what you do in this channel is amazingly inspiring, and you've changed the lives of thousands of scientist-to-be from all over the world. lots of love and admire from istanbul. we can't thank you enough!
Hello from Bangkok! Firstly, thanks for all your wonderful videos! I’m a ‘hard’ materials crystallographer trying to learn biochemistry. I really appreciate all the hard work that you have put into this. Best wishes for a wonderful New Year!
As I mentioned briefly, atoms are made up of protons (+ charged), electrons (- charged), & neutrons. The electrons get a lot more attention in biochemistry because they’re negatively charged and charge makes molecules want to do things like go towards or flee from other molecules. When oppositely-charged (even partly-charged) regions of molecules come together, they can form weak, non-covalent, bonds including sometimes a type of bond called a hydrogen bond.
The number of electrons can vary and this can lead to an imbalance with the number of protons, causing a molecule to be charged (ionic). For example, a neutral molecule that gains an electron becomes negatively charged (anionic) and if it loses an electron it becomes positively charged (cationic). So, water, if it picks up an H⁺ it will become a hydronium ion (H₃O⁺) and if it gives up a proton it will become a hydroxide ion (OH⁻).
note: we often refer to H⁺ as a proton since hydrogen only has one proton, so if it loses an electron you’re just left with a proton (and neutron(s)). It can get somewhat confusing…
Electrons also get more attention because they’re the part of atoms that atoms share to form covalent bonds (the strong bonds that link together adjacent atoms in molecules). Neutrons, on the other hand, are neutral, and they’re in the atom’s central nucleus, too far away to interact with other atoms. So normally we don’t think about them much - they’re just kinda there in the background. But the number of neutrons can vary without changing the identity of the atom, and we call these different version “isotopes.” So, an atom with 1 proton is always hydrogen no matter how many electrons or neutrons it has. Though, of course, atoms can only hold a certain number of these. When atoms have more neutrons than they can handle, they’re radioactive & can decay to a less neutron-y state, letting of radiation in the process. And we can take advantage of this to radiolabel things like RNA to track it. more here: bit.ly/2VtYSG7 But not all heavier atoms are radioactive. Hydrogen can hold 2 neutrons. The “normal hydrogen” actually doesn’t have any neutrons - just a proton and an electron. We call this form protium - add an electron and you get a hydride ion (OH⁻) - remove an electron and you get a proton (H⁺), which normally hangs out with water as a hydronium ion (H₃O⁺) - add a neutron and you get deuterium - add two neutrons & you get tritium, which IS radioactive Unlike radiolabeling, where we use radioactive isotopes, deuterium isn’t radioactive - it’s stable, just “different” from normal H. So deuterated water is heavy but not “hot” (slang for radioactive) I remember in biochemistry & chemistry classes H’s would just seem to come & go out of nowhere in equations & mechanisms and it drove me crazy. But turns out hydrogen really does come and go quite readily - and frequently - if it’s attached to the “right things” - hydrogen is constantly being exchanged and we can take advantage of this to see where exchange is occurring and more significantly where it is NOT occurring Water can exist as H₂O or H⁺ and OH⁻ and that H⁺ usually grabs on to another H₂O to give you H₃O⁺ (hydronium ion). So you have an OH⁻ able to take an H & H₂O & H₃O⁺ willing to give an H. They can give and take from each other (other water molecules) or they can give and take H’s from other things. Same goes for deuterated water - it acts the same as normal water because the other molecules “ignore the neutrons” as well. So, D₂O gives you D⁺ and DO⁻. And that DO⁻ can pull off the normal H, allowing it to get swapped out. But the DO⁻ has to find that H to pull off, so it has to be solvent-accessible, and “unoccupied.” And, in order for us to be able to detect it it can’t be sooo swappable that it swaps back when we do the post-labeling stuff, which uses normal water. Proteins have a lot of hydrogens, and there are several places you’ll find them. Most of them are attached to carbons, and these H don’t like to leave without a really good reason to - those are unlikely to just swap out for a hydrogen from the water. So the exchange rates for H in C-H bonds are too small to measure. The H’s in side chain functional groups, like those in hydroxyl (-OH) and carboxyl (-COOH) groups have the opposite problem. They swap out so rapidly that when you quench the reaction in a normal water-based solution, they swap back to the light form, leaving no evidence that any change occurred in between. It’s like when you take them out of the bath, some regions dry off before you even knew they were wet. But all hope’s not lost - there’s another place that you find H’s in proteins - in the amide (-(C=O)-NH-) functional groups in the generic backbone (aka backbone hydrogens). All the letters have it except for proline, whose side chain “loops back” to bind the N so the N doesn’t have electrons to share with the H. Some letters also have exchangeable H’s in N-H’s in their side chains as well (e.g. lysine and arginine). The H’s in the N-H backbone bonds are exchangeable at a measurable rate - if they’re accessible that is. A lot of the time these H’s are tied up in hydrogen bonds with other atoms. In fact, a lot of protein structure comes from these H’s H-bonding to the carbonyl (C=O) oxygens of the backbones of other letters in other parts of the proteins. Such backbone-backbone interactions give the protein its “secondary structure” - things like alpha helices and beta strands.
What are hydrogen bonds? Basically, when atoms share electrons in covalent bonds, they don’t always do so fairly. One of them may hog the shared electrons and we say the hogger is electronegative. Oxygen and nitrogen are two of those really hoggy ones, so when they bond to hydrogen, they hog the electron pair they’re supposed to be sharing with the hydrogen, making them slightly negative (δ-) and leaving the hydrogen slightly positive (δ+). O & N also have “lone pairs” of electrons that attract such hydrogens. When an electronegative atom with a lone pair (like the O in a carbonyl, which is an O double-bonded to a carbon) is attracted to an H attached to an electronegative thing, you get a hydrogen bond (H-bond).
H-bonds are not covalent (no actual electron sharing, just attractions) so they’re not as strong as the covalent bonds that actually give the protein it’s primary structure (connect the letters in linear fashion). But they can add up to really glue the protein together. There’s structural strength in numbers - and there are lots H-bonds in proteins!
In highly structured regions of the protein, those H’s won’t be available for swapping. Though if you wait long enough those bonds can break and reform as the protein “breathes” and this offers a chance to sneak in. Speaking of time, what’s normally done is you deuterate for several different lengths of time - the longer it takes for an H to get swapped, the harder it is to find and/or the more tied-up it is. To stop it you “take away the DO⁻” by adding acid, which neutralizes the DO⁻ and lower the temperature, depriving molecules have of the energy needed to do all that swapping
If you do this quenching at different timepoints (e.g. 30s, 1 min, 2min, 5 min) you can get a sense as to how dynamic various regions are. The less protected a region is, the faster it will get heavier, and the heavier it will get. Some things you can do with HDX-MS: at the large scale - global HDX measures mass of the whole protein (no cutting it up first). you can do things like compare w/& without binding partner (ligand) -> tells you about overall binding (does it bind or not) under different conditions (e.g. at low pH, high pH, low salt, high salt, etc.) and/or w/different introduced protein mutations (e.g. if you think a residue is important for binding & you change that residue to a different letter, will you still get binding) at the finer scale - “local HDX-MS” (with cutting*) - look at changes in specific regions of the protein. This is what I’m doing.
The protein is deuterated and then cut by a pretty promiscuous protease called pepsin which is immobilized on beads (resin) in a tiny column. The peptides are captured on a pre-column before they can escape too far. This pre-column concentrates things and lets you wash off salts and other non-peptide stuff. Then you let them go them go, sending them into another tiny column which will separate them based on how much they want to interact with the resin in the column (the stationary phase) versus the liquid they’re dissolved in (the mobile phase). It’s similar in concept to the preparative-scale protein chromatography I do, but on a much smaller scale (these analytical-scale columns are only like 1mm internal diameter and 5cm long!), at much higher pressures, and with different buffers.
The columns used for mass spec are typically “reverse-phase” so they’re nonpolar, hydrophobic. The peptides glob onto them and then you elute from them gradually with a gradient of an organic solvent like acetonitrile - the more hydrophobic the stuck-on peptides are, the more nonpolar you need to make the solvent in order to convince them that the solvent is better than the resin, so the further into the gradient they’ll come off.
The column helps make the data less overlappy and so you can better tell signals apart and it also provides a level of extra information we can use to tell what signals correspond to the same peptide under different conditions (once you introduce deuteration, you’re “messing up” the ability to ID the peptides based just on their m/z so you need these extra retention time info).
The peptides come off from (elute from) the column and then get ionized by that electrospray thing and then they pass through a gas-filled ion mobility thing to further separate the ions based on their size and then they get filtered so that only peptides with specific m/z ratios can reach the detector. The detector generates an electric signal that gets drawn on a graph and stuff, and that’s about all the detail I’m gonna attempt to give as to the actual process!
*Instead of cutting with enzymes (the bottom-up approach) you can break it in the gas phase in the “top-down” approach - instead of cutting the protein in lots of places, each time it just cuts it in 1 place giving you 2 big pieces. But “each” of those pieces is cut somewhere different so you get different sized pieces and then you can compare their weights I analyzed a protein with some different binding partners and looking to see what changes. For each condition, the sample was quenched at 4 timepoints and it was run in triplicate so I have a lot of data to comb through. In “pre-runs” they ran undeuterated, but still cut up, samples of the protein in order to identify the various peptides (protein pieces) in a step called protein mapping. This allows them to check that “all” of the protein is covered and determine the retention times (how long it takes the peptide to come off the column) for each peptide in order to tell the software to keep an eye out for. Now my job is to help the software identify the actual peptide signals from the background noise and compare them.
The software identified the peptides based on their mobility in an ion phase, their retention on the column, and their falling within a possible m/z range (calculated based on the undeuterated sample’s m/z signal and the possible deuterium uptake). But then it needs help to figure out the m/z signals corresponding to those peptides, because there can be interfering signals from other things that happen to fall within that range of parameters you told the software to look for. Each “peptide” has spectral plots with this data, but you have to kinda help pick out the right sticks - those that actually correspond to the peptide.
One of the things that really confused me about peptide mass spec plots is that instead of each peptide having a single peak/stick, like you might see with a small molecule, you see a sort of asymmetric curve called a mass envelope. The reason for this is that peptides have lots of atoms and it’s not only hydrogen that has neutral isotopes. Carbon, for example, also has low levels of naturally-occurring isotopes (mainly some C13 scattered in with the “normal” C12) and any molecule contains a random mix of them (based on the isotopes natural abundances, which for carbon is ≈ 98.9% C12 and ≈ 1.1%. C13). Thus, the “same” peptide sequence-wise can have slightly different masses and you get an envelope representing the isotopic distribution. The “centroid” is the center of the data under that distribution.
Because the x-axis is m/z (mass over charge), the distance between the sticks in the envelope will depend on the charge state. If you’re looking at a +1 charged peptide, the sticks will be “1” apart, +2 they’ll be 1/2= 0.5 apart, +4 and they’ll be 1/4=0.25 apart, etc. The bigger the peptide, the more opportunities there are to get charged, so you can get high charged states which make the sticks really close together and harder to accurately pick out, but the spacing relationship can help you identify which sticks are the ones you’re interested in (thankfully for each spectrum, you only have to pick out one of the sticks and it will automatically pick the others in the envelope based on the spacing).
That relationship between the sticks won’t change when the sample gets deuterated, but the whole mass envelope will shift to the right. The more deuterium is taken up, the more it will shift, and the difference between the centroids of the undeuterated and the deuterated is the deuterium uptake. In addition to the interesting stuff (H-bonding, etc.) that uptake will depend on how big the peptide is. The bigger the peptide, the bigger the theoretical maximum uptake, which can leave smaller more exposed peptides looking wimpier. So it can be helpful to convert the y axis to relative uptake, which will tell what fraction of the theoretically-exchangeable H’s were exchanged. This evens the playing field between peptides of different sizes so you can better compare.
This sentence describes what consumed my time for a long while in grad school… “Whereas a human is much slower at making all the manipulations required to do the actual deuterium incorporation determination, a human can very rapidly provide quality control for thousands of software-determined deuterium incorporation determinations per hour, ascertaining if the software has performed well or not.”
But the rapidly wasn’t so rapid in my case, mainly due to a laggy server…
Recommended reading: Oganesyan, I.; Lento, C.; Wilson, D. J. Contemporary Hydrogen Deuterium Exchange Mass Spectrometry. Methods 2018, 144, 27-42. doi.org/10.1016/j.ymeth.2018.04.023. Masson et al. Recommendations for Performing, Interpreting and Reporting Hydrogen Deuterium Exchange Mass Spectrometry (HDX-MS) Experiments. Nature Methods 2019, 16 (7), 595-602. doi.org/10.1038/s41592-019-0459-y. Hamuro, Y. Tutorial: Chemistry of Hydrogen/Deuterium Exchange Mass Spectrometry. Journal of the American Society for Mass Spectrometry 2020, 32 (1), 133-151. doi.org/10.1021/JASMS.0C00260. My research paper: Bibel, B., Elkayam, E., Silletti, S., Komives, E. A., & Joshua-Tor, L. (2022). Target binding triggers hierarchical phosphorylation of human Argonaute-2 to promote target release. eLife, 11, e76908. doi.org/10.7554/eLife.76908 More on RNAi: Blog: bit.ly/microRNARNAi TH-cam: th-cam.com/video/7XHXF0x2uKA/w-d-xo.html more on H-bonds: bit.ly/frizzandmolecularattractions
more about all sorts of things: #365DaysOfScience All (with topics listed) 👉 bit.ly/2OllAB0 or search blog: thebumblingbiochemist.com
what you do in this channel is amazingly inspiring, and you've changed the lives of thousands of scientist-to-be from all over the world. lots of love and admire from istanbul. we can't thank you enough!
oh my gosh. How incredibly kind of you. This truly means sooooo much to me to know I'm helping people. Thank you! Happy New Year!
Hello mam, can you upload some videos on cell culture experiments
If you search my playlists you should find several. Hope they help!
Hello from Bangkok! Firstly, thanks for all your wonderful videos! I’m a ‘hard’ materials crystallographer trying to learn biochemistry. I really appreciate all the hard work that you have put into this.
Best wishes for a wonderful New Year!
So happy you found this and my other content helpful. Thank you so much for letting me know - it means a lot. Happy New Year!
As I mentioned briefly, atoms are made up of protons (+ charged), electrons (- charged), & neutrons. The electrons get a lot more attention in biochemistry because they’re negatively charged and charge makes molecules want to do things like go towards or flee from other molecules. When oppositely-charged (even partly-charged) regions of molecules come together, they can form weak, non-covalent, bonds including sometimes a type of bond called a hydrogen bond.
The number of electrons can vary and this can lead to an imbalance with the number of protons, causing a molecule to be charged (ionic). For example, a neutral molecule that gains an electron becomes negatively charged (anionic) and if it loses an electron it becomes positively charged (cationic). So, water, if it picks up an H⁺ it will become a hydronium ion (H₃O⁺) and if it gives up a proton it will become a hydroxide ion (OH⁻).
note: we often refer to H⁺ as a proton since hydrogen only has one proton, so if it loses an electron you’re just left with a proton (and neutron(s)). It can get somewhat confusing…
Electrons also get more attention because they’re the part of atoms that atoms share to form covalent bonds (the strong bonds that link together adjacent atoms in molecules).
Neutrons, on the other hand, are neutral, and they’re in the atom’s central nucleus, too far away to interact with other atoms. So normally we don’t think about them much - they’re just kinda there in the background.
But the number of neutrons can vary without changing the identity of the atom, and we call these different version “isotopes.” So, an atom with 1 proton is always hydrogen no matter how many electrons or neutrons it has. Though, of course, atoms can only hold a certain number of these. When atoms have more neutrons than they can handle, they’re radioactive & can decay to a less neutron-y state, letting of radiation in the process. And we can take advantage of this to radiolabel things like RNA to track it. more here: bit.ly/2VtYSG7
But not all heavier atoms are radioactive. Hydrogen can hold 2 neutrons. The “normal hydrogen” actually doesn’t have any neutrons - just a proton and an electron. We call this form protium
- add an electron and you get a hydride ion (OH⁻)
- remove an electron and you get a proton (H⁺), which normally hangs out with water as a hydronium ion (H₃O⁺)
- add a neutron and you get deuterium
- add two neutrons & you get tritium, which IS radioactive
Unlike radiolabeling, where we use radioactive isotopes, deuterium isn’t radioactive - it’s stable, just “different” from normal H. So deuterated water is heavy but not “hot” (slang for radioactive)
I remember in biochemistry & chemistry classes H’s would just seem to come & go out of nowhere in equations & mechanisms and it drove me crazy. But turns out hydrogen really does come and go quite readily - and frequently - if it’s attached to the “right things” - hydrogen is constantly being exchanged and we can take advantage of this to see where exchange is occurring and more significantly where it is NOT occurring
Water can exist as H₂O or H⁺ and OH⁻ and that H⁺ usually grabs on to another H₂O to give you H₃O⁺ (hydronium ion). So you have an OH⁻ able to take an H & H₂O & H₃O⁺ willing to give an H. They can give and take from each other (other water molecules) or they can give and take H’s from other things.
Same goes for deuterated water - it acts the same as normal water because the other molecules “ignore the neutrons” as well. So, D₂O gives you D⁺ and DO⁻. And that DO⁻ can pull off the normal H, allowing it to get swapped out. But the DO⁻ has to find that H to pull off, so it has to be solvent-accessible, and “unoccupied.” And, in order for us to be able to detect it it can’t be sooo swappable that it swaps back when we do the post-labeling stuff, which uses normal water.
Proteins have a lot of hydrogens, and there are several places you’ll find them. Most of them are attached to carbons, and these H don’t like to leave without a really good reason to - those are unlikely to just swap out for a hydrogen from the water. So the exchange rates for H in C-H bonds are too small to measure.
The H’s in side chain functional groups, like those in hydroxyl (-OH) and carboxyl (-COOH) groups have the opposite problem. They swap out so rapidly that when you quench the reaction in a normal water-based solution, they swap back to the light form, leaving no evidence that any change occurred in between. It’s like when you take them out of the bath, some regions dry off before you even knew they were wet.
But all hope’s not lost - there’s another place that you find H’s in proteins - in the amide (-(C=O)-NH-) functional groups in the generic backbone (aka backbone hydrogens). All the letters have it except for proline, whose side chain “loops back” to bind the N so the N doesn’t have electrons to share with the H. Some letters also have exchangeable H’s in N-H’s in their side chains as well (e.g. lysine and arginine).
The H’s in the N-H backbone bonds are exchangeable at a measurable rate - if they’re accessible that is. A lot of the time these H’s are tied up in hydrogen bonds with other atoms. In fact, a lot of protein structure comes from these H’s H-bonding to the carbonyl (C=O) oxygens of the backbones of other letters in other parts of the proteins. Such backbone-backbone interactions give the protein its “secondary structure” - things like alpha helices and beta strands.
What are hydrogen bonds? Basically, when atoms share electrons in covalent bonds, they don’t always do so fairly. One of them may hog the shared electrons and we say the hogger is electronegative. Oxygen and nitrogen are two of those really hoggy ones, so when they bond to hydrogen, they hog the electron pair they’re supposed to be sharing with the hydrogen, making them slightly negative (δ-) and leaving the hydrogen slightly positive (δ+). O & N also have “lone pairs” of electrons that attract such hydrogens. When an electronegative atom with a lone pair (like the O in a carbonyl, which is an O double-bonded to a carbon) is attracted to an H attached to an electronegative thing, you get a hydrogen bond (H-bond).
H-bonds are not covalent (no actual electron sharing, just attractions) so they’re not as strong as the covalent bonds that actually give the protein it’s primary structure (connect the letters in linear fashion). But they can add up to really glue the protein together. There’s structural strength in numbers - and there are lots H-bonds in proteins!
In highly structured regions of the protein, those H’s won’t be available for swapping. Though if you wait long enough those bonds can break and reform as the protein “breathes” and this offers a chance to sneak in.
Speaking of time, what’s normally done is you deuterate for several different lengths of time - the longer it takes for an H to get swapped, the harder it is to find and/or the more tied-up it is. To stop it you “take away the DO⁻” by adding acid, which neutralizes the DO⁻ and lower the temperature, depriving molecules have of the energy needed to do all that swapping
If you do this quenching at different timepoints (e.g. 30s, 1 min, 2min, 5 min) you can get a sense as to how dynamic various regions are. The less protected a region is, the faster it will get heavier, and the heavier it will get.
Some things you can do with HDX-MS:
at the large scale - global HDX measures mass of the whole protein (no cutting it up first). you can do things like compare w/& without binding partner (ligand) -> tells you about overall binding (does it bind or not) under different conditions (e.g. at low pH, high pH, low salt, high salt, etc.) and/or w/different introduced protein mutations (e.g. if you think a residue is important for binding & you change that residue to a different letter, will you still get binding)
at the finer scale - “local HDX-MS” (with cutting*) - look at changes in specific regions of the protein. This is what I’m doing.
The protein is deuterated and then cut by a pretty promiscuous protease called pepsin which is immobilized on beads (resin) in a tiny column. The peptides are captured on a pre-column before they can escape too far. This pre-column concentrates things and lets you wash off salts and other non-peptide stuff. Then you let them go them go, sending them into another tiny column which will separate them based on how much they want to interact with the resin in the column (the stationary phase) versus the liquid they’re dissolved in (the mobile phase). It’s similar in concept to the preparative-scale protein chromatography I do, but on a much smaller scale (these analytical-scale columns are only like 1mm internal diameter and 5cm long!), at much higher pressures, and with different buffers.
The columns used for mass spec are typically “reverse-phase” so they’re nonpolar, hydrophobic. The peptides glob onto them and then you elute from them gradually with a gradient of an organic solvent like acetonitrile - the more hydrophobic the stuck-on peptides are, the more nonpolar you need to make the solvent in order to convince them that the solvent is better than the resin, so the further into the gradient they’ll come off.
The column helps make the data less overlappy and so you can better tell signals apart and it also provides a level of extra information we can use to tell what signals correspond to the same peptide under different conditions (once you introduce deuteration, you’re “messing up” the ability to ID the peptides based just on their m/z so you need these extra retention time info).
The peptides come off from (elute from) the column and then get ionized by that electrospray thing and then they pass through a gas-filled ion mobility thing to further separate the ions based on their size and then they get filtered so that only peptides with specific m/z ratios can reach the detector. The detector generates an electric signal that gets drawn on a graph and stuff, and that’s about all the detail I’m gonna attempt to give as to the actual process!
*Instead of cutting with enzymes (the bottom-up approach) you can break it in the gas phase in the “top-down” approach - instead of cutting the protein in lots of places, each time it just cuts it in 1 place giving you 2 big pieces. But “each” of those pieces is cut somewhere different so you get different sized pieces and then you can compare their weights
I analyzed a protein with some different binding partners and looking to see what changes. For each condition, the sample was quenched at 4 timepoints and it was run in triplicate so I have a lot of data to comb through. In “pre-runs” they ran undeuterated, but still cut up, samples of the protein in order to identify the various peptides (protein pieces) in a step called protein mapping. This allows them to check that “all” of the protein is covered and determine the retention times (how long it takes the peptide to come off the column) for each peptide in order to tell the software to keep an eye out for. Now my job is to help the software identify the actual peptide signals from the background noise and compare them.
The software identified the peptides based on their mobility in an ion phase, their retention on the column, and their falling within a possible m/z range (calculated based on the undeuterated sample’s m/z signal and the possible deuterium uptake). But then it needs help to figure out the m/z signals corresponding to those peptides, because there can be interfering signals from other things that happen to fall within that range of parameters you told the software to look for. Each “peptide” has spectral plots with this data, but you have to kinda help pick out the right sticks - those that actually correspond to the peptide.
One of the things that really confused me about peptide mass spec plots is that instead of each peptide having a single peak/stick, like you might see with a small molecule, you see a sort of asymmetric curve called a mass envelope. The reason for this is that peptides have lots of atoms and it’s not only hydrogen that has neutral isotopes. Carbon, for example, also has low levels of naturally-occurring isotopes (mainly some C13 scattered in with the “normal” C12) and any molecule contains a random mix of them (based on the isotopes natural abundances, which for carbon is ≈ 98.9% C12 and ≈ 1.1%. C13). Thus, the “same” peptide sequence-wise can have slightly different masses and you get an envelope representing the isotopic distribution. The “centroid” is the center of the data under that distribution.
Because the x-axis is m/z (mass over charge), the distance between the sticks in the envelope will depend on the charge state. If you’re looking at a +1 charged peptide, the sticks will be “1” apart, +2 they’ll be 1/2= 0.5 apart, +4 and they’ll be 1/4=0.25 apart, etc. The bigger the peptide, the more opportunities there are to get charged, so you can get high charged states which make the sticks really close together and harder to accurately pick out, but the spacing relationship can help you identify which sticks are the ones you’re interested in (thankfully for each spectrum, you only have to pick out one of the sticks and it will automatically pick the others in the envelope based on the spacing).
That relationship between the sticks won’t change when the sample gets deuterated, but the whole mass envelope will shift to the right. The more deuterium is taken up, the more it will shift, and the difference between the centroids of the undeuterated and the deuterated is the deuterium uptake. In addition to the interesting stuff (H-bonding, etc.) that uptake will depend on how big the peptide is. The bigger the peptide, the bigger the theoretical maximum uptake, which can leave smaller more exposed peptides looking wimpier. So it can be helpful to convert the y axis to relative uptake, which will tell what fraction of the theoretically-exchangeable H’s were exchanged. This evens the playing field between peptides of different sizes so you can better compare.
This sentence describes what consumed my time for a long while in grad school… “Whereas a human is much slower at making all the manipulations required to do the actual deuterium incorporation determination, a human can very rapidly provide quality control for thousands of software-determined deuterium incorporation determinations per hour, ascertaining if the software has performed well or not.”
But the rapidly wasn’t so rapid in my case, mainly due to a laggy server…
Recommended reading:
Oganesyan, I.; Lento, C.; Wilson, D. J. Contemporary Hydrogen Deuterium Exchange Mass Spectrometry. Methods 2018, 144, 27-42. doi.org/10.1016/j.ymeth.2018.04.023.
Masson et al. Recommendations for Performing, Interpreting and Reporting Hydrogen Deuterium Exchange Mass Spectrometry (HDX-MS) Experiments. Nature Methods 2019, 16 (7), 595-602. doi.org/10.1038/s41592-019-0459-y.
Hamuro, Y. Tutorial: Chemistry of Hydrogen/Deuterium Exchange Mass Spectrometry. Journal of the American Society for Mass Spectrometry 2020, 32 (1), 133-151. doi.org/10.1021/JASMS.0C00260.
My research paper: Bibel, B., Elkayam, E., Silletti, S., Komives, E. A., & Joshua-Tor, L. (2022). Target binding triggers hierarchical phosphorylation of human Argonaute-2 to promote target release. eLife, 11, e76908. doi.org/10.7554/eLife.76908
More on RNAi: Blog: bit.ly/microRNARNAi TH-cam: th-cam.com/video/7XHXF0x2uKA/w-d-xo.html
more on H-bonds: bit.ly/frizzandmolecularattractions
more about all sorts of things: #365DaysOfScience All (with topics listed) 👉 bit.ly/2OllAB0 or search blog: thebumblingbiochemist.com