This is a post by Christine Miller, Research Assistant at Harvard University, Amy Wagers Lab:
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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.
Passaging
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.
Antibiotic Use
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.
Last notes
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.
Note: StemCultures facilitates posting on this blog, but the views and accounts expressed herein are those of the author(s) or interviewee(s) and not the views or accounts of StemCultures its officers or directors whose views and accounts may or may not be similar or identical. StemCultures, its officers and directors do not express any opinion regarding any product or service by virtue of reference to such product or service in this blog.
Cell culture media is composed of a complex mixture of salts, carbohydrates, vitamins, amino acids, metabolic precursors, growth factors, hormones, and trace elements. Formulations range from simple, basic mixtures containing the minimum requirements for growing many cell lines to complex serum-free mixtures specific for growing a single fastidious cell line. Carbohydrates are supplied primarily in the form of glucose. In some instances, galactose is used in place of glucose as it is metabolized at a slower rate which decreases the amount of lactic acid build up. Other carbon sources include amino acids (particularly L-glutamine) and pyruvate.
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ATCC Media Brochure
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Most cell culture media are sold as basal media. Basal medium is a complex mixture of salts, carbohydrates, vitamins, amino acids, metabolic precursors and trace elements. To enhance cell growth, you must add growth factors, hormones, and other proteins to the basal medium. Often this is accomplished by supplementing the basal medium with serum. Serum should be added to the basal medium at the concentration specified on the product sheet for the cell culture you are growing.
In some cases additional supplements must be added as indicated on the cell culture's product sheet or in the catalog description. In general, if the medium formulation states that the medium has been adjusted to contain certain components, these components have already been added to the medium at the stated concentrations. However, if the medium formulation states that it is to be supplemented with certain components, then those components must be added to the medium by the user.
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ATCC Animal Cell Culture Guide
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Sodium bicarbonate is a buffer used to stabilize pH. Cells in culture produce CO2 but require only small amounts of the compound for growth and survival. CO2 affects the pH of medium. Increasing atmospheric CO2 decreases the pH of the medium. Decreasing the atmospheric CO2 increases the pH of the medium. As cells metabolize, they produce CO2 thus causing the media to become more acidic. If the medium contains phenol red, the color of the medium becomes more yellow. In culture media, dissolved CO2 is in equilibrium with bicarbonate ions and many cell culture media formulations take advantage of this CO2/bicarbonate reaction to buffer the pH of the media. CO2 dissolves freely into the culture media and reacts with water to form carbonic acid. As the cells metabolize and produce more CO2, the pH of the medium decreases (becomes more acidic).
Sodium bicarbonate, NaHCO3, is used as a buffer. Sodium bicarbonate dissociates into sodium and bicarbonate ions. By increasing the bicarbonate ions, the buffer drives the top equation to the left and thus increases the pH. The concentration of the sodium bicarbonate in the medium must be matched with the level of CO2 in the atmosphere above the medium. For media containing 1.5 to 2.2 g/L sodium bicarbonate, use 5% CO2. For media containing 3.7 g/L sodium bicarbonate, use 10% CO2. If the concentration of sodium bicarbonate is too high for the CO2 atmosphere in the incubator, the media becomes more alkaline (the pH increases).
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ATCC Animal Cell Culture Guide
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Some cell lines may be maintained satisfactorily on an alternative medium such as:
You can usually determine if a medium is satisfactory by using it with the cell line in question for 3 to 5 passages. However, cultures established at very low concentrations (e.g., cloning) usually require CO2 in the gas phase. An alternative to using a CO2 incubator is to have a 5% CO2 gas tank at your work site and stream filtered CO2 into the gas phase above the medium prior to sealing the flask.
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ATCC Animal Cell Culture Guide
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Insulin is a polypeptide hormone found in both vertebrates and invertebrates. It is a growth hormone that promotes uptake of glucose and amino acids. The Dictionary of Cell Biology defines insulin as:
1. Hormone found in mammalian serum.
2. Secreted by B cells of the pancreas in response to high blood sugar levels.
3. A mitogen-activator, which is any chemical specifically stimulating a eukaryotic cell to enter S phase of the cell cycle, that commits the cell to G2 and mitosis.
4. Has sequence homologies with other growth factors.
5. Frequent addition to cell culture media for demanding cell types.
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Antibiotics and/or antimycotic agents are added to cell culture media as a prophylactic to prevent contamination, as a cure once contamination is found, to induce the expression of recombinant proteins, or to maintain selective pressure on transfected DNA.
ATCC does not use antibiotics or anti-mycotics for routine cell culture. Long-term use of antibiotics or anti-mycotics may mask the presence of low levels of microbial or mycoplasma contamination. In addition, some antibiotics and anti-mycotics are toxic and may affect the recovery and proliferation of some cell lines.
However, one may elect to introduce antibiotics for short periods to primary cultures or as a safeguard while propagating specific valuable stocks to produce working stocks. If you do elect to use an antibiotic in your medium, ATCC recommends using a Penicillin-Streptomycin solution at a final concentration of 50-100 I.U./ml penicillin and 50-100 µg/ml streptomycin. ATCC offers a Penicillin-Streptomycin solution (ATCC® 30- &#;). This sterile solution can be added at 0.5 to 1 ml of solution per 100 mL of cell culture media for a final concentration of 50 to 100 I.U./mL penicillin and 50 to 100 µg/mL streptomycin.
While ATCC avoids use of the following two agents, commonly used concentrations are as follows. Gentamicin sulfate is an antibiotic and is used at 50 to 100 µg/ml culture medium. The anti-mycotic amphotericin B is used at 2.5 µg /ml culture medium. See Chapter 7 in Methods in Enzymology: Cell Culture, () Vol. 58, W. B. Jacoby and I. H. Pasten, eds. (Academic Press, New York).
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ATCC Animal Cell Culture Guide
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Many factors make serum-free media an attractive alternative to media containing serum:
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ATCC Media Brochure
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Conditioned medium is spent media harvested from cultured cells. It contains metabolites, growth factors, and extracellular matrix proteins secreted into the medium by the cultured cells. Examples of each might include: metabolites such as glucose, amino acids, and nucleosides; growth factors such as interleukins, EGF (epidermal growth factor), and PDGF (platelet-derived growth factor); and matrix proteins such as collagen, fibronectin, and various proteoglycans. Most cell lines requiring conditioned medium are dependent on at least one major constituent found in the conditioned medium
For example, one of the major growth factors present in LADMAC conditioned medium is CSF-1 (colony stimulating factor 1). CSF-1 assists with macrophage progenitor proliferation and differentiation. LADMAC conditioned medium is obtained from the LADMAC cell line ATCC® CRL- &#; and is added to the growth media used to propagate ATCC® CRL- &#;, ATCC® CRL- &#;, ATCC® CRL-&#;, and ATCC® CRL-&#;, all of which express colony stimulating factor 1 receptors.
Mouse embryonic stem cells (ESC) also benefit from growth-promoting components found in conditioned medium. LIF (leukemia inhibitory factor) is secreted into medium by near-primary cultures of mouse embryonic fibroblasts (MEF), and is an essential component in maintaining the undifferentiated state of ESCs. For example, ATCC® SCRC- &#; requires both the use of a mouse embryonic fibroblast feeder layer, as well as additional supplementation with purified LIF (Chemicon, ESG) to maintain the undifferentiated state of the embryonic stem cells.
It is important to bear in mind that omission of conditioned medium from a cell line requiring this type of supplementation should be carefully evaluated. While most cell lines are, in fact, dependent on at least one major constituent found in the conditioned medium, replacing the conditioned medium with a recombinant, or purified, form of this one, single growth factor may yield unfavorable results such as altered expression phenotypes, slowed growth, quiescence or even complete loss of the culture. Any of these responses would indicate that the conditioned medium contains more than just a single growth factor essential to the healthy maintenance and growth of the cells (Freshney RI, ).
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ATCC Animal Cell Culture Guide
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ATCC generally lists in the product description catalog and product sheet either the medium recommended by the originator of the cell line or a standard medium formulation that has been found to be effective otherwise. Media formulations vary widely among commercial suppliers even for media with similar, if not identical names. A change in media or absence of an additive from the recommended formula could affect growth, recovery and/or function of the cell line. It is very important to read catalog descriptions and media bottle labels carefully since media mix-ups are a leading cause of cell culture problems. On the ATCC website, the product page of the cells, in the "Culture Method" tab, you will find the recommended media listed for the cell line.
The formulations to the ATCC media can be on the ATCC website, www.atcc.org on the media's product page. Or you may contact media manufacturers for complete formulations of the most common cell culture media.
Hyclone media formulations of the most common cell culture media can be found at https://promo.gelifesciences.com/gl/hyclone/products/classical-liquid-media.html
ATCC Animal Cell Culture Guide
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Whether you add medium or split your suspension culture will depend on the optimal density range for logarithmic growth, the number of cells in your flask and the amount of medium needed to reduce the cell density. To determine how much medium to use, follow these steps:
For example, assume your suspension cell line should be maintained between 2 x 105 and 1 x 106 viable cells/ml. If your medium volume is 10 ml, and your cell count is only about 3 x 105 viable cells/ml, then, you have several choices.
If you chose option #3, your vessel would now hold 15 ml of medium. Depending on the vessel size, that is probably okay. However, if you do your cell count and determine that the cell density is about 1 x 106 viable cells/ml, then you would have to add 40 mls to bring the cell density back to 2 x 105 viable cells/ml. This would result in a total of 50 ml and the culture would probably have to be split into multiple vessels.
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ATCC Animal Cell Culture Guide
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