In my last post, I talked about essential factors you need to consider when designing a protocol for long-term cryopreservation of cells, tissues and other biosamples. However, I left out what may be the most important point of all: Methodology!
How do you actually freeze the material?
Once again, I want to stress how important it is to tune your cryopreservation method to the specific sample type you need to preserve. For example, a protocol that works well forPBMC may not give be the best option for RBC . Cooling rate has to be determined on sample specific basis.
In general terms, there are five methods typically employed to bring a sample down to cryo-level temperatures. Each requires a different set of equipment and can have distinct limitations.
Step-down freezing was popular when I was working in the lab and remains so today. It’s a slow process that is designed to mimic a very protracted supercooling–samples are gently brought down in temperature over the course of many hours or even an entire day –but it can be implemented fairly easily with standard laboratory equipment. A typical step-down process might go from 4°C to -20°C, then to -80°C, and eventually into cryo storage.
There is one significant drawback, however. Step-down freezing does not account for the latent heat of fusion (an increase in temperature as ice begins to from in the sample). As a result, step-down freezing isn’t the best methodology for certain sample types.
Blast freezing is a method designed for speed rather than maximum viability, and it’s used to decrease a specific volume of material by a set temperature in a fixed amount of time. Blast freezing is commonly used for large amounts of material, like blood bags or large volumes of protein.
It’s important to note that there are purpose-built blast freezers designed for this process. A normal ultralow temperature freezer (which is designed to keep a previously frozen sample cold) can’t replace a blast freezer (which is designed decrease a samples core temperature rapidly).
Direct plunge freezing is one of the easiest methods to implement in the lab. All you have to do is immerse the sample in liquid nitrogen or dry ice for a certain amount of time and then place it in a storage vessel. Direct plunge freezing isn’t optimal for sample recovery, though. It often leads to intracellular ice formation, one of the main factors contributing to unviable cells upon thawing. This technique is okay for less critical samples or difficult to replace samples, such as immortalized cell lines.
Slow Freezing using a programmable freezer (or CRF) is the best option for biobanks concerned about compliance with standard operating procedures or accreditation needs. CRFs are controlled from the freezer itself or an external computer and can provide an audit trail detailing how the freeze occurred. CRFs can also account for the latent heat of fusion released as a sample undergoes nucleation and can carefully step down the temperature afterwards to prevent cellular dehydration.
CRFs are widely used by academic and industry labs. They’re also important in the production of vaccines and pharmaceuticals and have helped shape the IVF industry. As I see it, any application looking for consistency, uniformity and documentation can benefit from a controlled rate freezer.
Vitrification as a technique is last on my list because it isn’t as well developed as the other methods for all sample types, although it has been deemed acceptable for preparation of oocytes. Vitrification involves using high concentrations of cryoprotectants to prevent ice from forming at all—it’s kind of like plunge freezing with highly perfused CPA. During vitrification, the sample transitions directly into a glass-like state when cooled. This technique has a lot of promise, but is currently difficult to scale.
For a more detailed discussion about cryopreservation freezing methods, check out my webinar “Avoid the Icebergs” on Thermo Fisher Scientific’s Cell Culture Cafe.