process will show up at the endpoint assay for traction forces.

One assay that we can perform with cells that have been cultured on MPAs is time-lapse imaging to quantitatively examine the dynamics of cellular contractility. In particular, that the dynamics of cellular contractility in response to soluble factors is interesting. Soluble factors such as biological compounds and synthetic compounds are known to increase, diminish, or have unknown effects on cellular contractility.

For this live cell experiment, we will be using this inverted epi fluorescence microscope. On the microscope stage, I have a small incubator box has been hooked up to a thermal regulator to maintain the temperature at 37 degrees Celsius with 5%CO2. We'll be imaging at high magnification with a 60 x plan APO objective.

Now we will go prepare the the impact substrate for imaging. The 60 x objective that we are using for imaging has a working distance of 200 microns, which is much thinner than the thickness of the tissue culture dish in which we seeded the cells earlier. Therefore, we cannot, we cannot image cells on the implants in the tissue culture dish.

To get around this problem, we use these glass bottom dishes. I will now transfer the substrate into the glass bottom dish upside down so that the cells and the tips of the posts are at the interface of the glass Bottom dish. Therefore, when this is placed on the microscope, the working distance between the cells and the objective is less than 200 microns.

Now we're ready to go put this on a microscope for imaging. I'll now open up the live cell incubator chamber, place the sample inside and replace the incubator chamber lid. At this point, I will scan the substrate for a suitable cell.

To image. The most efficient way to do this is to use a face phase one 10 x objective, which will allow me to see a, a wider field of the substrate and find a cell more quickly. My criteria for good cell is for a single cell.

It should not be touching any neighboring cells, and it also should be about average in size compared to all the other cells on the substrate, I found a suitable cell to look at. The next thing I will do is switch the objective to 60 x to look at the cell at high magnification. To do that, we defocus the objective, so it's to not bump the stage during rotation.

During rotating the turret open up the chamber, partially rotate the 60 x objective interview 60 x 60 x objective is a oil immersion lens, so we have to place a drop of immersion oil on the objective lens. Now we will finish rotating the turt to bring the 60 x into position. We will close the incubator chamber and refocus the objective to bring the cell back into focus.

Once I fine tune the position and focus of the cell, I will switch to the CCD camera so that to preview the cell on the computer screen and then start the time-lapse imaging. For this experiment, we'll be, I'll be collecting both phase and fluorescent images at an interval of one minute, so the time lapse imaging has started and I will let it run for 10 minutes or 10 frames, and that at that point I will add serum 10 minutes has now elapsed. At this time, I will open up the chamber and add serum to the dish to stimulate contractility in the cell.

The timer's still running, so I have to be quick in doing this Now, we will continue imaging the cell for 30 minutes to follow any changes in contractility. After we have finished time lapse imaging, the images can be sequenced into a movie shown. Here are two movies of the phase and fluorescence images.

On the left is a phase movie of a, of a mostly transparent cell outlined in black. The cell is spread across the tips of 20 posts, which are the circular structures arranged in a rectangular lattice. On the right is a fluorescent movie of the same cell, but only the posts are visible.

If I play back these movies, you will see that the posts attached to the cell are initially weak, def deflected, and deviate very little from the rectangular lattice. Towards the middle of the movie, I added serum to the media. The serum stimulates the cell to increase in contractility.

As the cell exerts greater traction forces on the underlying posts, the posts along the perimeter of the cell begin to deflect inward. In the fluorescent movie, you now see posts deviate significantly from their positions in the rectangular lattice. As I would describe later, the displacement of these post positions in these images can be used to analyze the traction forces of this cell.

Previously, we saw how we generated the mpad substrates and how we functionalize them in order to culture cells. Now you will have noticed that I placed the finished mpad substrates into three different cell culture dishes. The cells have been growing overnight, and now what I'm going to do is treat each individual cell culture dish with a different soluble agent that modulates contractility in a known way.

We can then measure in a quantitative way the traction forces that result After fixing the cells up 30 minutes. After adding the soluble factors 30 minutes after adding the contractility inhibitors, it is now time to fix the cells with paraform aldehyde to lock in the current state of traction forces. To do this, it is important to use paraform aldehyde containing calcium and magnesium, and we use paraform aldehyde at a volume percentage of 3.7%The method for immersing the substrates in paraform aldehyde, sorry, is the same as the method for immersing the substrates in this, in the different RINs during functionalization.

That is, it is important not to let the substrates dry out as this may lead to post collapse. After I transfer the, I've transferred the substrates, I let them incubate in the paraform aldehyde for 20 minutes. After the 20 minutes is up, we remove the substrates from the paraform aldehyde now and now that the cells have been fixed, we can process these samples for standard immuno staining.

To do so, we place the substrates face up on param and pipette on enough PBS to cover the substrate and keep it underneath the PBS. We then rinse a substrate three times with PBS to remove any paraform aldehyde that may be left in the culture. After rinsing the substrate three times with PBS, we can transfer the substrate for downstream immuno staining.

We do this by handling the substrate with a pair of tweezers and inverting on the first step in immuno staining, and that is a permeation agent. In this case, we're using 0.2%Triton X in PBS. However, the user can use whatever ization agent is most appropriate for their needs.

We repeat this for all our substrates and this is how we handle the substrates during the immuno stain process. Following immuno staining, the substrates are mounted onto a standard one inch by three inch glass slide, which can then be processed on any epi fluorescence microscope, be it inverted or upright. By processing the samples on a confocal microscope, we can focus on the bottom of the posts to obtain the un deflected that is null positions.

We can then focus up to the tips of the posts, snap another image of the posts, and, and by comparing the top versus the bottom image, we can measure the deflections of the posts and calculate the traction forces of the cells that, that the cells generated in order to produce such deflections. By doing this process for many different cells on several different conditions, we can then compare the traction forces generated in the different conditions. Now that we have collected images of both live and fixed cells or next task is to measure the deflection, to measure the traction forces in the cells.

To do this, to do this, we have written a computer program in matlab. These are the three images that we will use in this analysis. The, the first image is a phase image of the cell.

It is helpful for identifying the location of the cell. The second image is a fluorescent image of the tops of the posts. We use this image to locate the positions of the, of the posts at at their tip.

As you can see, some of these posts are deflected and deviate from the, the rectangular lattice in which most of the posts are located in such as this post. And this post. The third image we'll use is the base image, which captures the position of the posts at a, at the, at a focal plane that crosses through the, through their base.

At this level, the both deflected and un deflected posts should be located in their rest position, un, un deflected positions. In the rectangular lattice, you can see that the posts that were deflected in the second image at their base, they're in their FL position. This image is used to find the position of the post it, it's, it's under reflected position.

Next, we load the MATLAB program, which will prompt us to load these three images. Give the file a name. You see the phase image.

We will draw a box around the indicating where the cell is, so the program knows which posts to analyze. The program then performs in intensity, thresholding and ization of the cropped image to determine the OID coordinates of the posts. At the left are ized images of posts from the top image while the right are ized images of the same posts from the base image.

After obtaining the posts, OID coordinates, the program can then, can then determine the displacement vectors between the tip and base of each post. These post displacement vectors can be converted into force vectors by multiplying them by the spring constant of the post. We can analyze these cellular forces in a number of ways, such as average force per cell or average force per post.

One useful result is a quiver plot from which we can visualize the spatial distribution of forces. This is the cell that we just analyzed and you can see I've overlaid yellow arrows on the posts that correlate with the, the magnitude of the force and point in the direction of the force. By repeating this image analysis process for sequences images such as those that are required in our time lapse movie, we can generate a temporal sequence.

Here is a movie of the cell that we had stimulated with serum. As you can see, just as the serum was added, the cell increasing contractility and deflected the posts even more. In addition, we can quantify forces for greater numbers of cells that have been subjected to different conditions and then fixed stained as exemplified by these quiver plots showing a cell treated with a control compound versus cells treated with contractility inhibitors.

As you can. As you can see, the the force vectors on the, in on the cell in the control case are greater than the greater in length in magnitude than the force vectors. In the two cases where the contractility inhibitors were used, we, we've covered a lot of material material here to describe thepath substrate from micro fabrication to sample preparation, experimentation and analysis to demonstrate the versatile versatility of these substrates.

We've described two different assays, live cell imaging and fixed sample assay, and we've done analysis of cells obtained from both of these assays and shown quiver plots and quiver movies. With this powerful tool, one can do variety of spatial and temporal analysis of cell contractility. As you saw, this method drew on many different disciplines, including those from micro fabrication, basic cell biology, surface science, image analysis and processing, and many others.

And it's precisely because its method is grounded in several different engineering techniques that the parameters such as post geometry, soluble factors induced and methods to analyze post deflections can be customized depending on the user's needs. It is our hope that the visual demonstration of this method that we outlined in the past 20, 30 minutes can be used as a starting point for other and and deeper analyses into the regulation, functional consequences, and generation of cellular attraction forces.