Buy another pump rack from Fynch
Final length 5.7 inches (144.78mm)
2X - 2x8 ribbon cable
Do for both of the aluminum rails. Diagram can be found here.
Cut to final length 5.7 inches (144.78mm)
If necessary, drill out the hole on each side so that your M6 x 1mm tap can work
Tap each side with M6 x 1mm tap
Unscrew 4 screws from top of original pump mount
Remove 20-20 aluminum tube
Replace with 40-20 tube. Screw in
Add 3D printed extenders
Screw in 3D printed extenders
Slot pump rack in
Plug in ribbon cables
3D printed (SLS, nylon PA-12) vial cap with ports for nylon tubing
3D printed (SLS, nylon PA-12) bubbler body
Nylon tubing (1/8" OD)
Stainless steel frit - 5 micron filter, 1/4" Diameter, 1/16" Thick
(Recommended) Rubber stopper
(Recommended) O ring
(Recommended) Push to connect fittings for testing bubblers
The nylon tubing will straighten in the autoclave
The length of the tubing connecting the lid to the bubbler will be dependent on the culture volume of your experiments, which is determined by the length of your efflux straw. As a good starting point, a 2 inch efflux straw will correspond to a 20mL culture volume, and a 2.5 inch length of tubing connecting to the bubbler will be ideal for that volume. If your culture volume is different than this, the best way to find an ideal tubing length is by experiment:
Cut a length of nylon tubing and fit it in the cap and 3D printed bubbler holder. Make sure that you cut so that your bubbler is below your efflux straw!
HOWEVER, use caution with how low you set the bubbler; if the bottom of the bubbler body is below the 15mL line in the vial, it will begin to interfere with OD readings
Using the length of tubing you found as a template, cut equal lengths of tubing for the remainder of bubblers you will be making.
It's a good idea to make an excess of bubblers (25% or so) relative to the number of complete vial caps you'll be making, because the bubbler assembly process is tricky and you may have some that are not useable (see section 3)
Mixing tray (here a small weighboat)
Disposable applicator
Foreceps/tweezers
Frit
3D printed frit holder
Nylon tubing cut to length
Use gloves to avoid getting epoxy on your hands
Follow the epoxy directions to mix up a small amount of epoxy in a disposable dish or a piece of cardboard. Use even pressure to get even amounts of resin and hardener.
Use a pipette tip or other similarly sized applicator to put the minimum amount of epoxy around the rim of the 3D printed bubbler body where the frit will be placed
Avoid using too much epoxy around the rim, as that will tend to block air flow through the frit
Avoid using too little expoxy around the rim, or having uneven coverage, as that will lead to large bubbles escaping through the rim and defeating the bubbling action of the frit. It's a balance!
Using tweezers, carefully set the frit in place on the bubbler body - it should be a slight pressure fit.
Try to avoid getting epoxy anywhere outside of the rim of the frit as this will reduce air flow through the frit, and produce fewer bubbles.
Do your best to set the frit on the bubbler body as parallel as possible to the frit holder rim. If you place it at an angle, it will likely be very hard to press in. If this happens, your best bet is to take the frit off with tweezers and try setting it in place again.
Add epoxy to one of the ends of the previously cut nylon tube. Get the epoxy around the outside of the last 1/4-1/2" of the tube, with roughly even coverage. If you get epoxy inside the tube, you can blow it out or use a paperclip or similar object to clear it.
Set that end of the tube into the bubbler body, and give it a twist to ensure the epoxy is spread evenly at the joint. Don't force the tube all the way down into the bubbler body, there should be a lip that catches the tube with slight pressure.
Set bubbler upright to cure overnight
To our best guess, the stainless steel bubbler frits need to be passivated, but they are incompletely passivated when they leave the factory. Without this step, they will produce larger (worse) bubbles, likely due to a more hydrophobic surface.
Passivate the frits by submerging in liquid
Option 1: Soak in water overnight
Option 2: Soak in LB-Miller media or 5% citric acid for 1 hour. Lemon juice will do in a pinch.
Create a test system for the bubblers
You can attach the normal 1/16" ID eVOLVER tubing to the end of the nylon, but this may take more time per bubbler as the silicone tubing is hard to put on the nylon tubes
Especially if you have a lot of bubblers to test, you may want to use a push to connect fitting for easy swapping of bubblers
Connect the bubblers to your air distributor (ie your 1:16 air supply)
Check that your bubblers work well in rich media like LB, YPD, or BHI. As a rule of thumb, the less salt and peptides your media has dissolved in it, the larger your bubbles will be.
If one of your bubblers are not bubbling at all, check your tubing connections and try to run air through just that bubbler. If it's still not producing bubbles at 5psi of back pressure, it's likely blocked with epoxy, and is not useable.
If one of your bubblers is producing a steady stream of large bubbles from the rim of the frit, it means there's an incomplete epoxy seal against the bubbler body. You can attempt to fix this by adding more epoxy on the outside, but it'll likely be easier to just make a new bubbler.
Similar to the assembly of the nylon tube into the bubbler body, apply expoy to the other end of the tube attached to your bubbler, insert into one of the ports on the bottom of the vial cap (whatever one corresponds to your preferred layout; my preference is for media in on the right, bubbler in the back, efflux straw on the left).
Again, be sure to check for epoxy blocking the tubing, and clear with a paperclip if it's blocked.
Set assembled caps upright (with a vial rack or spare vials) so that epoxy doesn't drip into the o-ring groove as it cures.
Allow epoxy to cure 24 hours before sterilizing via autoclave as part of experiment prep.
An in-vial aerator, which can be hooked up to a gas stream of your choice. Normal stir bar-based mixing in the eVOLVER does not provide enough gas exchange in many situations. These bubblers produce small bubbles that greatly increase gas exchange.
The assumption is that there will be variability between the bubblers that you make -- even though you will screen them before putting them in a vial cap! This is compensated for by bubbling much more than we need for a given microbe's gas consumption needs. Worse bubblers will not be a problem if the worst bubbler you have provides more than enough gas exchange.
Aerators are great substrates for biofilming. If your strain biofilms, make sure you swap in new bubblers when bubbles noticeably diminish. See Cleaning Protocol.
Because of biofilming, it is useful to have another set of caps/bubblers that can be swapped in without interrupting your experiment. Ideally, you would pre-autoclave these and keep them sterile in aluminum foil. Pause the experiment, check if the vials need to be changed because of biofilm and swap sterile caps and/or vials in.
Bubblers will produce different sized bubbles depending on media type. This is related to the amount of salts, proteins, carbohydrates, etc that are in the media. These can act as surfactants to create smaller bubbles. Therefore, largest bubbles will be in water and smallest bubbles will be in rich media, as can be seen below:
<Add picture of bubbler in water vs algae media vs LB>
Although autoclaving the lids with bubblers removes the risk of contaminating the next experiment, you'll ideally want to remove media residue and any biofilm that may have accumulated on the bubbler. This is best achieved in the following process, which utilizes 3 valves and Y connectors to control the pressure in the tubing connected to the bubbler: a valve to pressurize the bubbler, a valve to apply vacuum to the bubbler, and a valve to "neutralize" the bubbler to ambient pressure. This can also be achieved manually with a syringe.
Remove all lids from their vials
In a 1L beaker full of 10% bleach, pressurize the bubbler to purge any residual media/bacteria into the bleach
Apply vacuum to the bubbler to pull bleach into it
Neutralize the bubbler pressure once you see bleach heading up the tube connecting the bubbler to the lid
Repeat with the rest of your lids, and allow them to rest submerged in bleach for 30 minutes (don't leave them in bleach for longer than a few hours, as the steel in the bubblers will begin to rust)
Pour out the bleach, rinse the lids with water a few times, and refill the beaker with water
Pressurize each bubbler to purge the bleach inside each bubbler into the water
Apply vacuum to pull in water (or 70% ethanol), and then apply pressure to purge
Leave your lids to dry in a rack or put them in vials for your next experiment!
If residual salts on the bubbler is not a concern, a simpler protocol is to soak the lids in 10% bleach (without any purging or vacuum), and then blow out the resultant bleach/media mixture with an air supply or syringe, and then rinse with water.