My name is Ken Kotz. I am a postdoc here at the Bio MAs Resource Center affiliated with Massachusetts General Hospital. My main research project involves isolating neutrophils using an immuno affinity capture device with microfluidic structures.
We use this device at five different hospitals across the country to isolate RNA for downstream genomic analysis. Basically what you have is a master of SU eight photo resist on top of silicon. We're going to pour a two part elastomer A-P-D-M-S on top of that master and that PDMS will form a mold.
We remove the master mold from its case and place it into a Petri dish. The master is secured by tape on the bottom of a Petri dish. On the scale we weigh out a mixture of of PDMS, hardener and base resin.
The ratio is one part hardener to 10 parts resin. Typically, for our devices that have been cut out, we weigh out 20 to 30 grams of the resin and two to three grams respectively of the hardener. After pouring both the hardener and the resin into the small whey boat, we use a standard plastic fork to mix the two together.
You wanna make sure that you mix vigorously and fold the mixture on top of one another to ensure that the hardener is evenly distributed within the resin. Typically, we monitor how well the two have been mixed. By looking at the number of air bubbles, we try to get an even distribution of small air bubbles throughout the elastomer.
We then pour the PDMS on top of the SU eight master that's in the Petri dish. We place the master with the PDMS on top of it in a vacuum bell jar, and evacuate the chamber to degas the air that we've mixed into it. Typically, the degassing process takes 30 minutes to one hour.
Following degassing, we remove the devices from the vacuum chamber and place them in a, in a oven at 65 to 80 degrees c. Minimum cure time at the these temperatures is three to six hours. In our lab, we typically leave the devices in the oven overnight.
Typically, we use a number 11 surgical steel knife. We make straight cuts down about half a centimeter from the edge of the silicon master, and we cut around the edge of the master making a large disc. We peel the PDMS mold from the master and we place the feature side up in a clean Petri dish.
We take the liberated mold out of the Petri dish and we place it onto a cutting surface using either either a knife or a knife mounted onto a linear rail. We section the devices that are contained on the mold. Each section device is then placed back into the Petri dish for further processing.
Typically, to interface our devices with fluids, we will punch holes into the PDMS device. The device that we use to punch holes is a standard three mill syringe. At the tip of the three mill syringe is a blunt tip stainless steel needle.
Within the stainless steel needle is a wire that fits the inner diameter of the needle. That wire is attached to the plunger of the syringe By pulling back on the plunger and retracting that needle, you can punch a hole or punch a plug into the PDMS. Flip the PDMS device and eject it by pushing on the plunger.
The secret to punching devices is to keep the plunger as vertical as possible and not to rotate the punching device at all. When you punch through the PDMS, you lift the whole device, the whole PDMS device up with a pair of tweezers, flip it over and eject the plug with by pressing down on the plunger. You grab the plug with a pair of tweezers and dispose of it.
Then you retract the plunger again, flip the device back down and pull out the cutting device. You repeat this until you have all of your holes punched. So for the bonding process, we bond our PDMS devices to glass slides.
For our devices, we're going to non reversibly bond or or covalently attach the PDMS to glass slides. The way this is accomplished is by placing them into a, placing them into a plasma Asher and exposing them to a reactive oxygen plasma. So the steps in doing this are to take the PDMS device and place it onto the tray of the plasma Asher.
With the device features facing upwards, you can use any standard glass microscope slide. We use a slightly enlarged one and a half inch microscope slide to fit our device. You can use them straight out of the package.
We prefer to treat them in a piranha solution. This cleans off any organic contaminants that might be on the surface of the glass slide. After you place the device and the glass slide on top of the tray for the plasma Asher, you slide the tray into the plasma Asher and expose it to the reactive oxygen plasma.
Following completion of the program of the plasma Asher, we open the chamber and we remove the plate with our devices to the bench top. Using tweezers, we carefully lift up the PDMS device, being careful not to touch the bonding surfaces with our fingers. We flip the bonding side over onto the glass, onto the glass slide, and then we make sure that we've, that there are no air bubbles between the PDMS and the glass slide.
We place the PDMS device now bonded to the glass on a, on a hot plate for at 65 to 75 degrees C for 10 minutes to achieve a better chemical bond. So this part in, in this section, we are going to chemically modify the surfaces of the PDMS in the glass following bonding in the clean room, we're left with a surface A PMS surface that has reactive OL groups on, on the surface. We need this reactive surface is hydrophilic and will eventually diffuse away into the bulk of the PMS.
So surface chemistry is best accomplished immediately following plasma treatment, we're going to use a 5%solution of a me capto. It's a three me capto, propyl, trimeth oxy. It's a 5%solution by volume, and basically what we do is we inject it into the inlet and make sure that you completely fill the device without any air bubbles.
If any air bubbles are present, you push them out with tweezers. This, once the, you've injected the slan, you let it sit for 15 to 30 minutes. At room temperature, you typically, we inject a second volume of the slan 10 minutes into the reaction.
So after the cylin has reacted for approximately a half hour, we'll take a, we'll flush all the devices with three to four device volumes of pure ethanol. The excess that we eject will just wipe up with a chem wipe Following rinsing of the devices, we pick up the devices and place them onto a hot plate, which is set to a hundred degrees C, and we basically will allow the ethanol that's in there to evaporate. This helps a kneel the siling surface.
Once the device is dried, it can be stored in a desiccated for months or it can, you can take the device and immediately proceed to the next step. After the devices have cured, they've cured with a now a sine coating on both the glass and the PDMS surface. The sine is a a capto sine with the thiol group, which will react with our hetero bifunctional crosslinker, which is GMBS.
It's diluted in pure ethanol. It's water reactive, so you need to take care not to expose it to aqueous conditions. We inject it into the devices and we remove any air bubbles with tweezers as I'm doing here to make sure all the surfaces are wedded with this molecule.
We cover it and we let it react for a half hour. After the GMBS has reacted for a half an hour, we're going to flush the devices with de ionized water to remove any traces of ethanol that are in the device. And then we're gonna flow through neu travain, which is biotin binding protein.
The neu travain is then pre-diluted to a concentration of 10 micrograms per mil. If, if any air is accidentally injected into the device, the device will need to be re flushed with ethanol to effectively remove all the air ethanol wets, the device surfaces better and allows you to remove any air bubbles that were accidentally introduced into the device. And once the devices have been flushed with the ionized water, we'll typically fill the device with two to four device volumes of the diluted neutral evidence T travain solution.
The Nutt travain will react with the GMBS, which is immobilized to the surface through any primary means on the Nutt travain. This process can occur at room temperature for one hour. Typically, I place the devices in the cold room at four degrees C overnight.
Before we add the antibody, however, we want to flush out any, any Aden, which is unbound in in solution. And we do that by flowing through 1%solution BSA diluted in PBS. Because these chips are gonna be used for downstream genomic analysis, all solutions that we use are RNA free.
Typically, what we do is we will flush the device with four to five volumes of the 1%BSA solution. And then following that, we add our antibody. The antibody for this experiment is CD 66 B and antibodies specific to granulocytes in whole blood.
We use a concentration of anywhere from 10 to 25 micrograms per mil, and we'll inject the antibody. We'll inject 200 microliters of antibody into each device. We'll do two injections of a hundred microliters, one into each port of the device.
Typically, we'll inject a hundred microliters into one port, and we'll wait 30 minutes and inject an additional a hundred microliters into the opposite port.
