Cook Medical Team Lead, Senior Scientist Steven Mullen’s study on the changes in osmolality that occur in nonhumidified incubators was published in Human Reproduction in 2021, and his research also formed the basis of the presentations and workshops listed below.
For those who were unable to attend the original presentations, we collected the attendees’ questions and the presenters’ answers and grouped them into categories.
How can I measure osmolality?
The easiest way is with an osmometer. We use a Wescor Vapro® 5600 vapor pressure osmometer. Another common method involves the use of a freeze point machine. Others estimate evaporation and then extrapolate the osmolality change by measuring volume or weight changes, but that method may not be very accurate.
Would changing drops (dishes) on day 3 in a dry incubator with one-step culture media mitigate the evaporation?
Changing media on day 3 would avoid excessive evaporation, because the media would experience only 3 days of evaporation instead of 5 or 6 days. This is one way to avoid the issue, but there are others, such as using more oil, using denser oil, etc. Although changing media on day 3 will reduce the amount of evaporation, the osmolality will still increase over the 3 days. Thus, under certain conditions, the osmolality rise after 3 days may be great enough to be detrimental.
Does the osmolality increase with each day of culture?
If evaporation is occurring under the culture conditions that are used, then yes, osmolality would increase each day over time. It depends on the variables of the culture system: humidity, volume, oil volume, oil density, etc.
How many measurements did you perform for each timepoint? How consistent were your measurements?
The number of measurements for a specific time or factor combination varied throughout the study. When we performed such measurements, the values were almost always within 2 mOsm/kg of each other. The most common measurement was +/- 1 of the initial measurement, and it is not uncommon to get the same results again (i.e., the same osmolality when a volume of medium is measured twice).
The starting osmolality of different kinds of media varies. How would this initial difference affect the rate of change?
It shouldn’t affect the rate of change. The absolute values at any time will differ, but the rate of change shouldn’t differ.
Is there any relationship between osmolality and the amount of fungus present, especially in a nonhumidified incubator?
The only way that the presence of fungus might affect osmolality measurements is if the fungus were growing in the medium that was used for the osmolality measurements.
Do direct and indirect heat have different effects on media osmolality?
The only thing that might cause an effect is the time it takes for the medium and oil to warm, and then the rate of osmolality change should be an effect of temperature.
If we put culture media (cleavage, blastocyst) in an incubator without oil in a closed tube, how much does the osmolality change?
In the case of tightly closed tubes, the evaporation is minimal or none, so the osmolality rise won’t be an issue in this case. However, we have to keep in mind that culture media (cleavage and blastocyst media) need to be equilibrated and kept in the environment of gas exchange. This is important for maintaining a proper pH.
The slides you presented show the difference in osmolality change between dry and humidified incubators. Did you evaluate more significant endpoints, like fertilization rate, embryo quality, cumulative pregnancy rate, and live birth rate?
No, in the research we performed, the osmolality rise (mOsm/kg) was investigated in different conditions and culture setup only. Clinical evidence regarding fertilization rates, cleavage and blastocyst rates, and clinical pregnancy rates is available in well-known journals.
What is the relationship between pH and osmolality?
There is no direct relationship. If evaporation is occurring, the osmolality will increase. An increase in pH would also likely be due to the evaporation.
OSMOLALITY AND EMBRYO CULTURE
What were your observations about osmolality change during embryo development from fertilization to blastocyst development, especially in single-step medium?
While most data is from a research setting with no embryos present, there is little reason to think that the evaporation rate would be different if embryos were present in the media. The type of media wouldn’t really affect the evaporation rate (i.e., there should be no difference between single-step and sequential media). If single-step medium isn’t changed over the 5–6 days of culture, it will likely see more evaporation and a higher osmolality increase than a sequential medium that was changed from step 1 to step 2. Changing to a single-step medium is one possible solution for addressing this osmolality change. Another possible solution is adjusting other culture parameters to mitigate the rate of evaporation (adding more oil, changing the type of oil, etc.).
Do you think the embryo can adapt, to some extent, to shifts in osmolality?
Embryos do have a mechanism to regulate cell volume with their inorganic ions or organic osmolytes (amino acids, mainly glycine and glutamine at early cleavage embryo development). The postcompaction embryos (day 3 and 4) have an even more robust mechanism with different osmolytes and transporters. However, we have to bear in mind that this mechanism is able to work only within a narrow range. When osmolality in culture media exceeds 300–310 mOsm/kg, it might impact the embryos’ viability and inhibit embryo development in IVF culture.
Is there any alternative to oil that can be used as an antievaporation agent during embryo culture?
Not that I’m aware of. Mineral oil does help reduce evaporation. There are different types of oil, and some appear to reduce evaporation more effectively than others, such as higher density mineral oils. If the density isn’t listed on the commercial product, it can be difficult to discern which oil is more dense than another, because the nomenclature used to describe commercial oils—light vs. paraffin, etc.—is inconsistent.
What amount of oil is optimal for maintaining proper osmolality in dry incubators without a water jacket?
The answer varies according to the type of dish, the volume of media, the size of the microdrop, the type of oil, etc.
Using too little culture oil could have a negative effect, but could using too much oil also have a negative effect?
Using too much oil won’t have a negative effect, really, other than creating a risk of possible spillage in the lab when you’re walking with the dish. Excessive oil usage may prolong the amount of time required for pH equlibration, but it shouldn’t have a negative effect on osmolality.
How do you recommend that the embryo transfer be performed in order to avoid evaporation of the transfer media? Do you use oil on it?
We use a humidified chamber or a bell jar to cover the dish while we wait and then quickly load the catheter. We don’t use oil. We work quickly and use a larger volume of medium to try to mitigate the risk.
Do you prefer mineral oil over paraffin oil?
This is a hard question to answer, because it’s not always clear as to what the actual difference is. As long as the oil is nontoxic, we don’t have a strong preference and have used both. Nomenclature seems to vary by manufacturer.
We keep a 100 mL bottle of oil in the incubator. Is it harmful for the oil?
Perhaps. Keeping oil too long can lead to potential issues with accumulation of volatile organic compounds (VOCs) or peroxidation. It’s better to adhere to a short shelf life for oil than to store it long term.
Are small bubbles in the oil an indication of evaporation?
Not that I’m aware of. Very small bubbles in oil sometimes seem to be the result of CO2 offgassing when the dish has been out of the incubator for a few minutes. This may be what you are seeing.
Is it acceptable to keep oil in the incubator so that it’s ready for use?
Yes, but you should limit how long it is kept in the incubator, because it can accumulate VOCs or other toxins over time. Keeping oil in the incubator should have no effect on evaporation or osmolality.
Is there is any significant benefit to using washed and humidified oil?
Perhaps for toxicity control, but for evaporation and osmolality control, our data indicate that it provides no benefit.
Have you investigated how the chemical structure of the oil affects its hydrophobic and hydrophilic properies?
No. It would be interesting to investigate. I suspect that the chemical structure of the molecules affects the density of the oil and consequently the rate of evaporation that occurs when oils of different densities are used.
Is it advisable to use oil from one company and culture media from a different company? Will it affect the efficiency of oil?
I see no issues with using supplies from different companies.
Is it beneficial to let oil stabilize in a humid incubator for later use in a dry incubator?
Not that we have seen when we tried humidifying or washing oil. It’s not possible to saturate the oil. Evaporation still occurs at the same rate.
Which is the ideal oil, one with high viscosity or low viscosity?
It is difficult to say which oil is ideal. Higher viscosity oils tend to reduce the rate of evaporation, but only in nonhumidified incubation conditions. Lower viscosity oils tend to be easier to work with, in my opinion, but allow for faster rates of osmolality rise in nonhumidified incubation. For many situations, the value of one variable in a culture system depends on other variables.
Can you give us some brand names of high-density culture oils?
Oils marketed as “heavy” have higher density, generally speaking. I do not wish to name brands, because I don’t want the study to appear biased toward any particular brand or manufacturer.
Did you use washed or unwashed oil, and could this have influenced your outcomes?
I used unwashed oil. Other studies have shown that using washed oil has no effect on the rate of osmolality change.
Which is better to use for culture, paraffin oil or mineral oil?
There is really no difference between the two names. The main reasons companies sell one or the other is that the United States Pharmacopeia (USP) defines the substance as mineral oil while the British Pharmacopoeia (BP) defines the substance as paraffin oil. We meet the specifications for both. Essentially, you are talking abut a by-product of petroleum distillation that is composed mostly of alkanes and cyclic polycarbon chains. Cook Medical’s culture oil is a mineral oil. It is manufactured from oil that meets all the requirements of the USP and the National Formulary monograph for light mineral oil. Additionally, Cook’s culture oil is tested against the European Pharmacopoeia monograph for light liquid paraffin.
How do osmolarity, pH, and temperature differ with regard to the use of an oil overlay in closed vs. open culture systems?
These really shouldn’t differ with open or closed culture in terms of the setpoint that is achieved under proper conditions. Equilibration timing may differ and take longer with an oil overlay in terms of pH. Evaporation, temperature loss, and pH rise would be lower with an oil overlay in a closed system than in an open system. The equilibrium values should be the same. The oil overlay will increase the time it takes for the system to reach equilibrium with its surroundings, both when putting the dish into the incubator and taking it out.
Is drop shape affected by the hydrophobicity of the dish? Won’t the use of hydrophilic dishes result in a different drop shape than hydrophobic dishes?
The geometry of the drop can be influenced by various factors. The amount of oil over the top of the drop and the surface-area-to-volume ratio are likely key factors. So, how drops are made and shaped can be important. The media and dishes I used for my experiment are pretty standard, so the values of the surface area I determined should be close to what others will obtain in their system. However, there are likely to be circumstances, as you point out, that could change this.
What is the lowest culture media volume recommended for a drop culture system?
Impossible question to answer. The issue with very small volumes is that it becomes easy to inadvertently alter media composition through evaporation, etc.
What is your recommendation for proper media droplet size, amount of oil overlay, and type of oil?
This depends on the culture system. Best to measure and see what works. It also likely depends on the number of embryos per drop, etc.
What is an adequate volume of oil for a 50 µL droplet of culture media?
Depends on the drop shape and also the size of the dish. Based on my research, I would use enough oil so that the height of the oil layer was at least 4 mm.
Did you prepare the droplets with the overlay method? How do you avoid the change of osmolality from exposure of the media to air during dish preparation?
I did not use an oil overlay method. I prepared the media on a room-temperature surface and usually outside the hood to avoid air-flow-related evaporation. We also use an electronic repeat pipettor to prepare microdrops, which is very fast. Having said this, I am not concerned about small changes in osmolality during media preparation, simply because that should not affect the rate of change in osmolality, although it will affect the actual value at any given time. My reasoning for designing the study as I did was so that the results can be broadly applicable and not restricted to any specific factor that I chose, such as the medium’s starting osmolality.
Does 1 mL of oil completely cover the microdroplets of medium in a 35 mm dish?
When I was using the 35 mm dishes, I always used 3 mL of oil. Smaller volumes of oil were used in a 96 well dish, and I always used enough oil to cover the surface of the medium.
How did you make the droplets in the 35 mm dish to ensure that the surface area would always be the same?
I was very careful to make the drops consistently. In the study where I measured the drop surface area, I used the same technique that I do for making drops for the rest of the experiment. I used the average value for the drop surface areas that I directly measured when I estimated the surface-area-to-volume ratios for the drops that were not directly measured. It is important to note that I when I developed the model for the experiments, I used 96 well plates that have a cylindrical geometry in the well so that the surface area would be consistent for each treatment. There are variations in the drop size for the same volume of media, but the variability is pretty small relative to the size.
If you compare the difference in surface area of a drop in a well vs. a drop in a flat dish, do you think a culture dish with a well design is better than a flat dish?
When I used dishes with wells, the medium was not in the form of a drop. I ensured that the medium covered the entire surface of the well bottom so it would have a cylindrical geometry. The results seemed to suggest that whether the medium is in a cylindrical geometry, as it is in a well, or in a drop configuration, as is common in 35 mm dishes, the osmolality change will be the same for a given surface-area-to-volume ratio.
How should dishes be prepared to minimize changes in osmolality?
The dish and culture media preparation should take as little time as possible, one dish at a time. The room temperature of the IVF cabinet should be used. The culture media are taken from the fridge before the dish preparation, so a heated surface is not needed. In fact, a heated surface will increase the evaporation of the medium. If you are culturing in a dry incubator, the microdrops should be bigger (50 µL). If you are using humid incubator, the size of the drops doesn’t matter, because the evaporation does not occur as fast as in a dry incubator, so you can use 25 µL drops and still maintain low osmolality until the end of the culture. The layer of oil should be as thick as possible. A 6 mm layer of oil will decrease evaporation more effectively than a 2 mm layer of oil. When you prep the dish, the underlay method is superior to the overlay method, mainly because of the drop shape, so the surface-area-to-volume ratio will be probably smaller, which again will decrease the evaporation risk.
For noninvasive preimplantation genetic testing for aneuploidy (PGT-A), what is the recommended volume of the drop?
This would depend on the specific protocol for the noninvasive preimplantation genetic testing (NiPGT) system that the lab is using. I would comment only to say that they should follow their test protocol for that system.
How many embryos did you place in each microdroplet?
We didn’t use embryos in our research. If we had, I would’ve used one embryo per drop.
After culturing the embryo, should I cover the dish to reduce any changes to the osmolality?
Yes, I would suggest using the lid and covering the dish. The lid provides protection against evaporation and slows the rise of osmolality.
Does the type of culture dish affect the osmolality of culture media?
Numerous different dishes are available in the market. The dish type does not influence the osmolality directly. The amount of media and the amount of oil that is placed on the dish might impact the osmolality, as shown in the presentation.
If humidified incubators yield better results than dry incubators, and this applies to the nonhumidified time-lapse incubators as well, is it advisable to culture in nonhumidified timelapse incubators?
To be clear, it’s not that dry culture cannot work. In fact, many labs are very successful with dry incubators. Nonhumidified incubators can also help reduce issues of contamination. They can have improved temperature stability and environmental recovery. However, culture conditions should take into account the dry nature of the incubator and ensure that evaporation isn’t an issue. Steps can be taken to address it, like using more oil, using a denser oil, altering the media volume, doing media exchanges, etc.
It is typical to classify incubators as dry or humidified, but are all humidified incubators similarly effective? I’ve noticed that humidity can differ, and at different times, the same drop of media without oil will have a different evaporation rate.
Within the same dish, within the same humidified incubator, there really should not be any significant difference in the rate of evaporation between microdrops unless the microdrops are different sizes and aren’t covered by the same amount of oil. Under oil, evaporation is minimal in a humidified incubator. You can use a hygrometer to measure the humidity in an incubator. Different incubators may achieve different humidity levels, depending on their design. There are relatively inexpensive instruments that can be used to measure humidity and temperature. The challenge is whether they are compatible with use in your specific incubator. These instruments need to equilibrate within the incubator environment, and that can take several hours, so you should validate their use in your system if you intend to use them to measure humidity levels in your incubator.
Is there anything I can safely do to add humidity to a dry benchtop incubator?
Trying to humidify a normally dry incubator is risky, because it can potentially damage the internal electronics and may also impact the gas sensors, etc. I believe this is a primary reason why humidity is eliminated in many time-lapse incubators. Humidified benchtop incubators are available. If an incubator is meant to be used dry, then it is safer to adjust the media volume, amount of oil, type of oil, etc., to reduce evaporation rather than trying to humidify the incubator. I would advise against attempting to humidify an incubator that is designed to work without humidity.
38 years ago we were not only adding a pan with tri-distilled water in box incubators but were also, in our lab, using a heater to create a humidified environment for the embryos. We recognize that the oviduct is not dry, so why would anyone incubate in a dry environment?
The embryo is bathed in 100% moisture when in the microdrop, so I’m not sure that the humidity per se is a concern to embryo health. It is debatable whether the embryo is submerged in liquid in vivo like it is in vitro, and this is likely another topic unto itself (i.e., microfluidics, novel surface coatings to culture on, etc.). Although the embryo may experience a more moist environment in the female reproductive tract, the amount of humidity in an incubator does appear to impact evaporation, which can alter the chemical properties and formulation of the culture media, which in turn can impact media efficacy and embryo development.
Has anyone compared the economics of the newer time-lapse culture systems with the traditional culture conditions?
Some have done this. I have not personally. Time-lapse incubators can work well. The real advantage would likely be the use of the time-lapse data for embryo selection. While these are good incubators, other incubators work very well too in terms of environmental recovery and stability.
Of what use is a hygrometer in an IVF lab?
A hygrometer can measure the relative humidity of the environment.
What is your advice for protecting against osmolality increase in dry incubator?
I would recommend that you use the information that is available to assess the effects of different scenarios that you would feel comfortable using in your setting, and use whatever combination of factors results in the smallest rate of osmolality change.
In nonhumidified benchtop incubators, will there be any osmolality changes in one-step media overlaid with oil from day 0 to day 6?
Based on the research done by Steve Mullen, the rise of osmolality is a consequence of using a dry incubator, small culture media drops, and a thin layer of culture oil. The combination of all the above with single-step media will increase the risk of, or even speed up, osmolality rise in the culture.
What is the ideal volume of medium to use to culture embryos for 3 days?
I can’t really answer this question. There are multiple variables to consider. In terms of evaporation, the volume of media is just one variable. The amount of oil that is used and other factors can affect evaporation.
Do you perform volume assessment of cells every day?
We exchange media in our labs and don’t utilize uninterrupted culture. We didn’t adopt our paradigm because of evaporation concerns—we have always used a sequential system that is based on the research done at CCRM and on improved outcomes—but it has helped us avoid this issue of evaporation, since we do use dry incubators.
How can you avoid evaporation of HEPES media in a center well for oocyte retrieval? Cover it with oil?
That would be one way, yes.
Do you think preparing culture media in a cold plate or on top of an ice pack would be a good practice to avoid evaporation?
I don’t think this is needed. Preparing on a room-temperature surface and doing so quickly by limiting the number of dishes prepped at one time before adding oil is sufficient to avoid excessive evaporation during dish prep.
Since there is a change in osmolality with culture time, is it recommended to switch to sequential media rather than single-step media?
Not necessarily. Other methods can be used: humified incubation, increasing the amount of oil, adjusting the media volume, etc. Also, a single-step media can be changed during use.
Do you prefer to subculture on day 3 even after using single-step medium?
We exchange media in our labs and don’t utilize uninterrupted culture.
Single-step culture is not a very old technique. Is any data available on the safety of this culture system and the development of babies born as a result of it?
It can work very well, and there is no clear consensus on the superiority of one media type over another (sequential vs. single step).
Should we use higher volumes of media to avoid changes in osmolality during preparation, as long as we are not working in time lapse?
This may help. However, when you’re working in a humidified incubator, the osmolality change over time is quite small. It is probably a good idea to incorporate many processes into your protocol in order to reduce osmolality change.
Have you looked at the impact of protein supplements on osmolality?
No, we didn’t. During the experiments we used Cook Medical’s Sydney IVF Cleavage Medium, which contains human serum albumin.
What is the most important component in media that affects embryo development?
Cook Medical’s sequential culture media are composed in a way to support embryo development at each stage of embryo growth. The formulations are designed for specific stages of development, and all components are added for the combined effect. The most important of these components—the ones that support embryo metabolism—are carbohydrates and amino acids.
With the emergence of time-lapse culture systems, the current trend is toward the use of single-step medium for extended uninterrupted culture. Why is Cook Medical not developing a single-step medium to address this market need?
Our belief is that the most important need to meet is that of the developing embryo. The more recent trend toward single-step systems is driven largely by static culture systems (time lapse) and the efficiency and convenience of the lab. We believe that a sequential media system is the most appropriate system for supporting embryo development, because sequential media systems mimic nature. Sequential media are based on the natural changes that occur in the oviduct and the uterus during embryo development.
How often should one change to fresh culture media to maintain a normal osmolality?
In the case of sequential media, the medium and the dish are prepared for 2–3 days only, and after that, the dish will be changed and the medium renewed. In the case of single-step media, the dish will be kept 6–7 days. The experiment done by Steve Mullen shows a rise of 6.96 mOsm per day in a microdrop setup. Depending on the culture medium’s osmolality, the amount of culture medium used, the amount of oil used, and the type of incubator, you might be able to predict the time when the osmolality will be exceed the set limit. To avoid an osmolality rise above 30 mOsm/kg, changing the media after 2–3 days of culturing should help maintain a safe range.