Debbie Knight

A Day in the Life: March 1, 2012

In research log on March 1, 2012 at 9:00 am

From time to time, I will give a glimpse into the “glamorous” life of a research associate and talk about what I’m doing in the lab on a particular day. These entries I will call “A Day in the Life…”

Every once in a while, I have to make and sterilize cell culture media and buffers to use in the lab. Some media we buy already made — these are the more difficult-to-make media and it’s just easier to buy them than to make them ourselves. But I make quite a few in our lab — it’s often more economical to do so.

Since not everyone out there may know how this process is done, I thought I’d take you through the journey. In many ways it’s like working in your kitchen at home, only you get to wear “cool” lab safety gear such as a lab coat, safety glasses, gloves, and closed-toe shoes to avoid injuries. It’s not like similar dangers lurk in the kitchen setting, it’s just Occupational Safety and Health Administration (OSHA) doesn’t regulate how you run your kitchen at home. Knife throwing while wearing sandals at home? Sure! Knife throwing in the lab? No way! Not without protection!

So, today I’m making two cell culture media and a buffer. And there will be two ways in which I sterilize these solutions. The buffer, phosphate-buffered saline (PBS), is a commonly used solution in a cell culture lab. It is basically a solution of various salts combined in such a way to stay at a neutral pH — a pH that cultured cells can tolerate. There is nothing in the PBS solution that will be harmed if heated during the sterilization process, so I will be making the solution, distributing it into glass bottles, and steam sterilizing it.

Now, cell culture media contains several components that are sensitive to heat (such as glucose which could caramelize or amino acids which break down) . In this case, the media will have to be sterilized through a filter (which looks a lot like a piece of paper) that has a very small pore size — small enough to filter out any bacteria and fungus that might be lurking in the water used to make the media, but large enough to allow the liquid to flow through.

So, first I’ll talk about making PBS. And steam sterilization.

In our lab, we have a 3-ring binder with protocols and recipes — I call it our little black book.

My lab's "black book" of lab protocols and recipes

The largest component of the PBS solution is what most of us call “salt” — sodium chloride.

Sodium chloride (or table salt) is the main component of our phosphate-buffered saline

So, the first step is to measure it out. I’m making five liters, so I’ll need 40 grams of sodium chloride. I’ll also need 1 gram of potassium chloride, 5.75 gram of sodium phosphate, and 1 gram of potassium phosphate.

Measuring out sodium chloride on the analytical balance

So in the photo above, I’ve only measured 26.7203 grams of sodium chloride. I’m shooting for 40 grams, so clearly I’ll have to continue adding more to the weigh dish. Many labs differ on how close you have to come to the 40 gram mark — obviously, 40.0000 grams would be ideal, but in reality it’s difficult to get that kind of precision (one grain and you could go over the target weight). With a solution for cell culture, it’s not that critical to measure something that precisely. When I’m measuring chemicals for cell culture purposes, I shoot for accuracy to the second decimal place (i.e., 40.00 grams), so if the balance reads something like 40.0029 grams, I’m okay with that. As we say, tongue-in-cheek since our funding comes from the government: close enough for government work. (No disrespect to the hardworking American taxpayers out there)

After measuring out all the chemicals, one at a time, and placing them in a container large enough to hold  five liters of liquid, I make sure I clean off the balance. This is important! If chemicals, especially salts, are left on the metal weighing pan, they can corrode it. It ain’t pretty and is not the proper way to take care of this delicate instrument. In the first lab I worked in at this university, I was constantly reminding my lab mates to clean the balance. The balance looked pretty good when I left to work in another lab. Within a couple of months (without me to apparently remind them to clean the balance), the weigh pan was corroded and badly pitted from all the chemicals that were left behind due to poor lab hygiene. (Bad, former lab mates, bad!)

Cleaning the weighing pan on the analytical balance is important to remember (and yes, it is a make up brush!)

So, after weighing the chemicals, adding water to the container is next. Once the water is added, the solution is stirred with a teflon-coated magnetic stir bar that is placed inside the container. The mixing helps the chemicals dissolve as well as to disperse everything evenly.

Adding water to make the buffered solution. The mixing container is placed on a magnetic stir plate. A magnetic stir bar placed in the container is used to mix the solution.

Now, let’s talk about water purity. Tap water will do in a pinch, but it may have things in it like chlorine and fluoride, not to mention trace minerals, pesticides, and other contaminants. These can vary from day to day, so if you use tap water, be mindful that it could affect the welfare of your cell cultures. The next grade of water in our lab is deionized water which flows from a faucet next to the regular tap water. This is okay, but I prefer water I use to make lab reagents to be a little more purified, so I use water that has been double-distilled. I currently order our water from a university lab store, but our department will be putting in a reverse osmosis water purification system as a shared resource soon. (A plastic jug this water comes in was fashioned into a mixing container in the above photo — re-using/recycling)

Decanting water from our "bottled water" supply

The pH of the solution, once all the chemicals have dissolved, is important. The cultured cells can only tolerate a narrow pH range, all hovering around a neutral pH (i.e., pH 7.2). More often than not, the pH will have to be tweaked using an acid solution (generally hydrochloric acid) or a basic solution (generally sodium hydroxide). I say “generally” here because there are some solutions that require different acid/base pairings for pH adjustments.

Hydrochloric acid is often used to adjust the pH of lab solutions

So the way pH is measured is using an instrument called a pH meter. Our meter needs to be calibrated before each use to ensure it is working properly and that our pH measurement is right. The probe goes into the solution and relays information to the meter in real time, so as you adjust the pH with an acid or base, the numbers on the meter will change. In this case, we want the pH to be 7.2, which after some “magic” with the acid/base solutions, the meter will read as such.

pH probe measuring the solution's pH. The meter gives the read out for this measurement.

pH meter reading of 7.2

Because I made this solution in a plastic container, there’s a chance that the magnetic stir bar has shaved off a few flakes of plastic. This is undesirable in the PBS solution. So, I pour the solution through a few layers of gauze to filter out such debris.

Filtering out debris while dispensing solution to glass bottles

Once the solution is dispensed into the glass bottles, the caps are loosely threaded into place. The caps need to be loose because during the steam sterilization process, the bottle, cap and the liquid contents expand with heat and pressure. If the lid is on too tightly, the bottle may burst (no where else for all that pressure to go). If the lid is on too loosely, the cap may blow off due to the pressure within the bottle. I generally tighten them down and then unthread them a half turn or so. Ideally the cap should jiggle without falling off.

I then put a piece of aluminum foil over the top of the bottle. This acts as a dust cap when the bottle is stored for use, keeping out stuff that may settle out of the air. Many labs use a tiny piece of foil that just covers the lid. I prefer the foil to cover the lid and the “neck” of the bottle. I should note that there is the rare lab that do not use any foil at all because they believe that small amounts of the aluminum leach into their solutions during the sterilization process and affect their cell cultures. This could be true, I suppose, but I have not observed any noticeable problems with our cell cultures using the foil dust caps.

Some labs use a foil cover that shields the cap and the neck of the bottle from dust (left). Some labs prefer to use just enough foil to cover the bottle cap (right). I prefer the style on the left when storing cell culture solutions

A lab that had used the autoclave before me prefers to cover only the lid with foil -- some of the coverage is not complete as evidenced by the glimpses of the black bottle cap. This is not how I learned to do this.

In the photo immediately above, you’ll notice that there is a piece of brown and off-white tape stuck to the aluminum foil covers. This is what we call “autoclave tape.” The tape originally looks like masking tape (see photo below). As the tape is exposed to heat or steam, there are thin heat-sensitive bands that turn brown. In the lab, we use the tape to help identify glassware and solutions that have been sterilized using a steam sterilizer (or autoclave). (I had one student who actually thought we drew the brown stripes on the tape with a marker). While the brown stripes tell you the tape has been exposed to heat, it does not tell you whether the autoclave cycle ran correctly or whether the contents are truly sterilized.

Autoclave tape is added to the tops of the bottles

One other thing that should be mentioned here. Bottles with liquids in them should be placed in an inch or so of water in a pan. This helps disperse the heat around the bottle and, should the bottle burst during the autoclaving process, the bottle’s contents and any broken glass will remain in the pan rather than muck up the inside of the steam sterilizer and possibly clog the small drain at the bottom of the sterilizer.

Because I made five liters of PBS and I dispensed it into ten 500-ml glass bottles, this is a little too heavy for a plastic autoclave pan so I used a metal pan. Again, there is about an inch of water in this pan to help disperse heat during the sterilization process.

The autoclave is down the hall from my lab, so I have to transport it. I could carry it by hand (but it’s heavy and bulky). I prefer to use a cart — mostly for the return trip when pan and bottles are really hot.

Transporting the bottles and pan to the autoclave room on a cart

Steam sterilizers, or autoclaves, vary greatly.  They can be old school, where you batten down the door using a crank. The one I use is pretty automated — you just load it, select the program you want, and go.

Getting ready to place (or "load") the pan into the autoclave

Loaded!

With the autoclave door closed, the sterilization cycle can begin

After the door is shut, you select an autoclave program. Selecting a program on this autoclave is pretty simple. We only have two choices. A cycle for “liquids” and a cycle for things like glassware or pipet tips called “gravity.” Both programs bring the autoclave chamber to a pressure of 18 pounds per square inch and a temperature of 122 degrees Celsius using steam. They both hold that temperature and pressure for 20 minutes. This should be adequate to kill most bacteria and fungus that might be in or on the stuff being sterilized. I say “most” because some bacteria form what is called an endospore (like a seed) that is not harmed at this temperature or pressure. If the endospore has the opportunity to come out of stasis, then the bacteria can grow in or on the supposedly sterilized stuff.

But I digress. The difference between the two autoclave programs is how they vent the steam inside the autoclave chamber. If you have liquids inside the bottles, you don’t want to vent all the pressure at once. If you do, the liquid will essentially “boil” off and rather than 500 ml of solution in a bottle, you might have only half that remaining after a fast venting (believe me, I’ve done it, that’s what happens). You want the pressure to slowly be released when you’re doing a liquid load. So, if you select the “liquid” program, it stops the flow of steam and lets the contents of the chamber (i.e.,your liquids) cool slowly. If you just have glassware (and no liquid inside that glassware), you can use the “gravity” cycle. This cycle vents the steam/pressure pretty quickly (and makes a pretty remarkable sound sometimes as it’s venting).

The cycles (or programs) for autoclaving

Starting the cycle with the push of a button

Once at proper pressure and temperature, the cycle counts down the time

After the cycle is complete, this autoclave gives an audible buzz as well as a visual notification.

Autoclave cycle is complete

Pressure gauge indicates there is no pressure in the autoclave chamber

At this point, the door can be opened. But care should be taken — there is still a bit of steam that will escape when you open the door. I can’t tell you how many times I have forgotten this and gotten a minor burn on my forearm or singed my hair a little by not remembering this little tidbit.

Also, you should don a pair of autoclave gloves (kind of like oven mitts) before removing the pan of autoclaved stuff. The gloves we have are a very bright orange and make quite a fashion statement, let me tell you!

Wearing the very fashionable autoclave gloves when removing hot pan from the steam sterilizer

Now, many labs feel the stuff that just came out of the autoclave is hot enough to thwart any possible contaminants from blowing under the cap and into the glass bottles. I’m more paranoid and leave nothing to chance, especially when it comes to cell culture reagents. I either use foil or disposable bench paper to cover the pan when I transport it from the autoclave room to my lab. Like I said, I’m a little paranoid, so this is probably overkill.

Covered pan before transporting sterile bottles through the hallways to the lab

Once in the lab, I need to transfer the bottles out of the one-inch layer of water in the bottom of the pan and onto the lab bench so after the bottles cool, they are dry so I can apply labels to them. Again, I’m a little paranoid, so I use bench paper to seal them in until I can tighten the bottle caps.

Cooling bottles, all tucked in and protected against dust

Once the bottles have cooled, I tighten the caps and throw on some labels. PBS is a clear solution so it could be confused with sterile water or another buffer. I indicate the date the PBS was made on the label, so if there’s problems with the batch, I can find all the bottles from the batch and dispose of them. Of course, I’m so paranoid, that rarely is there a problem with our PBS. But you never can be too careful.

Adding labels to sterilized bottles of PBS

The bottles are then stored on a shelf. PBS can be stored at room temperature or in the refrigerator for quite a while. The five liters I made will probably only last a couple of months before I have to make more.

So, that’s how you sterilize a buffered solution. But what about culture media?

The first step for sterilizing culture media is to have freshly steam-sterilized glass bottles. I’ll be filter-sterilizing media into these sterilized bottles after the bottles have cooled.

In this case, the empty glass bottles are light, so I can use a plastic pan instead of the metal one.

A word about plastic and the autoclave — not all plastic can take the heat! You have to be careful that you use the right kind of plastic or you could end up with a bit melted plastic mess (and believe me, it’s happened on more than one occasion in the labs I’ve worked in). When this happens, I call it “autoclave art.” Some of this “art” turns out pretty cool — almost frame-worthy art. But more often, it just oozes throughout the autoclave chamber and has to be chiseled off. That’s not pretty. The best advice I can offer? When in doubt, don’t put it in the autoclave. Or be sure to have a tried-and-true autoclavable pan to hold the questionable plastic item during its maiden voyage in the sterilizer. One problem with plastic autoclave pans is that after many autoclave cycles, it becomes brittle and discolored. The pan in the photo below has been through several cycles and the handle is cracking.

Plastic pans, after several cycles in the autoclave, become brittle and crack (note cracked handle)

Sterilized empty bottles coming out of the autoclave

Note that the pan I’m removing from the autoclave in the photo above is newer — it’s not discolored or brittle … yet.

Once the bottles have cooled to room temperature, I make the media. We purchase powdered media from Life Technologies (formerly known as Invitrogen which was formerly known as Gibco), but there are other companies out there that make powdered media as well. My lab was locked into this company’s media long before I arrived on the scene.

So the first step, simple enough, is dumping the contents of the little packet into a container. Each packet makes one liter of culture medium. This particular medium I need to add a little sodium bicarbonate to help buffer the solution and hold it at a neutral pH.

Pouring powdered media into a mixing container

I’m making two different kinds of media. One is called DMEM (Dulbecco’s minimal essential media) and RPMI-1640 (it’s named after the institute it was made at: Rosswell Park Memorial Institute). These two media have components in which different cultured cells prefer to grow.

Both of these media have a chemical dye called phenol red which acts as a pH indicator. It’s a “red” color when the pH is neutral (like pH 7). It turns “magenta” (or deep red) when the media is more basic (like pH 8 or higher) and “orange” when the pH is acidic (like pH 6 or below). This indicator helps you tell if the media that’s been sitting on the refrigerator shelf is okay to use or not. It also tells you what sort of environment the cultured cells are growing in. If there is bacterial contamination or the cultured cells are packed in the culture flask, the media may turn “yellow” or “orange.” The cultured cells are not going to be happy for long under these conditions. If the incubator in which the cells are grown is not set up properly, the media may turn “magenta.” Again, the cultured cells will not be happy for long. So, pH is important and this is why the indicator dye has been added to the media.

The pH indicator dye (phenol red) in these media turns "orange" under acid conditions (when pH is below 7) like the container on the left or red when pH is more neutral (when pH hovers around 7) like the container on the right

When making the media, a pH adjustment using an acid solution like hydrochloric acid or a base solution like sodium hydroxide is used.

The pH of this cell culture medium has been adjusted to 7.2 (This was the container on the left in the previous photo, note the change in color)

Once the solution is at the appropriate pH, it can be filter sterilized. This filter comes sterile and has a pore size of 0.2 microns — small enough to prevent bacteria, fungus, and some viruses from passing through, but large enough to allow the now-sterile liquid to pass through. This sterilization process is done inside a biosafety cabinet which circulates hepa-filtered air to provide a sterile environment.

Filter-sterilizing culture media in the biosafety cabinet. This filtering system uses a vacuum pump to pull the medium (red) through the tubing and into the filter unit (blue). The red fluid inside the glass bottle has passed through the filter and is considered sterile.

Filter sterilizing cell culture medium, DMEM

As I indicated before, I am making two different media. One difference between DMEM and RPMI-1640 is that RPMI-1640 has less phenol red in it, making it appear as more of a salmon-pink color rather than a Kool-Aid red. By the way, even though the media looks like Kool-Aid, I can assure you it doesn’t taste like it — it tastes salty. How do I know this? Well, I accidentally splashed some on my lip once.

Two media, filter-sterilized. RPMI-1640 on the left is more salmon pink colored than the DMEM on the right because it contains less phenol red (a pH indicator dye)

I number the bottles in the order they are filtered because I also make a little tester bottle to assure that the contents of this batch are sterile. The numbering helps me keep track of which bottles are sampled in which tester. Why do a sterility test on media that was just filter sterilized? Well, sometimes in handling the bottles and the filter unit, contamination can be introduced. Also, sometimes the filter has a defect or there’s a surge in the vacuum pressure that causes a small tear in the filter — this can also lead to contamination. The media that cultured cells grow in is very rich — and if a bacteria and fungus are present, they can grow like crazy. They love the stuff. So, it is important to be sure that the culture medium is free of such contamination. Testing a small sample of the culture medium helps to ensure that the entire batch has been properly sterilized.

After the caps are tightened, the bottles of media are stored in a walk-in refrigerator. I use different colors of labeling tape to quickly identify different media formulations. Yellow for DMEM, aqua for RPMI-1640, green for M-199, orange for B-medium. Culture medium has a limited shelf life, so it is made in smaller batches than buffers (which have a longer shelf life).

Cell culture media stored in a walk-in refrigerator. Different colored labeling tape helps quickly identify different media formulations

And there you have it. How to sterilize cell culture solutions in the lab. Exciting, I know.

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  1. […] For more on autoclaves, see my previous post […]

  2. […] For more on autoclaves, see my previous post […]

  3. Hi, it’s good to know there are people as paranoid as I am… I keep my medium and PBS bottles in individual plastic containers and only open them inside the biosafety cabinet, the problem is finding containers of suitable size…

  4. seems a bit over the top to waste all that aluminum foil to be honest. I never do that and never get contamination.

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