Hi there. I'm Mitre a Dunham, a professor at the University of Washington Yeast Resource Center in my lab. We're interested in studying genome evolution.
One of the tools we use to study this process is experimental evolution where we evolve cultures of yeast in the laboratory and then ask how their genomes change. One of the devices we use to study this is the Hemostat. The hemostat is essentially a culture vessel in which you can grow yeast cultures for many hundreds of generations months in the lab, and then ask how the cells adapted.
This hemostat here is a custom glass flow hemostat, and as you might imagine, these are expensive and take up a lot of space. So recently in my lab we developed the mini hemostat. The mini hemostat lets us do all of the things with the large volume hemostats, but for less money and in less space.
My graduate student, Aaron Miller, is going to take you on a tour of how to build and use the mini hemostat array. Hey everyone. I'm Aaron Miller, a graduate student in the Dunham Lab.
In this video series, I will show you how to make and use an array of mini stats. If you have any questions that are not answered in these videos, please refer to the minis stat manual. The mini stat chamber is made up of a 50 mil glass tube, stopped with a silicon.
Cork air is delivered to the chamber by way of a long spinal tap needle, which reaches to the bottom of the tube and delivers robust airflow. Sampling is controlled by a second needle, which routes the culture to collection chamber for analysis. The length of this needle determines the working culture volume.
A third and far shorter needle is used to add media to the culture chamber. This activity determines the dilution rate and directly controls the rate of cell growth. The 20 mil culture volume stands out as being small enough that it can be greatly multiplex and still fit in a relatively small space, and yet large enough that samples produced will be sufficient.
For D-N-A-R-N-A and metabolite characterization air is pumped from an aquarium pump into a gas washing bottle. This disperses the Airstream into small bubbles to hydrate the air, which is then routed to the culture chambers. These bubbles not only aerate the media, but they also work to keep the cells suspended and from clumping together, media is delivered into the mini stat from tubing that connects a media car with the media delivery needle.
On the minis stat, the flow rate is controlled by a multiplex peristaltic pump, which massages media through pump tubing that is part of the media line. When the culture level reaches the culture sampling needle, the positive pressure created by the airflow pushes the culture out through the effluent track and into culture collection chambers. The temperature of the inner growth chamber is regulated by an external heat source, in this case, a heat block, although a water bath or incubator would also work.
In this video, I present instructions on how to assemble an array of four mini stats. Four represents a basic unit of the mini stat and by repeated application of these instructions, you can scale up this approach to run as many as 32 mini stats on a given pump. The second video in the mini stat series provides complete designs and methods for running evolution in competition based experimental studies and mini stats, as well as instructions of how to clean up after you're done.
We'll start with cleaning and assembling the culture chambers to clean the tubes. We wash with approximately 10 mils of 90%ethanol and then with 10 mils of water. To physically remove contaminants, I also take three Kim wipes and scrub the inside of the tube.
Then I wash one last time to remove any lint left by the Kim wipe. The culture tubes are generally stored upside down when not in use. Next, we mark the level for the desired culture volume.
We use a culture volume of 20 mils. Additionally, we mark the 21 and 19 mil volume marks to give a sense for variance from the ideal to start. Fill each tube with 21 mils and mark the water level for each.
Then remove one mil and mark the 20 mil mark and perform this again for the 19 mil mark. Before performing this next part, put onto eye protection and always wear eye protection and move cautiously when working with exposed needles. Next, set aside a single culture tube that we'll use to hold each of the corks as we pierce them.
With the three needles used in our minis stat design, this tube will get filled with bits of corks, so don't use it for experiments until you've had a chance to clean it fully firmly. Place the cork in the tube. Take the longest needle, the one with the pink backing and place its tip slightly offset from the center of the cork so that there's still room for the other needles.
Brace the cork and push the needle through such that the needle runs roughly parallel to the side of the glass chamber. There's an insert that comes with this needle. Save this because we'll use it later.
Next, take the media needle which has yellow backing and place it a similar distance from the center of the cork. About a third of the way around the cork circumference, brace the cork and push the needle through. Lastly, take the white backed effluent needle and pierce the cork in the remaining space such that the needle is roughly parallel to the wall of the glass culture chamber.
Push the needle in only halfway as this is the needle you'll use to set the culture volume, take the needle insert from the pink bag needle and use it to clean out the air and effluent needles. Then go in with tweezers and recover any piece of corks stuck to the needles on the corks underside. Finally, place the assembly into a clean and marked culture tube.
Do this for all four culture tubes. We will now build and arrange all non autoclave parts. Hydrating the culture chamber is important to prevent evaporation.
We have tried a number of methods and find that simply bubbling air through a 10 mil tope significantly decreases evaporation during the batch phase of growth at the beginning of our experiments. To start, use some pliers and break the end off. Next, remove the cotton plug from the backside and place it through the single hole silicon stopper.
Place this on top of the one liter filter flask. Then place an adapter and do a four centimeter long piece of large tubing and attach this to the top. Next, place a connector into another large piece of tubing around 10 centimeters in length and place this on the nozzle on the side of the filter flask.
After air is humidified in the hydrating chamber. It is split between four mini stats using a four port manifold. Take four four centimeter pieces of large tubing and place them on each port of the manifold.
Next, attach a long piece of tubing to the left side of the manifold and attach the other side of this tube to the hydrating chamber. For the setup shown here, this piece of tubing is 45 centimeters long. The important thing is that this tube is more than long enough to span the distance between the manifold and the hydrating chamber to power airflow.
We use aquarium pumps which pump just over 300 mils per minute of filtered and humidified air into each chamber. Attach a piece of small tubing of sufficient length to reach from the pump to the adapter at the top of the humidifying chamber. Varying this tube length within reason doesn't seem to affect flow rate into the chambers.
Next, we set up the peristaltic pump. For our experiments. In yeast, we use a media flow rate of 0.17 volumes per hour, which roughly correlates with a pump rate of 7.5 RPM on our pump.
Fine tuning and flow rate can be individually tuned using the occlusion nozzle on the left side of the pump cassette. It is important to determine ahead of time which pump rate will provide the ideal dilution rate for your experiment. Likewise, it is important to figure out what's heading on the heat block correlates to your desired temperature.
We use 30 degrees Celsius to recreate the environment experienced by a given culture. We usually bubble air into a chamber filled with water to best reflect the temperature within each chamber. Now let's assemble the three types of tubing that carry air and media into and out of the culture chambers.
The air tubing connects the four port manifold to each of the minis stat chambers. Make sure your tubing is cut long enough to do this. For this demonstration, I'm using tubing that's 45 centimeters in length.
On one end, we place an air filter, which acts as a sterile barrier filter humidified air before it is routed into the bottom of each culture chamber. On the other end, we place a lure connector to attach this tube to the pink back needle. The effluent tube connects each culture chamber to their respective a hundred mil sampling bottle.
Make sure the tubing is of sufficient length to do so. In this demonstration, the tubing is 48 centimeters in length. Simply add a lure connector to one side and it's ready to go.
The media tubing, rats media from a sterile media source by way of a peristaltic pump to each chamber. Make sure you have tubing of sufficient length to reach from the media source to the pump and from the pump to the culture chamber. First, take the 64 centimeter long piece of large tubing and attach it to the female.
Quick connect. On the other end of this piece, place a large to small adapter and connect this to a long piece of small tubing. Next, we assemble a branch system to allow flow of media from one source to multiple chambers.
In this case, four, plug the small tube and quick connect into this branch set of tubes. Next, take each of the peristaltic pump tubes, add on adapter needles and screw and male lure locks to both sides. Then we connect each of these into the branch set of tubes.
The other end of the pump tubing is then connected to one last set of tubes, which in this demo are 42 centimeters in length. These lead to a final set of lu connectors, which are screwed into the yellow backed media delivery needles at the top of each culture chamber. Now we'll assemble the effluent sampling containers.
To date, we have been using 100 or 125 mil glass screw top bottles as sampling chambers and find that adding corks to the tops keeps things more organized and sterile. We found that the corks can be a bit messy unless you place something to help route the media into the bottle. For this purpose, I've been using 200 microliter tips that have been cut as shown, placed in one of the two holes in the base of the cork.
Next two small tubes, four centimeters in length are placed in the top of the cork and capped with air filters that allow gas exchange in the autoclave and keep the chamber sterile. Afterwards, the array of minis stats form a closed system. We autoclave this system to sterilize it and use filters and aluminum foil origami to ensure sterility between autoclaving and the time of use.
Take a small piece of foil and fold it in half. Over the quick connect of the media line, press it flat and fold the corner's back. Make sure to make this fold tight against the tubing.
To seal this end against biological contaminants, fold the foil onto itself again and pinch it closed. Now it's ready for the autoclave. Now we will connect the tubes to each of the culture chambers.
Take the divided end of the media line and connect them to each of the yellow back needles in the cork assembly. Do so sequentially so the media tubing does not get tangled. Next, take the four airlines and connect them to the pink back long needles in the cork assembly.
Then take the filtered ends and wrap them in a big piece of aluminum foil. Tightly enough that water will not get in, but not so tight that air cannot escape during autoclaving. Next, take the effluent lines and place filters on the ends of each of them.
These will act as temporary sterile barriers. Screw the opposite end of each of these lines into the white backed medium length needles on the cork assembly. As before, fold a large piece of foil onto the filtered end such that air can escape but water won't get in.
Mark the F LX line so you can tell them apart from the air lines, each of the sampling chambers will also be fitted with foil wrappings to let air out and protect from water. During autoclaving car preparation is covered at length in the mini stat manual assemble and foil, a 10 or five liter car to prepare it for the autoclave. Now place everything including the car into one or more.
Autoclave bins autoclave these on liquid cycle with a 20 minute sterilizing time. This typically takes an hour and a half to complete. In this second video of the mini stat series, you will learn how to run an experiment for evolution in competition based studies and mini stats and learn how to clean up after you're done.
Before running an experiment, you must assemble an autoclave all components of the array as demonstrated in video one. Additionally, you must make and filter media as outlined in the minis stat manual and prepare an overnight culture grown in the appropriate hemostat media. Fill each of the hydrating chambers with 700 mils of DD H2O.
Take the minis stat array and place it in the heating block. Place the airlines into the four port manifold and turn on the aquarium pump. Remove the air filter and place the culture sampling line into the autoclave sterile a hundred mil sampling bottle, un foil and quick.
Connect the media in line to the car. Then undo the clip massage air out of the line. Turn on the pump.
If no one else is using the pump, you can crank it up to 90 RPM and the chamber should be full within 30 minutes. If an experiment is already running, then let the culture chambers fill with media overnight. Once the culture is filled to about 35 mil, turn off the media pump.
The extra media buffers against evaporation effects during the batch phase of growth to prevent contamination during inoculation. Spray each mini stat culture cork assembly with a liberal quantity of 70%ethanol and let it sit for a minute. If you want, you can dab the sides to remove excess ethanol.
Put on safety goggles and use a 22 gauge needle and one mil syringe to inoculate the mini stat with a hundred microliters of overnight culture. Make sure to inject the inoculum down into the media and not just onto the mini stat chamber wall. Then allow the culture to grow for 30 hours.
This batch phase of growth is a common tactic used to establish a saturated culture where a key limiting nutrient is depleted at the onset of the experiment. After 30 hours, you'll need to set the culture volume to 20 mils to do this, decrease the height of the white backed effluent needle and turn off the air bubbling into the culture in turns until you've arrived at a culture volume of 20 mils when the bubbling is turned off. Alternatively, if the culture volume is too low, turn the pumps back on and wait until they fill to the 20 mil mark in the absence of airflow.
Now that the culture volume is set to 20 mils, turn the pump back to 7.5 RPM or whatever setting correlates to 0.17 volumes per hour on your pump sample into clean culture tubes on ice once or twice daily. As the experiment dictates place the sampling corks into the collection tubes on ice, I usually sample for two hours, which provides 6.8 mils of culture. If your pumps are set right, you'll want to record the time you turn the pumps on and started taking your time zero measurement as well as the duration of the measurement and the volume collected.
The effluent collected into the 100 mil sampling bottles from setting the culture volume is not of interest, so you can dump this out when the two hours is up. Place the sampling corks back onto the now empty a hundred mil bottles and you can proceed to get a cell count and perform any other assays you may desire on your samples. Each day.
I sample my cultures for two hours on ice. While that's happening, I measure the effluent that has gathered since the last sampling to calculate the dilution rate and volumes per hour and the number of generations since the last measurement. Once the two hour sampling is finished, place the sampling line back into the now empty a hundred mil effluent collection bottle.
Use the sampled culture to quantify cells per mil and other endpoints of interest in-depth analytical methods to characterize population composition through use of genetic markers, as well as characterization of DNA and RNA is covered in the mini stat manual. Also provided is a template worksheet that we use in the lab, which helps organize data collection and automate data analysis. As you finish running your experiment, it's important to take down and process all parts of the same day to ensure that media does not dry out and contaminate tubing or other mini stat parts.
On the last day of the experiment, turn off the media and the air pumps for the last sampling. I oftentimes remove all lines attached to the culture chamber and place all tubing and separate autoclave bins. I then place the culture chamber itself on ice and measure culture volume cells per mil, and any other end points of interest.
Next, take each type of tubing and rinse it thoroughly with RO water. For media tubing, I rinse for minutes with other types of tubing at least 30 seconds. Then use the air pump to blow any remaining water out of each piece of tubing.
After that, you can organize and hang all tubing to dry for use in later experiments. Also, all quick connect should be rinsed with water and lift to dry. To clean the cork assembly, flush each needle thoroughly with DDH two oh.
Use chem wipe. Wipe off any media which remains on the exterior of the needle as well. Then use the needle insert to aspirate and dispense water from the media and effluent needles.
Wipe the exterior of the needles one more time and they're ready to be used. In your next experiment, clean the culture chambers with water and ethanol and store them upside down as described in the first video. Additionally, you should rinse the car in its cork thoroughly with RO water three times.
Be sure to have removed the quick connect from the base of the car and run water through the spout. Before reusing tubing, cork assemblies or culture chambers, I typically wash them again with RO water and make sure to remove any water remaining in the tubing by blowing air through them with the air pump. I hope you found this instructive and we wish you luck with your experiments.
If you have any questions, please refer to the mini stat manual.
