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Mass photometry FAQs

What can mass photometry do?

At its simplest, mass photometry returns information about the distribution of mass species in a sample. This can be used to assess the monodispersity of your sample, conversely the distribution of oligomers in the sample.

How does it really work, though?

Mass photometry is a method in which changes in the refractive index of a liquid medium (the diluent) as biomolecules within the medium impact a glass surface are detectable using a laser microscope that is focused on that glass surface. These changes in refractive index manifest as shadows that are recorded, detected, then related to masses using calibration curves generated using protein stocks of known MW.

Can the technology tell me what portion of my protein is monomer, dimer, etc.?

You can get reasonable estimates of the distribution of oligomer species in a sample if you can reasonably eliminate unbinding events. Some explanation: biomolecules don’t usually bounce off the glass. They will impact and bind, then after some reside time, release and unbind. The software can tell binding events from unbinding events. Often, a molecule will bind and not unbind. This is what you want to get the best possible assessment of oligomeric species by percentage because otherwise, you have some species recycling themselves back into solution to create new binding events while other species remain adsorbed to the glass. Any mass species with rapid binding/unbinding will be overrepresented in the data.

If you need this type of assessment and observe considerable unbinding events, you should try different diluents. Try adjusting the pH up or down, try adjusting the salinity, and so forth until unbindings are reasonably eliminated.

What’s the range for detection?

Our instrument has a range from 40 kD to 5 MD. It can return viable results slightly below and above those endpoints for well-behaved, highly pure samples. “Well-behaved” means soluble, globular, folded, not highly charged, and existing in discrete mass species.

How much protein (biomolecule) do I need?

The final concentration in each experiment ranges from 0.5-5 ug/ml or around 1-100 nM. That is for a 10-20 ul volume. That works out to only 20 ng per experiment. Typically people will bring a few 10s-100s of ul of their sample at 0.1-1 mg/ml, then dilute 1:100 or more.

The importance of the final concentration of the sample during data acquisition cannot be overstated. It is not merely a matter of choosing whether to observe fewer or more individual impacts. With too few impacts, you lack confidence in the data. With too many impacts, the data become muddled, and the curves for the mass species overlap in the resulting histogram.

What buffers are compatible?

The technology is compatible with a wide range of buffers and salts. We have found that cryoprotectants (e.g. glycerol, sucrose) and detergents (e.g. Tween 20, Triton X-100) increase the background noise. Sometimes that background is too high and kills the experiment. If you must have any such constituent present, use the highest quality you can find.

We have also found that very high concentrations of both usual and unusual buffer reagents, e.g. >2 M salt, >4 M urea, >25% glycerol, can affect the data quality as well as the very ability to focus the instrument.

We like to start with PBS, TBS, HBS, etc. We keep them on hand for user convenience. If you have some other buffer that your buffer requires, bring a few ml of it.

Should I filter my own buffer?

Yes that is generally helpful, although the bigger factor is using better constituents to make your buffer in the first place. There have been occasions that users brought buffers that were made with impure ddH2O sources. We will be able to see immediately if a buffer is suitable for the technology.

Should I filter my sample?

Yes that is generally helpful.

How can I prepare for doing mass photometry?

Don’t make any dilutions ahead of time.

How long does it take to get data?

Once you are trained on the instrument, you can prepare some slides in a matter of 20-30 minutes, then get your first data inside of 10 minutes. We urge users to run replicates as well as multiple dilutions. So a single sample can quickly turn into 5-10 reads or more.

Can I analyze proteins that have concentration-dependent oligomerization effects?

We can try such samples, however they can be tricky. As noted above, the final concentration in each recording is roughly 0.5-5 ug/ml or 1-100 nM. If the effect you aim to observe is within that range, it will collide with the ability to actually record the necessary data.

Can I analyze fibers with mass photometry?

We have tried and it does not work. Detectable impacts are generally round in shape. Fibers impact the glass and cause all sorts of shapes, none of which are round. So their impacts are not single and detectable, and they cannot be related back to a mass.

Can I analyze molecular cages with mass photometry?

We have tried and it does not work. The impacts of the molecules on a glass coverslip are dependent on the mass of the molecule, not its size. So molecular cages will make recordable impacts, sometimes very clean looking impacts, but they will be analyzed as having a mass related to the molecules that compose them, not a mass directly relatable to the volume they enclose. For the experienced: when we ran a series of increasingly large molecular cages, we saw practically the same contrast values for all of them. This resulted in a calibration curve that was a straight vertical line.

How much does it cost?

We prefer to discuss this in person. Suffice to say, in this core lab, we key the cost of usage to the number of silicon gaskets used. Each gasket sheet provides 24-28 individual reads. How you use those reads is up to you – on various sample dilutions, replicate reads, different samples, different diluents, and so forth. You should also reserve at least one read to run a set of MW standards to make your own calibration curve, especially if you have a buffer containing some agent (e.g. glycerol) that might shift the apparent MW of everything upward or downward.

Can I analyze nucleic acids?

Nucleic acids tend not to make good impacts on the glass owing to their charge. That charge can be neutralized with the addition of spermidine. We keep 20x stocks of spermidine on hand for users at no extra charge. You can also try coating the glass with poly-lysine. We also keep poly-lysine solution on hand, though we have not had much luck with that technique.

Can the instrument calculate binding kinetic figures?

It can calculate KD figures but only for tight complexes such as antibody-antigen binding in the nM range. Refer to Sonn-Segev, Belacic, et al. 2020 Nature Comm 11: 1772 for more information. We have not (yet) done this in our core.