Cox Systems Biology

Contents

1. Safety
2. General principles of cell care
3. Sterile technique
4. Gas tanks and modular incubator chamber
5. Preparing stocks
6. Cultures in gel
7. Passage
8. Freezing, thawing, and maintaining lines

• Appendix 1: RG2 Tissue Culture Standard Operating Procedure
• Appendix 2: Trophoblast Organoid Media (TOM)
• Appendix 3: Stock solutions for TOM components
• Appendix 4: Dilution calculation examples

1. Safety

Many applications of 3D cell culture use primary human cells as their starting materials. This means that performing these cultures is Level 2 biohazardous work. Our lab’s guide for working safely in Risk Group 2 settings is here: RG2 SOP and in Appendix 1. There are some key take-away points from this guide, although all of it is important:

1. Perform all work inside a functioning BSC wearing proper PPE, including a lab coat with cuffs, and intact nitrile gloves
2. Clean the BSC with 70% ethanol and clean all objects with 70% ethanol when you put them into the BSC, and when you take them out
3. Dispose of liquid waste using the aspirator in the BSC. Ensure there is bleach in the aspirator and wait 30 minutes for it to inactivate hazards before emptying it.
4. Dispose of solid waste in a biohazard bag inside the BSC. Place this in the larger biohazard bin outside of the BSC after your work.
5. Clean glassware by collecting all dishwater into a large beaker filled with bleach in the sink. Wait 30 minutes for the bleach to inactivate hazards before emptying it.

While your cell culture work is important, keeping yourself and the people around you safe is the number one priority. If these goals ever conflict, always choose the living humans over the cells!

Back to top

2. General principles of cell care

Growing cells is basically like taking care of really weird, mostly aquatic, very fussy plants. They need to be kept at the proper temperature, they need to be fed regularly, and they need adequate humidity. They also dislike when they are too crowded together, or too lonely. When they get too crowded, you need to ‘split’ them apart in a process called passaging (see Chapter 7 for more information about passaging). Very importantly, they must also be kept healthy by growing them in the absence of pathogens (see Chapter 3 for more information about sterile technique).

Organoids are particular in ways that their 2D counterparts cannot even fathom. Their cores become necrotic if they are allowed to get too large. If the organoids are grown in gel (which our current trophoblast organoids are), their gel domes need to be kept stably polymerized during culture, and depolymerized for any manipulations of the culture (see Chapter 6 for more information about culturing in gel). If the organoids are grown too close to the bottom of the dish rather than in their 3D droplet, they may ‘touch down’ as they grow, which will lead them to differentiate rapidly and decrease their lifespan.

Cell and organoid cultures are kept at the proper temperature by growing them inside an incubator kept at 37^oC. Most incubators also have a water bath to maintain humidity of the cultures and CO₂ intake, but these features do not need to be used if you are performing cultures in a Modular Incubator Chamber, which is the case for our current trophoblast organoid cultures.

Organoid media needs to be prepared fresh every 2-3 days. The recipe for trophoblast organoid media that we use based on our current stock solutions can be found in Appendix 2. This media is made based on the protocol described by Turco et al, 2018, which can be found at 10.1038/s41586-018-0753-3. It contains many growth factors that ensure a population of trophoblast progenitor cells is maintained in our cultures, which is why they can be passaged, frozen, and thawed. You can learn more about making stock solutions for media (for supplementing any kind of culture) in Chapter 5.

Back to top

3. Sterile technique

One of the most important parts of cell culture is to keep the culture free of bacterial and fungal contaminants. This is achieved by always using sterile technique when working with the cultures.

While this ethanol will help prevent the risk of contamination, you cannot assume that it is 100% effective at removing contaminants from the surfaces that you clean with it. There are other steps you must take as well.

During work, do not rest your hands or arms on the grills of the BSC. This will prevent it from working effectively, which could not only lead to contamination of your cultures, but is also unsafe for you. It is challenging to hold your hands up for work in the BSC unsupported for long periods of time. Take short breaks when needed to prevent yourself from getting too fatigued and becoming unable to hold your hands up. Some people also find it helpful to rest their hands on top of tip boxes inside the BSC when they are not using them. I find it helpful to rest my forehead against the outer wall of the BSC while I work so that I can direct as much of my core strength as possible to my arms.

When you have open culture dishes or open containers with sterile contents inside, you risk contamination of the contents if you pass your hands/arms overtop of them. There are a few steps you can take to reduce the chance of this occurring. When you are working with an open container of sterile contents, pick it up and hold it above the surface of the BSC rather than leave it in a tube rack.

Keeping our cultures sterile is also made difficult by the fact that the source materials for them are primary human cells. These cells are very likely contaminated with bacteria, yeast, and other pathogens as well when they are obtained in the clinic. To try to kill whatever is coming in with our source materials, all of our culture media contains antibiotic-antimycotic solution.

Even when taking all of these steps, culture contamination/infection is still possible. Check each well every time you feed the cells to make sure this has not occurred. Bacterial contamination looks like small grains of sand or television static, and may have an odor. It can also appear as dark large colonies within culture gel. Yeast contamination looks like much smaller cells that bunch together like evil pearls. Mold contamination is visible to the naked eye as a white fuzzy growth. All forms of contamination often come with a change in the media colour indicator to yellow, or the media may appear cloudy. Discard any contaminated wells, and if possible discard the entire plate when contamination occurs in case it spread to other wells but is not yet detectable. If it is essential that you keep the contents of a plate where contamination occurred in other wells, move the contents to a fresh plate. Thoroughly clean the entire incubator and/or culture chamber with ethanol in the event of contamination.

Back to top

4. Gas tanks and modular incubator chamber

Working with a modular incubator chamber means that you will need to perfuse the chamber with mixed medical gas from a gas tank. To do this you will need to know how to use the gas tank and its regulator. Labelled pictures of these items are below. You will notice in the picture that the gas tanks are secured to the wall with belts. This is important for everyone’s safety as it makes sure that the highly pressurized tanks do not tip over. The tanks must be kept secured in these belts at all times.

The steps for perfusing the modular incubator chamber with mixed medical gas are:

Images showing the modular incubator chamber’s intake and outflow valves, and how to squish the intake valve when the tubing is stuck shut, are below.

When the gas tank you are using is empty, you need to place the regulator on a new, full tank. To do this, you need to use the big wrench, then place the regulator on a new tank and tighten its screw. Loosen the regulator control until it is almost completely off, open the regulator knob, then open the gas tank as little as possible. You will see the cylinder gauge reading display without any reading on the delivery gauge value display. Then, tighten the regulator control until the delivery gauge shows that gas is flowing and the flow meter shows a flow rate of 25 Litres per minute. Then, you can turn the gas flow off.

You can order more gas tanks from Linde (Praxair) by calling 1-800-225-8247. People from the company will come to drop off the new tanks in the correct place and take away the old empty tanks. If you need to move a compressed air tank, only do so using a shipping dolly that secures the tank with a strong metal chain.

Back to top

5. Preparing stocks

Many types of cell culture media, including that used for our trophoblast organoids, require supplementation with additional growth factors, nutrients, and other components that support culture maintenance. These additional components are often shipped to us in lyophilized powder form, and they then need to be reconstituted in solution. In each form (both powder and in solution) most factors are not stable for long term storage in the fridge. They need to be stored in the -20^oC freezer, but they will also be degraded if they undergo multiple freeze-thaws. This problem is solved by making concentrated stock solutions of each additional media component, then dividing the stock into lots of tiny tubes called aliquots, so you only need to thaw the amount you need each time you make media.

The amounts of each additional media component needed for 10mL of Trophoblast Organoid Media (TOM) in our lab can be found in Appendix 2, and the concentrations of their stock solutions and how to prepare them can be found in Appendix 3. This chapter will cover the general methods of preparing stock solutions for any kind of culture media.

Stock solutions should generally be made as concentrated as possible, while ensuring that the media component still dissolves (if it was supplied as a powder). This will help maintain the shelf life of the stock as well. When media components are supplied in liquid form, these can also often be divided into smaller aliquots and frozen at -20^oC, although some of the components supplied in liquid form will actually degrade if frozen. Always check the product datasheet that the supplier provides for their storage and reconstitution recommendations. Some media components are not soluble in standard solutions like PBS. When media components need to be dissolved in something that is toxic to cells like DMSO, it is especially important to make the stock solution as concentrated as possible. In deciding how to prepare stocks, you should also consider the fact that you are a human and you will not want to put small amounts of liquid into 300 tiny tubes for one media component. I usually try to arrange to make no more than 100 aliquots at a time for any given media component.

All tubes for aliquoting should also be sterile, and you should perform all of your aliquoting inside the BSC using sterile technique. You can put snap cap tubes into an empty pipette tip box and bring them to sterilization services on MSB’s 4^th floor for autoclaving to ensure that they are sterile. For any powders that you have to weigh, use a sealed conical tube instead of a weigh boat to ensure that the media component remains sterile. When you are preparing aliquots for culture media, the protocol you are following may provide the desired concentration in grams (or milligrams, nanograms, etc.) or in moles (or millimoles, nanomoles, etc.) per mL (or per Litre if it is provided as a molar M amount). You will typically want to put the entire vial of powdered stock into a stock solution, but you may also encounter situations where you will have more powdered stock than you want to dissolve for aliquoting.

For step-by-step examples of how to perform dilution calculations, see Appendix 4.

Back to top

6. Cultures in gel

Our trophoblast organoids are grown in Growth Factor-Reduced Matrigel. This product is derived from a mouse tumor cell line called EHS cells and consists of many extracellular matrix components including collagens and laminins. Because these basement membrane components are present, many types of epithelial organoids grown in these gels will “flip” their polarity, since the cells think the gel is their basement membrane. This is true of our trophoblast organoids as well—unlike placental villi, trophoblast organoids grown in gel have a cytotrophoblast shell that encases a syncytiotrophoblast core.

EHS-derived ECM gels are unique because they stay in liquid form and can be pipetted when they are ice-cold, but they polymerize to be able to hold their structure when warmed. To use the gel, you will take an aliquot out of the freezer and allow it to thaw on ice for a few hours. It must be kept ice cold as it thaws to be able to use it, because once it is warmed and it polymerizes you will not be able to use it anymore. You will then need to keep the tube of thawed gel on ice as you pipette volumes out of it and place any tubes you will put the gel into on ice as well. The gel should only be removed from ice during the brief (as brief as possible) times it is inside a pipette tip and when it is placed on the plate for culture. When pipetting the gel, you will want to avoid making bubbles as much as possible, because the gel will hold on to the bubbles forever once it polymerizes. To prevent bubbling, watch as it is taken up into your pipette tip to make sure you don’t remove the tip too early, then when you dispense the gel, never go to the second pipette stop. While you will lose small volumes of gel this way, only going to the first stop will make a big difference in preventing bubbles. It is worth it!

When aliquoting the gel, the 10mL vial that the company ships will need to be thawed overnight on ice in the fridge. Keep the tubes in a ziplock as well to prevent any of the ice water from breaching and contaminating the tubes. Right before you aliquot, I recommend placing all aliquot tubes on ice, and using pipette tips that have been kept in the freezer overnight to reduce the chance of the gel polymerizing prematurely.

Some people use EHS-derived ECM gel to coat culture dishes in a thin layer to support 2D cultures. For 3D cultures, domes of polymerized gel are prepared in the centre of the wells of the culture dish, and warm culture media is overlaid on top after the gel domes have polymerized. For our organoid setup, 40mL volumes of ECM gel are pipetted into the centre of the wells of a 24-well plate. I recommend that you use 24-well plates made by Sarstedt, as I have been unable to have the gel domes become sufficiently solid when I have tried to use plates from other manufacturers. It is difficult to describe the process of making a gel dome. I recommend trying to make some practice domes for yourself if someone has some extra gel that has been left out of the freezer in the fridge for a couple of days that would not be usable otherwise.  

Many protocols recommend allowing the gel domes to polymerize inside of the 37^oC incubator before pipetting warm media on top of them. I have found that room temperature is sufficient for dome polymerization. With my use of the modular incubator chambers inside my standard incubator, anything I warm in my standard incubator dries out very quickly, so I prefer to polymerize the gel on the bench when possible.

Back to top

7. Passage

When organoids growing in gel become too crowded within their gel domes, or when individual organoids become too large, you need to move them to new gel domes so they have enough space to live and receive adequate nutrients. You can do this in a process called “passaging”, also known as “splitting”. Through this process, you will not only allow your organoid cultures to continue, but you can create the conditions so that increased numbers of organoids can grow by “expanding” your cultures.

When you passage organoids growing in gel, you can do what I call a “hard pass” or a “soft pass”.

A hard pass is necessary when the organoids are becoming too large, and involves breaking up the large organoids into smaller pieces. Breaking up the organoids through a hard pass is also what you need to do if you want to expand the cultures. To get to the stage where the organoids can be broken up, you will need to completely depolymerize their gel domes. To do this you use Organoid Harvesting Solution, which is a gentle method that uses surfactant rather than enzymes to breakup the gel, ensuring the organoids do not break down into single cells during treatment. You can place the cultures in Organoid Harvesting Solution in the plate you performed cultures in, or in a 15mL conical tube.

To harvest the organoids within the culture plate, you remove the culture media, wash the dome once with room temperature PBS, then place 500mL of Organoid Harvesting Solution in each well.  You can then disrupt the gel dome with your pipette tip, and then put the whole plate in the fridge for one hour. If there are cultures growing in the plate that you do not want to have in the fridge for this hour, you should harvest the organoids in 15mL conical tubes.

If the domes are stable but you want to move them to a 15mL tube, you should first remove the culture media and wash once with PBS, but when you place 500mL of Organoid Harvesting Solution in the well, scrape across the bottom of the well with your pipette tip to completely detach the dome, then pipette it into the new conical and place it on ice. You may need to add a bit more Organoid Harvesting Solution to the empty well, scrape it again, and move this to the conical tube as well if it appears as though gel remains in the bottom of the well after your first try. The conical can then be taken off ice and moved to the fridge for one hour.

If the domes are not stable, scrape the well with your pipette tip to detach the portion of the dome that remains stuck, and then transfer the dome and its media into a 15mL tube. Bring the total volume up to 10mL with cold media or PBS, then spin. Once the spin is done, aspirate the supernatant, and resuspend the remaining gel/organoids in Organoid Harvesting Solution. If you decide to put more than one dome in the same tube (this is a great option to limit the amount of tubes you use, as long as all of the domes are from the same donor), add 1mL of Organoid Harvesting Solution for every 4 domes. Otherwise, the volume of 500mL will work if you have 1-3 domes’ worth of gel in the tube. You can get away with using less Harvesting Solution when the organoids are in tubes because doing the spin before adding the Harvesting Solution will allow for partial removal of some of the gel.

In any case, after the Organoid Harvesting Solution has been on the organoids for one hour in the fridge, you will need to remove the solution. Move the organoids in solution to 15mL conical tubes if they are not already in tubes, then bring the volume in the tube up to 10mL with cold DMEM/F12. Spin at 1600rpm at 4^oC for 5 minutes, remove the supernatant, then wash again with cold DMEM/F12 by resuspending the organoids and bringing the total volume up to 2mL and repeating the spin. You can then resuspend the organoids in the appropriate volume of Cultrex gel.

A soft pass is what you do when breaking up the organoids is not necessary, but the organoids have become over-crowded. You do not need Organoid Recovery Solution to do a soft pass.  Remove the old culture media and place 500mL of cold DMEM/F12 into the well. Scrape back and forth with the pipette tip to detach the gel dome from the bottom, then transfer the well contents to a 15mL conical tube on ice. To ensure that all of the gel is removed, place another 500mL cold DMEM/F12 into the well and scrape the bottom of the well again, and move the contents to the same tube. You can then top up the total volume of the tube so that there is 2mL of cold DMEM/F12 for each gel dome that is put in the same tube. Spin at 1600rpm at 4^oC for 5 minutes, then remove the supernatant.

This first spin will allow for incomplete removal of the gel, and you will see some remain at the bottom of the tube, but the organoids will be located below the remaining gel at the very bottom of the tube. At this stage you can either add new gel if the volume of gel to be added is significantly higher than what remains, carefully pipette out the gel without disrupting the organoids if it is abundant and you are confident you can do so, or repeat the spin to remove even more gel before new gel is added.

Back to top

8. Freezing, thawing, and maintaining lines

The trophoblast organoids that we grow in our lab can be frozen and thawed, essentially creating ‘cell’ lines that allow for reproducibility of our experiments and thorough validation of each line.

The organoids are originally derived from primary trophoblast cells that we obtain from human placental samples. If you are trying to start a new organoid line, prepare the cultures so that there are no more than 50 000 trophoblast cells per 40mL gel dome. To determine how many cells you have in a sample, you need to count them using a hemocytometer. To count using a hemocytometer, take 10mL of your (thoroughly mixed) sample and combine it with 10mL of Trypan Blue in a small snap cap tube. Then, load 10mL of this mixture into each chamber of the hemocytometer so that the liquid disperses under the glass and across its grids. Bring the hemocytometer over to the microscope and count the 4 corner squares of the grid. Then calculate the amount of cells per mL for your sample by following the instructions below, and measure the correct volume to result in <50 000 cells per dome.

You can also initiate cultures using organoids that you have already frozen. Freezing organoids is stressful for them, so the organoids should be as happy as possible before they are frozen to increase the chances that they will recover once thawed. I have found that organoids that have not gone through too many passages yet (less than 4) tend to recover best after being frozen and thawed. To freeze organoids you need to go through the same steps that you would use for passaging to remove most or all of their gel, then resuspend them in freezing media rather than normal trophoblast organoid media. Freezing media is made up of 70% complete trophoblast organoid media, 20% FBS, and 10% DMSO. Because DMSO is toxic to cells (but essential for preventing osmotic shock during freezing and thawing), you should add DMSO last and get the organoids into the freezer as quickly as possible after adding it. Once the organoids are resuspended in freezing media, they will need to be moved to clearly labelled cryovials.

I try to set up my freezings so that the contents of one cryovial will contain enough organoids to populate two gel domes in a volume of 1mL of freezing media. The cryovials should then be placed in a slow freeze chamber and moved to the -80^oC freezer. Once at least 24 hours have gone by and the organoids in the slow freeze chamber have completely frozen at this temperature, they can be moved to our liquid nitrogen tank (“LN2”) for long term storage.

In preparing freezings, it is helpful to consider what our current organoid stocks look like: which ones are running low, and which ones do we have plenty of already? You do not need to spend time making freezings of organoid lines that we already have abundant stocks of, but it is generally good to try to make a new freezing each time you thaw one so the line can be maintained. If there are lines that are running low, it is great to try to make extra freezings! For additional safeguard of our organoid lines, make freezings on different days rather than all at one time. This way, if something goes wrong with one of the freezing preps (such as contamination, or the organoids being more vulnerable on that day for another reason that prevents their recovery after thaw) you will have other freezings that will have a better chance of succeeding.

To thaw a cryovial containing organoids or primary cells for culturing, you will need to obtain it from LN2 storage. ALWAYS wear a face shield, nitrile gloves, and freezer mittens when moving cryovials to LN2 or taking cryovials out of LN2 to thaw as there is risk of its contents exploding. We have an inventory of cryovials in LN2 with their locations (straw, box number, and grid display showing each vial’s location in the box). It is best to consult the inventory before you go to take a vial out so you can do so efficiently to prevent unintentional thaw of other cryovials. Please cross the vial you are thawing off of the inventory as well so we know it is not there anymore. Once you have located the cryovial you want to thaw, remove it from the box and very briefly remove its lid to depressurize it in the event that there is liquid nitrogen that got inside of it. The risks associated with doing this are very minimal since the sample will stay completely frozen if you do this quickly enough, and the risks associated with an exploding cryovial are greater than these. Next, place the tube inside of a Styrofoam box for transport so that if it explodes, the Styrofoam will help to absorb the shock.

When you bring the cryovial that you are thawing over to the BSC, open it and add 500mL of DMEM/F12 to help speed up the thaw and give the cells some nutrients. Hold the tube while it is thawing so it is warmed in your hands—it is best that the thaw occurs quickly once it has begun. After the whole vial has thawed, move its contents to a 15mL conical tube and slowly bring the total volume in the tube up to 10mL with additional DMEM/F12. Washing with lots of media will give the cells nutrients quickly while also ensuring complete removal of DMSO. Spin the tubes at 1600rpm for 5 minutes, remove the supernatant, resuspend in 2mL of DMEM/F12, and repeat the spin. Then, place the tube on ice and resuspend the organoids in 80mL of Cultrex gel. Divide the organoid suspension between two wells of a 24-well plate.

Back to top

Appendix 1: RG2 Tissue Culture Standard Operating Procedure

RG2 Tissue Culture SOP

Treat both risk group 1 (RG1) and risk group 2 (RG2) cells as RG2 for ease of operation.

1. Enter containment zone (tissue culture lab).
2. Don lab coat and nitrile gloves. If any work will be performed outside the Class II, Type A2 biological safety cabinet (BSC), also done face/eye protection.
3. Follow Canadian Biosafety Handbook (CBH) section 11.4.1 for preparing to work in the BSC:
   1. Turn on blower.
   2. Open sash to proper height.
   3. Adjust chair so that underarms are level with bottom of sash.
   4. Confirm inward airflow by holding KimWipe at middle of edge of sash. Wait 5 minutes before next steps
   5. Disinfect interior surfaces of BSC with 70% ethanol including all items that remain in the BSC (pipettes, pipette tip bucket, tip boxes, aspirator). Check to make sure there is enough room in aspirator for your purposes. If not, empty in the sink and add ~500mL Lavo 12 bleach.
   6. Disinfect all required items with 70% ethanol and place them in the BSC.
      1. Do not block the front/rear grilles.
      2. Place all items required for experiment inside BSC so that movement of arms in/out of BSC is minimized. Exclusion: serological pipettes
         1. Hook 70% ethanol spray bottle on BSC opening near dominant hand
         2. Place waste container/bag inside BSC towards rear.
      3. If right-handed, organize clean items on right and dirty items (e.g., waste container) on left. (Reverse if left-handed.). Excluding pipette tip bucket – place on the same side as your dominant hand
   7. Wait 5 minutes for ethanol to dry, air to purge and airflow to stabilize.
4. Follow Canadian Biosafety Handbook (CBH) section 11.4.2 for culturing cells in the BSC:
   1. Perform work as close to the rear of the BSC as possible.
   2. Do not rest arms on grille or work surface.
   3. Minimize disruption to air curtain by avoiding excessive movement of arms.
      1. Move arms in/out of BSC slowly and perpendicular to front opening.
   4. Only allow one person to work in the BSC at a time.
   5. Minimize air curtain disruptions by not allowing other individuals to walk back and forth behind BSC user.
   6. In the case of a spill, decontaminate all items in the BSC (see spill procedures).
5.  Decontaminate liquid waste by aspirating it with glass pipette into aspirator containing abundant Lavo 12 bleach. Do not empty aspirator unless 30 minutes has passed before its last use to ensure sufficient contact time. 
6. Follow Canadian Biosafety Handbook (CBH) section 11.4.3 for completing work in the BSC:
   1. Put used pipette tips inside waste bag. Close the waste bag and decontaminate its exterior with 70% ethanol. Spray abundant ethanol onto outside and inside of pipette tip bucket. NOTE: perform these steps any time pipette tip bucket or waste bag is filled.
   2. Decontaminate all items in BSC with 70% ethanol.
   3. Wait 5 minutes for ethanol contact time and for air to purge.
   4. Wash all dishes/instruments by removing them from BSC one at a time.
      1. Use sparklean or dish soap as appropriate for material.
      2. Wash items carefully with all water from wash directed into large beaker containing Lavo 12.
      3. When beaker is full, wait 30 minutes before emptying into sink and washing subsequent items. 
      4. Place on a clean blue absorbent pad for drying and keep separate from dirty dishes
      5. Ensure the eye wash station is not obscured by dishes
   5. Remove waste bag from BSC and dispose of it in biohazardous waste stream (yellow bucket).
   6. Remove as many items as possible from BSC leaving pipettes, pipette tip bucket, pipette tip boxes, and aspirator. Pipette boxes remain in the BSC to ensure they are kept as sterile as possible, since open boxes are not airtight
   7. Decontaminate all interior surfaces of BSC with 70% ethanol.
   8. Allow 5 minutes for 70% ethanol contact time.
   9. Close sash.
   10. Turn off blower.
7. Doff personal protective equipment (PPE).
8. Exit containment zone.

Back to top

Appendix 2: Trophoblast Organoid Media (TOM)

With our stock solutions, 10mL of Trophoblast Organoid Media (TOM) can be made by supplementing it with the following components:

	Amount	Final concentration
Antibiotic-antimycotic	100mL	1X
N-2 supplement	100mL (entire aliquot)	1X
B-27 supplement	200mL (entire aliquot)	1X
FGF2	10mL (entire aliquot)	100ng/mL
R-spondin 1	8mL (entire aliquot)	80ng/mL
L-Glutamine (Q)	100mL	2mM
N-aceltyl-L-cysteine (C)	10mL	1.25mM
EGF	5mL	50ng/mL
HGF	5mL	50ng/mL
Y27632 (Y2)	4mL (or 10mL if <one week post-thaw)	2uM
CHIR99021	1.5mL	1.5uM
A-83-01	1mL	500nM
Prostaglandin E2 (PGE2)	1mL	2.5uM

Back to top

Appendix 3: Stock solutions for TOM components

	Concentration	Reconstitution	Aliquot size
Antibiotic-antimycotic	As supplied	n/a	5mL
N-2 supplement	As supplied	n/a	100mL
B-27 supplement	As supplied	n/a	200mL
FGF2	100 ng/mL	PBS with 1% BSA	10mL
R-spondin 1	80 ng/mL	PBS with 1% BSA	8mL
L-Glutamine (Q)	200 nanomoles/mL	n/a	1.5mL
N-aceltyl-L-cysteine (C)	1.25 mmoles/mL	DMSO	250mL
EGF	100 ng/mL	PBS with 1% BSA	20mL
HGF	100 ng/mL	PBS with 1% BSA	10mL
Y27632 (Y2)	5 nanomoles/mL	DMSO	20mL
CHIR99021	10 nanomoles/mL	DMSO	10mL
A-83-01	5 mmoles/mL	DMSO	500 and 10mL
Prostaglandin E2 (PGE2)	2.5 nanomoles/mL	DMSO	10mL

Back to top

Appendix 4: Dilution calculation examples

Back to top

Citation Guide

APA
IEEEE

Back to top

Last updated: 12/2025