This is a post by Christine Miller, Research Assistant at Harvard University, Amy Wagers Lab:
I think it is fair to say that most people who have experience with cell culture know that there is at least some degree of “black magic” that goes into getting a particular protocol to work. In my experience, I’ve found this to be especially the case with iPS/ hES cell culture. In this series of blog posts I hope to shed a little light on this “black magic,” to talk about what I’ve found works, and hopefully to generate a platform for others to share their secrets as well.
Even though iPS cell culture is a relatively new technology, there are already tons of protocols for culturing them—each with its own variations on the amounts of reagents to add to culture media, methods of passaging, ways of freezing down lines, and the list goes on. Clearly, there are countless variables to test if you want to optimize your culture strategy. In addition, however, I have found that not only are there variables in technique, there are also lots of differences in iPS lines, even when they are all reprogrammed from normal patients. These differences may be illuminated in ways like pluripotency tests, where one line may take exactly six weeks to form a clear teratoma while another line may not exhibit tumors until 10 to 12 weeks. This might not sound too surprising on paper, but when you have injected a couple different lines on the same day and six weeks down the road all your lines except for one have teratomas, it is easy to think that that last line just didn’t work. If you wait another couple weeks, you may be surprised to find a cage full of mice with teratomas. Also, differences in lines may become very obvious while trying to differentiate iPS cells down a particular lineage. Currently I have been working on driving cells down the hematpoietic lineage, and I’ve found that the culture conditions for differentiating one line are quite different from differentiating another line. Even variables as small as the line’s growth rate or passaging timing may be different. My point with all this is simply that you should be aware that these differences exist and to be open-minded if your experiences with one line do not translate 100% to those with another line.
So, moving on to the good stuff—how to deal with some of these variables. I’m going to give you the “Dummies” edition of what specifically I have found to work well culturing my cells.
iMEFs vs. Matrigel
There are two main ways to culture iPS cells: you can culture them on a feeder layer using irradiated mouse embryonic fibroblasts (iMEF), or you can culture them feeder free. Depending on your desired application, both methods have their benefits.
Here is the breakdown of what I have found using iMEFs.
iMEFs are really great if you are thawing a line you’ve never worked with before. They are reliable in culture for a good 10 days, which should give you enough time to see a couple small colonies form. Also, the iPS colonies formed on the iMEFs will be nice and uniformly shaped, so you will be able to clearly identify where your colonies are and where differentiation (if any) is occurring. However, there are a couple things that must be taken into account using iMEFs. First, you must use good-quality iMEFs. If they are not high quality, they will not provide the appropriate feeder layer and support that your iPS colonies need, resulting in failure to seed or improper seeding that leads to differentiation. You want your colonies to fit fairly snugly between the iMEFs so that they can stay contained and undifferentiated. However, if they are too snug (the iMEFs are plated too densely), the colonies will grow vertically and risk differentiating on account of not having the space to expand horizontally. I’ve used both homemade and purchased iMEFs and have found for my needs it is more cost effective to buy them. I get them from global stem (cat # CF-1 MEF), and it costs $24 for a vial of 2M cells. I plate them at 200k/well of a six-well plate and do this by splitting one iMEF vial over 10 six-well plate wells (ie: 1 and 2/3 plates). I’ve tried plating anywhere from 100K to 300k, and 300k was definitely way too much, but 100k was a bit too sparse for my iPS cells to seed well. Making sure they are evenly spread out over the plate is also really important, so be sure to do the “T” motion at least three times in the hood and then at least one more time in the incubator. My only comment about the homemade iMEFs is that, unless you are making them to share with many others (and therefore can take turns harvesting, irradiating, and preparing them), it’s a lot of work and may not necessarily save you that much money. The main downside to using iMEFs is that it’s a much more time-consuming process. In order to passage or seed iPS cells onto them, first you would need to gelatin-coat your plates, which will take a minimum of four hours to set. Then you can plate your iMEFs, but those need to sit overnight in order to plate properly. Ultimately, then, this means that preparing your plate needs to start one or two days before you want to plate your iPS cells on it.
Feeder free, on the other hand, is very quick to prepare, taking only one to two hours to set. The main products on the market right now for this are Matrigel (BD), CELLstart (Invitrogen), and Vitronectin XF (Stem Cell Technologies). I have only tired Matrigel, but from the descriptions of CELLstart and Vitronectin, they sound very similar. Another benefit of using one of these feeder-free systems is that they are quite a bit more streamlined and simple. The media usually comes as part of a kit, where you only have to add a couple of things (if anything) in. There is usually some sort of standardization with these systems allowing you to purchase not only your media but also a recommended passaging reagent and freezing reagent, which can be nice as well. My last comment about working feeder free is to make sure you are buying the hES-grade material. The first time I ordered Matrigel, I didn’t realize that there were differences in grade and purchased a non-ES cell-grade one. After about six days in culture, the Matrigel would degrade and my colonies would lift off the plate with the matrix and basically be completely destroyed.
There are two main methods for passaging hES and iPS cells: using an enzyme or manually detaching the colonies. Depending on the status of your plate and colonies, one method may be more useful to you than the other. In my experience, when you have fewer than 20 colonies (per well in a six-well plate), it is much better to passage manually. This gives you much more control over what you are detaching from the plate and bringing over to your fresh plate. This should also then be passaged at a 1:1 split unless the colonies you have are pretty large. Even though there are lots of different methods and tools you can use for manual passaging, I’ve found the most effective way to do this is to just use a p200 pipette and tips. This seems to be the perfect size to allow you to score larger colonies into sections while also scraping up the smaller colonies with one scratch. I’ve tried using Pasteur pipettes with the tips curved using a Bunsen burner, but this seems to yield too blunt and irregular tips. I’ve also tried using different-sized needles to break iMEFs off the colonies and score the colonies into smaller pieces, but this often scrapes plastic off the bottom of the plate, getting pieces of plastic mixed into the colony.
If you have over 20 colonies per well, I think it is much easier to go with an enzymatic passage. If you are using a feeder layer, the quality of the colonies may be slightly worse than with manual passaging, because you are picking up the iMEFs in addition to your colonies, which can result in your colonies forming large clumps in the new plate rather than seeding nicely into the new feeder layer. For hES and iPS cell culture, the enzyme used shouldn’t break the cells into a single cell suspension (like Trypsin/EDTA); rather they should be broken into smaller clumps for optimal seeding and growth capabilities. I have used 1mg/ml collagenase type IV (Invitrogen) diluted with DMEM-F12 for passaging with a feeder layer, and Dispase (Stem Cell Technologies) also at 1mg/ml when passaging from Matrigel. Both collagenase and Dispase keep the cells in clumps. Once prepared, collagenase only stores for two weeks, so be sure to not hold on to it for any longer than that; the enzyme becomes weak and ineffective. Even if you are passaging enzymatically, I have found it very helpful to do a little colony cleaning manually beforehand. Getting rid of partially differentiated colonies, breaking up larger colonies into smaller pieces, and teasing away some iMEFS can really make a big difference in the quality of your cells.
When you are removing the cells from the plate after the enzyme treatment, a really effective method for scraping is the “car wash” method. This is done using a 5ml glass serological pipette. While tilting the culture plate slightly forward so that the media forms a pool at the bottom half of the well, you pull up the media, and while releasing the media, scrape in a zigzag pattern from the top of the well towards the bottom. By releasing the media while gently scraping, you help keep the colony-removal process gentle and the cells in bigger clumps. Once you have cleared the top half of the well, flip the plate around so that the other half is on top and repeat the process.
Many people have very different views from mine on the use of antibiotics for hES/iPS culture. I feel very strongly about culturing antibiotic-free, and I will explain why. First of all, it allows you to have more control over the status of your cells. If there is any sort of breach in sterility, without antibiotics, you will immediately know and be able to deal with it by getting rid of the contaminated plates. You never have to live questioning if a plate is infected or not, meanwhile exposing your other plates to the potential infection. Everything is very clear; infections are obvious and therefore can be dealt with swiftly, without jeopardizing the rest of your cells. One of the few times in my iPS culture experience that I was using an antibiotic in my media, after not knowing whether a particular plate was infected, one by one all of my plates became infected and I literally lost every single culture I had. Now, I know that this is probably a pretty extreme case, but in any event, it demonstrated what can happen when antibiotics are battling a bacterial infection. Since the infection was not obvious, I continued to expose my cells to contamination unknowingly and therefore contaminated everything.
Secondly, using an antibiotic can mask mycoplasma infections. Usually, mycoplasma infections are accompanied by other infections—or rather, when the sterility of your cultures is breached, mycoplasma can also be introduced, and they are typically introduced with other infections such as bacteria. If the antibiotic successfully fights off the bacterial infection, your cells will still have the mycoplasma infection, which is typically only detected by using specific mycoplasma detection tests (by taking spent media and testing it). Last year, our iPS facility tested positive for mycoplasma. This was a total disaster. We had to throw away all cell cultures, close down the core for fumigation, and literally throw away all disposable materials in the room including reagents and media. It left us out of commission for a whole month. Not only was this an unbelievably expensive endeavor, it also made us lose valuable time, resources, and in some cases permanently lose cell lines. At the time when this happened, we were all using antibiotics in our media; since then, we have made it a room rule to not use them. Since we have been antibiotic free, we have also been mycoplasma free. Not using antibiotics also helps reinforce good practices in sterile technique, forcing you to be ultra careful with your cells and keep your surroundings very clean. This helps eliminate some variables in culturing, since you have more control over your environment and therefore over culture conditions.
I think one of the most important things to remember with iPS cell culture is to be patient. Especially if you are just thawing cells for the first time! Even if it looks like there are no colonies, I would be willing to bet that if you keep feeding and wait, you will see at least one. Sometimes this can be a slow and frustrating process, but just keep at it and you’ll eventually get some great cultures. I was the first person in my lab to do human iPS cells work, so I truly understand how difficult it can be getting things up and running. There are a lot of helpful resources online, and as the stem cell community grows, the resources grow also. The HSCI iPS core has their protocols available online (http://www.hsci.harvard.edu/ipscore/node/8), which I have found to work well. WiCell has many helpful resources and protocols available online as well. I use their protocol for teratoma assays, and it pretty much works without fail (http://www.wicell.org/home/support/stem-cell-protocols/stem-cell-protocols.cmsx).
So, to wrap things up, if you are new to hES/iPS culture, I hope this has lifted the curtain a bit on culture techniques, hopefully helping to eliminate at least a couple variables while you get started. If you have tips of your own now (or later!), please do share! Pooling our secrets, we can help each other out and make some real scientific progress.
Christine Miller is Research Assistant at Harvard University, Joslin Diabetes Center, Amy Wagers Lab.
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