Reconstitution of Basic Mitotic Spindles in Spherical Emulsion Droplets

The overall goal of this procedure is to reconstitute mitotic spindle-like structures within the geometrical confinement of water in oil emulsion droplets. This method can help answer key questions in the mitotic spindle assembly field, such as how are spindles positioned and assembled, and what is the specific contribution of individual components to these processes? The main advantage of this technique is that we use a bottom-up approach to reconstitute spindle-like structures within the geometrical confinement that mimics the shape of a mitotic cell.

This method can also be used to study other processes that are dependent on cellular confinement. Mix 10 parts PDMS pre-polymer with one part of curing agent in a total mass of about 40 grams. Then, place the mix in a vacuum chamber for about 30 minutes to remove air bubbles.

Meanwhile, wrap aluminum foil around the mold to create a one centimeter deep cup with a four inch diameter. When ready, pour about 3/4 of the PDMS mix into the mold. Then, return the PDMS to a vacuum chamber for another 30 minutes.

Following degassing, cure the PDMS for an hour at 100 degrees Celsius. Meanwhile, prepare some glass slides for spin coating by dusting them off with compressed air. Then, spin coat the remaining PDMS mix onto the slides.

Cure these slides for an hour at 100 degrees Celsius to harden the PDMS. Next, gently strip off the PDMS using a razor blade, and punch the required holes out of the PDMS strips to make microfluidic chips. Now, corona treat the microfluidic chip and a PDMS-coated glass slide for a few seconds.

Finally, place a microfluidic chip onto each glass slide with the channels facing down, and bake the chips overnight at 100 degrees Celsius. First, using chloroform-washed glassware, prepare about 250 micrograms of chloroform-dissolved lipids. Carefully dry the lipid mixture with inert gas.

And then place it in a vacuum chamber for an hour. Then, dissolve the lipids in mineral oil and 2.5%surfactant to 0.5 milligrams per milliliter, which is about 500 microliters. To completely dissolve the lipids, sonicate the mix for 30 minutes at 40 kilohertz.

Next, spin coat PDMS onto cover glasses with a thickness that matches the microscope objective. Then, spin coat PDMS onto glass slides. Now, cure the PDMS-coated cover glasses and glass slides for an hour at 100 degrees Celsius.

Then, make the flow cells by closely spacing thin slices of laboratory sealing film onto the PDMS-coated glass slides. Position three millimeter strips about two millimeters apart. Then, cover the flow cells with a PDMS-coated cover slip, and seal the assembly by melting the film at 100 degrees Celsius for a quick minute.

Once heated, gently press the cover slip down and seal it with Valap. Next, prepare a PDMS cup for long-term imaging. Punch a four millimeter diameter hole in a three millimeter thick slice of PDMS.

Then, corona treat the PDMS slice and a PDMS-coated cover glass. Once treated, position them on top of each other, and bake the assembly overnight at 100 degrees Celsius. Monitor the formation of droplets on an inverted brightfield microscope.

Connect the lipid oil phase to the pressure controller, and increase the pressure until a drop of oil exits from the peak tubing. Connect the peak tubing to inlet two of the microfluidics chip. Then, completely fill the microfluidic chips with the lipid oil phase from inlet two.

Next, introduce MRB-80 based water phase from inlet one. Control the droplet size by changing lipid oil phase and water phase pressures to create droplets with a diameter of about 15 microns. About 800 millibar for the lipid oil phase, and 200 millibar for the water phase, is a good starting point.

After obtaining the desired droplet size, completely fill the flow cell with droplets. These droplets are formed using small volumes of the water phase. This means that the microfluidic setup needs to run quickly in order not to lose the entire water phase before droplets of the correct size have been formed.

Once filled, carefully close the ends of the flow cell using Valap. If the droplets do not stop moving, then the seal isn’t complete, or air bubbles may have been introduced. For long-term imaging, transfer the droplets into a PDMS cup, and cover with a layer of oil lipid mixture.

Thaw the centrosomes at room temperature, and put them at 37 degrees Celsius for 20 minutes to ensure proper microtubule nucleation. While waiting, prepare the assay mix on ice. This should contain tubulin, GTP, an oxygen scavenger system, molecular force generators such as microtubule cross-linkers, ATP, and ATP regenerating system.

Once mixed, spin down the sample in the cooled Airfuge rotor at 30 psi for three minutes. Then, add the pre-heated centrosomes to the mix. Optimize the amount of centrosomes added to get one or two centrosomes per droplet.

Then, use this mixture to produce emulsion droplets as previously described. In order to recruit dynein to biotinylated lipids at the droplet cortex, include GFP-dynein TMR, and streptavidin, to the water phase. On a spinning disk confocal microscope, visualize microtubule growth after 30 minutes at 26 degrees Celsius.

While imaging, microtubule growth can be promoted by increasing the temperature to 28 or even 30 degrees Celsius. For Z-projections, take stacks with one micron intervals, which should require about 20 images per droplet. Go to the main camera panel, click on Edit Z, and set Z Step to 1.0.

Click on Next to store the settings. Next, set the maximum linear EM Gain by clicking on the Camera tab in the Acquisition panel, and sliding the EM Gain bar to 300. Then, set Exposure Time to 200 milliseconds in the same tab.

Click on Record to store the settings. For live imaging experiments, make Z Projections every two minutes for two hours by reducing the exposure times to about 100 milliseconds, and increasing the Z intervals to two microns. For live imaging experiments, use a PDMS cup instead of a regular flow cell for maximum immobilization of the sample.

Using the described protocols, aster formation was studied in water and oil emulsion droplets containing centrosomes. Initially, the centrosomes freely diffuse within the confined volumes. After about 20 to 30 minutes, the first microtubules become visible, and centrosome diffusion becomes restricted as microtubules grow against the cortex in all directions.

When the microtubules grow longer than half of the droplet’s diameter, the centrosomes get pushed to opposing boundaries, with microtubules growing along the droplet cortex. Without streptavidin, dymeine is diffuse within the droplet. However, with streptavidin, within about 10 minutes of droplet formation, dynein gets linked to the biotinylated lipids.

When viewing fluorescent tubulin in the presence of cortical dynein, it is clear that the centrosomes are centrally positioned. Whereas in the absence of dynein, the centrosomes get pushed to opposite sides of the droplet. This is probably due to dynein promoting microtubule catastrophes and cortical pulling forces, which results in centering of the asters.

After watching this video, you should have a good understanding of how to use microfluidic technologies to create spindle-like structures in spherical emulsion droplets. Once mastered, droplet formation and imaging can be done in two to three hours when performed properly. Using this procedure, other spindle assembly factors can be encapsulated in order to study their effect on spindle morphology and positioning.

Precautions such as wearing gloves should always be taken while using chloroform or PDMS.