 There are advent of new technologies which are really trying to provide high throughput solution and much more reproducible and reliable results. One way of doing high throughput assays is to tag the proteins of interest with different type of tags or regions like fluorescent tags or radioisotopes or chemiluminescent tags and then trying to monitor the signals obtained from them. However, if you are adding a chemical moiety on your proteins of interest, it may provide some artifact results. It may alter the protein binding or the protein functions as well. So, there is lot of emphasis about how to do the protein studies in the label free manner and there is advent of new technologies which are trying to achieve the protein-protein interaction protein other biomolecular interaction studies by using label free platforms. In this slide, today we have invited a guest Dr. Shiv who is head of business operations at Nanotemper Technologies to talk to you about a latest technology MST which is micro scale thermophoresis for quantifying the binding events and studying the biomolecular interactions. Dr. Shiv will also provide you a demonstration of how to use MST technologies for biophysical assays. Towards the end, one of the colleagues an application scientist will explain the basic principles of Tycho which is used to check the protein quality and purity in a very very short time. So, in this manner you will have a good understanding about how to first test your proteins of interest quantify them to check their quantification and also their purity level and then further study how they can bind with another protein of interest or a drug molecule of interest. So, let us have this lecture with Dr. Shiv. I will quickly start my presentation without wasting much time, but before that I would like to know like are you doing any biophysical assays for the interaction studies protein protein protein small molecule. If you are working on those lines for sure this talk should be very interesting to you. So, I am sure like you are working on those lines, okay, okay, great, okay. So this is something like you know a biophysical technique that determines the binding affinities. So, as I said like you know this is the technique about a binding affinities how you determine the binding affinity between two biomolecules. It can be anything protein protein protein small molecule protein ligand like anything you can remain the binding affinity. So, but before that I would like to quickly introduce myself as myself Shiv Ramayana Lepeta you can call me Shiv currently taking care of a nano tempered business in India we are based in Bangalore and my other two colleagues Saji Millan application specialist and my other colleague Ram is also here with us. So, before going into the details of the technology I would like to give a quick introduction and how our company has formed which would be very interesting. The technology what I am talking about is thermophoresis. So, we thermophoresis have you ever heard of it before yeah we say that like you know whenever there is a heat the molecules will move from hotter region to the colder region. But it will be a director moment. So, this principle was discovered by these two gentlemen 150 years ago, but there was no any application it was only a physical principle, but there was no any application until the two CEOs of our company who filed a patent in 2008 they said that the thermophoresis can be established at a micro scale level which can be used to determine the binding affinities. So, from there the company has started and as I said like the first instrument which was launched in 2010 which is in front of you they said that using this thermophoresis principle you can determine the binding affinities. Later on different versions came into the picture which I am really not interested, but other than this we have a one more technology which measures the protein stability. Like you know you just wanted no till at what temperature your protein is stable like what you call that TM melting temperature of your protein. So, these are the two technologies what company has and as I said like these two instruments are very handy just beside me on that side and this side maybe during this talk is over we will also have a quick demonstration and we will see how easy it is to run a experiment without any optimization. So, these are the two CEOs of our company as I said like they are 31 year old who has actually filed a patent and started this company in 2010 and now we are successfully spread across the globe and as you can see a coffee machine in between them we say that our machines are very simple they are like a very robust like a coffee machine like you know it's a vending machine like you want to do an assay everything you just prepare and keep it and 15 minutes of time everything is ready on your table we will see is it as well. So, with this quick introduction of the company as I said like you know it started in a small garage in 2008 and we slowly spread across the world and we have more than 700 instruments across the world and the best part of this company is like you know you see from 2010 to 2017 we got 1700 publications in all repeated generals. So, that is how we had been connecting more to the scientists it's completely a technology driven company and it has been estimated that 15 million experiments are done using our technologies till date. So, I think this is the more than introduction of the company what I can give and now I am just coming to the technology as I said it is a thermophoresis we have established at a micro scale at a very small temperature gradient we have demonstrated this thermophoresis. This is called at a micro scale thermophoresis in a short form you call it as an MST. So, but before going into the details I just want to elaborate more on the basic principle what is thermophoresis we all know what is thermo what is electrophoresis. So, in the temperature in the like in the electric field the ions or molecules will move based on their sizes if there is any gradient in between them like an acrylamide or like any kind of a gel. So, similarly thermophoresis is the movement of the molecules in the temperature gradient thermo means temperature, phoresis movement the movement of the molecules in the temperature gradient is called as a thermophoresis. But this thermophoresis is sensitive to for three things size, charge and the hydration shell. So, as an electrophoresis you are only seeing the size right, but here this movement is dependent on three important things size, charge and the hydration shell. So, if any of these things changes during the molecular interaction the thermophoresis behavior changes. So, that is the basic principle what we have been using it maybe I will go more into the detail. Size, charge and the hydration shell. So, normally when a protein protein binds the size changes, when a protein DNA binds the charge changes. And when a protein small molecule binds the charge never changes or size never changes, but still one water molecule is displaced upon the binding. So, it all depends how it is binding and all this stuff. So, this technique has become very sensitive for all these three parameters and there will be a change in the thermophoresis behavior when there is any change in this parameters yeah. So, generally as I said like you know thermophoresis the moment you know you put a heat all the molecules move away from the heat that is a general principle. As you can see here this is just a cover slip slide which has been coated with a LMU the moment you are shining with a laser all the fluorescent molecules are moving away. So, that is the reason why you are able to see the LMU this is a Munich University. So, that is called a positive thermophoresis. The molecules moving away from the heat which is a general property is called as a positive thermophoresis. Similarly, there will be like you know the vice versa of it what you call it as a negative thermophoresis sorry. So, in the negative thermophoresis what happens is the molecule move towards the heat. So, it is like example glass beads when you shine some laser or you when you heat them all the molecules move towards the heat that is called the negative thermophoresis. So, in principle when you are doing a thermophoresis experiment you always see these two kind of a phenomenon like either they move towards the heat or they move away from the heat. But in our principle you can measure both their movement. How we measure maybe I can take you to the next slide. So, if you look into the first slide the two gentlemen who has proposed the theory of thermophoresis they state that the steady state concentration of the molecules before and heating before and heating or directly dependent upon the three parameters that is size, charge and the hydration shell. So, as I said like you know before heating whenever you are shining some heat the moment of the molecules that has gone towards the hotter region and the molecules in the colder region their ratio is always dependent upon these three parameters that is what the law of thermophoresis says. And all our experiments have been highly validated and it follows this principle. So, with this basic principle of thermophoresis we built up an instrument which has two important things one is a laser which creates a temperature for thermophoresis and here one of the binding partner is fluorescently labeled. So, just to see the fluorescence you have a fluorescent detection unit and the biggest advantage in this technique is you are not immobilizing anything as in other techniques. So, you directly determine the binding affinity in the buffer and you require only four to five microliters of the sample and the time. So, normally like you do a dilution series up to like 12 to 16 dilutions and every dilution take 40 seconds to scan. So, in 12 to 15 minutes of time you are able to determine the binding affinity without any compatibility with the buffers. There is no any crosstalk, there is no any like you know buffer interferences in your assay. There is the biggest advantage of this technique and many people ask the temperature you are creating how much is the temperature we create. It's a very steep gradient as you can see in the slide 20 to 23 degrees. So, that temperature doesn't affect any of your molecules. So, in that way what we do is you fix one of your partner and titrate the second one of a different dilutions and irradiate with the laser and you see how the biomolecules move away upon the interaction and when they are not interacting. So, that differences will help us to determine the binding affinity. So, let's see more in detail in the next slide. See if you look into the typical signal what is actually happening. As I said like you are fixing one of the partner and the second partner you are titrating it right. So, on every concentration on every dilution whatever you are taking it you are radiating the laser. So, as you can see here for the first five seconds all the molecules are uniformly distributed. This is generally what is in principle it is there. The moment you put the laser on, IR laser on what is happening is all the molecules are moving away from the heat and it states that within 30 seconds they attain the steady state. So, for 30 seconds I am putting the laser on okay. So, what is happening is all the molecules are moving away from the heat and within 30 seconds they attain the steady state as you can see here and after that I am putting the laser off. Since there is a fluorophore again there will be a like you know regaining of the fluorescence. So, again your signal will shoot up right fine. So, now I have a sample where one of the binding partner and a different concentration of the ligand is mixed and that has been taken in a small glass capillary and on every glass capillary I am taking this thermophoresis reading. First five seconds it is very uniformly distributed after five seconds you are putting the laser. Till 30 seconds we assume that it attains the steady state and after that you are putting the laser off. So, in this way for all the different dilutions whatever I am taking it I will collect the thermophoresis signal right. So, this is how typically as I said like you know in the top you can see and each and every glass capillary I have a partner along with the ligand which is like you know a different concentration. So, now every of the glass capillary is going for the thermophoresis measurement you see here. So, there is a change in the movement. Can you tell why this change is happening? Yeah upon binding either the size or a charge is changing. So, that why the movement is also changing. If the movement is not changing that means that there is no any binding at all. Then you have a signals all are overlapping one or the other as a clutter. But here as you see here the as and how there is a binding the different concentration of the ligand is coming the movement is changing right. So, this is how we track the difference between the unbound state and the bound state. The unbound is where it is not binding and in the bound state is the where the biomolecule is actually interacting. So, in this way we are actually capturing this small differences in the thermophoretic behavior to determine the binding affinity. So, once you determine all the 16 thermophoretic curves for all the different ligand concentrations you just go to the dose fit curve as you can see here. This is the baseline where typically there is no binding and here this is the saturation and the midpoint is generally as a Kd is what we call it as a binding affinity. So, where your molecules are half bound and half unbound is called your Kd. So, typically by just extracting the data from the thermophoresis curve we are fitting the dose fit curve to get a Kd value. So, this is the basic principle what it is underlying the instrument. Just by the thermophoretic behavior changes from the bound state to the unbound state you are able to determine the Kd. Since this technique is very sensitive for size, charge and the hydration shell you always get a differences in the thermophoretic behavior which will help you determine the binding affinity. The biggest advantage is you are taking a smaller volumes and you have no interferences with the buffer and there is no any limitation with the sizes. So, this is the basic principle I am sure if you have any questions you can stop me here. The rest are like how the examples like how protein-protein interacts I have some good data I want to show to you. So, this is the basic principle how this instrument works and how we determine the binding affinities. So, with this you see a wide variety of samples from ionic interactions to the liposome interactions nanodisc any kind of interactions can be studied in this instrument. So, there is no any limitation of size here. There is no any limitation of any of the properties of the molecule here. So, anything even like ionic interactions you can imagine their ions are almost in a Dalton's and even if you know liposomes or ribosomal complexes they are in a mega Dalton's. Even if there is an interaction between these things still the techniques works very nicely. As I said like you know we got 1700 publications in the last five years we have got all the supporting data with all different kind of interactions we have. So, if you are working on a specific like protein protein protein small molecule you can still discuss I can help you out like you know how we can do this I say another stuff. So, this is the main USB of this technique without any limitation of the sizes you can work with a wide variety of the samples to get the binding affinity. So, typically this technique is comparable to the SPR have you ever heard of SPR I think many people might be using the SPR as well. So, this technique is a very much comparable to the SPR and ITC isothermal calorimetry. So, in the chemistry people will be just measuring the heat change heat exchange to determine the binding affinity. So, the biggest advantage over all these techniques is as I said like you know we did a protein DNA interactions and you see in all the three techniques you are almost getting the same KD we got 29 nanomolar SPR 24 and ITC we got 30 nanomolar. So, whatever the binding affinity you determine in the MST is very similar to the ITC and SPR assays but what are the biggest advantage? So, as we are all familiar with the SPR you always have a chip you immobilize one of your protein and pump your different buffers and ligands and it involves a lot of method development and method optimization and of course the cost for every assay. And in the ITC we know like you know you have a huge pump and then you need to fill your pump and then start putting it up. So, their higher volumes of the sample is required. So, if you look into the biggest advantage of the MST is you don't require the higher volume of the sample we look with the always the lower concentration of the sample and the assay is done in 10 minutes of time with no cost. The glass capillaries are only the consumables what we use for doing the assay and it will cost you five euros per assay. So, at the end of the assay if you look on like to perform one KD measurement you require five euros. So, that is the biggest advantage the smaller volumes, time and the overall cost of the experiment. So, this has been a highly like you know worked out with all the three experiments as you can see here the cost you know it's almost like one tenth is what we generally do it for an SPR and other assay experiments. So, maybe people working with SPR still might have some questions you can still feel free to ask me. So, as I said like you know we do a labeling to one of the binding partner. So, we recently came up with a kit where you can label your his tag. So, at the six histidine position you can label your protein. So, in such a way what is the biggest advantage is like you know you need not purify your protein if your protein is in the cell lysate and it is already his tagged just simply add the dye and do the dieterations. So, in that way even in the cell lysates if you want to have a specific interaction studies this can be successfully done. So, now I will just take you to some examples of like you know how this application has been used in a different variety of interactions. One is the dimerization. So, when you are talking about the protein-protein interaction the same protein when it interacts it becomes a dimer. So, it's extremely difficult in other techniques like maybe in SPR and ITC when the same protein you are immobilizing and same protein you are running through you get a lot of artifacts. So, but here labeling one of the protein and titrating the same protein you can determine the dimerization KD as well. As you can see here you can clearly see a clean fit of KD coming up here. So, even the protein dimerizations and protein timers can be easily studied using this technology. And other most interactions people are interested is protein nucleic acids. So, I am sure you are well aware of this Chris Packer mechanism. So, here the RNA to the nucleus protein has been studied and this paper hit the nature and here in this paper they quoted that this is only biophysical technique that has been to determine the binding affinities. So, even the cutting edge technologies also MSD has become much handy tool to determine the binding affinity. So, maybe traditionally people does EMSA. So, have anybody done EMSA here? Great. So, you know like what's a pain of doing an EMSA there and you always get a qualitative you never get a quantitative information there. So, in the same studies which are done with the EMSA we try to label the DNA like a Psi 5 or Psi 6 or whatever you have the dye and then titrate your other things and you are able to determine the KD which is very much similar. So, you are with a smaller volume of the sample you are quantitatively determine the KD over this all this EMSA and radioactive labeling everything is all been like you know overlooked when you do this kind of assays here. So, other example is a protein ionic interactions these are always a very special interactions as I said like you know when a protein and ion interacts there is no size at all there is no any size making any difference the ion is a very small in size and your protein is a very big. So, that time the size doesn't change at all but still when a protein goes and binds there is a displacement of one water molecule at least outside the protein or anything. So, even with this small change we are able to determine the binding affinity this is the biggest advantage as you can see this is a one calcium binding protein which has been done and when the same experiment was done in the presence of a magnesium there was no binding as a positive and negative control this experiment has been done and as you can see here this is from the Max Black Institute and the person who has developed the ITC who is like you know known to be either father of ITC. So, always when you are working with the nano molar, picomolar interactions and a very small size molecules this addresses very nicely or any other techniques. So, and again this is like you know a protein small molecule interaction here you have a ribosomal protein complex in the red color which is like you know few mega Dalton's and a small molecule which is about like you know 120 Dalton's the cancer drug. So, when it binds the size is absolutely not like you know making any difference like if you imagine like mega Dalton protein and 120 Dalton small molecule interacts the size doesn't change at all but in principle if you look into this structural information like you know the water molecule is actually being displaced which we call it as a hydration shell because of which we are determining the thermophoresis change and the binding affinity. So, on other experiments many people who are working with the enzymes they might be more interested with the like this kind of what you call the inhibitor studies, inhibitor screenings. So, generally as you know like you know you do your binding with one specific what do you call this? The tracer molecule and once you know the KD of it and again you titrate the second one. So, wherever it displaces then you always know the binding like competitive experiments can also be easily studied. We have a well robust softwares to support all this kind of studies at the end of the day. So, this is again the same example with the high molecular weight protein complexes as I said like you know few mega Dalton proteins when in fact with the proteins still you are able to determine the binding affinities very easily. And the other challenging thing is like if anybody is working on the membrane proteins hope you know what is the pain of like making the membrane proteins more stable it requires a detergents. So, any other techniques like you know when you have a detergents you always have an buffer incompatibility. For suppose you have an STS if I have some any like a foaming agents in your things then obviously you get a lot of artifacts. So, here in this technique as I said like we are using only 3 to 4 microliters of the sample that thing can be nullified. So, even membrane proteins can successfully be studied here as you can see the paper again hit the nature where the membrane brown protein has been expressed with the GFP and that particular protein has been studied for the microscale thermophoresis. So, in this way any variety of sample it need not be a protein protein any kind of molecular interactions protein carbohydrate protein iron anything anything in between them can be studied using the microscale thermophoresis. As I said this is one example where the cell isate like when you are working with the biological fluids like maybe a serum cell isates you have a thousands of protein. So, you can even bypass by doing them a not doing any purification. If your protein of interest is always with a histag you add the dye which is very specifically binds to the histag and then titrate the other partner. So, in this way even for disease markers many people working with the proteomics are also using the MST as a complementary tool to screen the different samples for the analysis here. So, this is again just the example like how we get the data here. So, as I said like you know any one of the binding partner should be labeled it can be A or it can be B there is no a mandate that you know you should label only a protein you can label anything you can label a small molecule you can label a protein any partner can be labeled but the binding affinity doesn't change at all. So, that is the main USB of this. So, this has some examples what I want to tell for a variety of interactions how this techniques works so successfully. So, any questions or any other things you wanted me to know I shall be glad to address. So, now just as I said you are aware of the principle how it does it and you know what variety of applications can be done on this. Now, I will just quickly take to the software like you know how this instrument like how it can be operated and all these things maybe in a nutshell I just want to tell this software is an artificial intelligence driven. So, that means that the moment you set up the assay it runs on its own and once the analysis is over the software automatically assesses whether this data is good enough or not is there something like you know still you are not getting a proper results or because of any reasons it will automatically give you a suggestions. Okay, if this is not a binding fit then it will tell you please change the buffer you change the pH or you change your like you know the protein concentrations. So, like this automatically software does the assessment. So, in a way it is very much easier of course like you know as a user we always know technically what is happening with the software by looking to the data we can always assess but still software will help you to make a judgment on your analysis and you always get all the data in a single page as you see here this is a clean binding fit. So, and these are all the thermophoretic traces of all your dilutions what you have made it. So, first suppose like you know your samples are making an aggregates. So, first suppose like upon a particular concentration when it is interacting with the ligand it becomes an aggregates. So, then aggregates means then the sample will be non-homogeneous. The thermophoresis behavior also will be very like random the curve likes you know looks very very random I will just show how it looks like. See sorry see whenever there is an aggregation see you see here that the curve is completely like you know a wavy. So, like this qualitatively we can always judge the aggregation of your sample also. So, in other experiments when a sample is aggregating it is already in contact with the instrument it spoils the next run it spoils the next things. So, the biggest advantage here is you are using a different independent capillaries for every dilution. So, if one capillary is having an aggregate you can just discard them and you know at what concentration your protein is actually aggregating. So, in this way qualitatively you can judge the aggregation in your samples by using this kind of a technique. So, maybe this all these things I do not want to bore you much because as I said like it is more for a users who are actually doing it. In principle the software is completely a artificial intelligence driven it gives you a solutions at the end of the day like how you need to move on. So, this is the just I just want to summarize quickly we are already associated with the top premium institutes across the globe and I am very glad that in India we are associated with the IIT Hyderabad, RCB, Novozymes and many other customers who are using this technology. And as I said like you know in all kind of generals you can find the MST papers like in nature we have more than 15 papers, science all the best generals have acknowledged this is a biophysical tool that actually be a very useful tool in measuring the binding affinities. So, here are some customers who had been using a lot of SPR, ITC experiments and now made a shift to the techniques and found how easy it is. Like if you are doing any screening like you know you are doing a drug screening you have a small molecules libraries hundreds of thousands of molecules. So, I am sure like in other techniques will be a nightmare. So, here since you are using 10 to 12 minutes of time you can always screen them very quickly. And these are our some customers who has used it and gave a very good feedback and we have more than 10 publications from Indian customers in the last 24 months. The final slide is always what I say as I said in the beginning it is a company of scientists. So, along with the it's not just telling about the instrument it is all about giving a good support. So, you are always bundled with a good scientist to support you and develop the assays on your own. So, that's all I want to tell since we already are going to give a quick demo maybe like you know we can just quickly share some questions you have or if you are working on any other things what are the challenges you are facing I shall be very glad to help you out. What kind of samples you do on SPR basically at the moment? ITC, okay. So, for sure like you know as I said like now we have a small a positive control kit. It is a DNA aptamer and a small molecule ATP basically ADP and the DNA aptamer. So, anybody is interested they can just come in front and then just do it because I said like doesn't require any much training much thing to understand. As I said the basic principle is thermophoresis and how the every dilution there's a difference is what we need to understand and rest everything is done by the software automatically. See that's what I'm saying in principle anything is possible alkaloid can you tell me an example? Curcumin. So, very recently I think as I said like 15 minutes of the time is the measurement time. So, maybe people can come friend and this want or you can sit there and see how are you wish. So, we have a control where we are just going to load into the capillaries and do the like an MSTSA and we'll go to this one the stability one. So, there also the things are ready it takes only three minutes to measure the stability of your protein that is the biggest advantage. So, maybe she will give a small talk on few slides what are the basic principle and what are the applications. So, anybody working with the protein can use this instrument for quickly checking like you know the functionality of your protein whether your protein is really functional or not okay. So, these are the glass capillaries what we use for that say you can see these are the simple glass capillaries which can accommodate maximum of five to six microliters you just dip into the solution. So, normally you take a PCR steps and then like you know fix your partner A and titrate the ligand like generally like 12 to 16 dilutions is what we prepare and all the 12 to 16 dilutions each dilution you take in the glass capillary. So, and this glass capillary will go for the measurement. So, and here you have already like you know we already mixed the DNA aptamer and ATP here. So, what I'm just going to do is now you know and just dipping the glass capillary and then loading it. So, it's all like once you load it within 10 minutes of time you can do it. So, normally you know proteins are very like you know unstable at a room temperature. So, the DNA has been used the DNA is very stable here. So, for the for the generally for demos I prefer to do with the DNA because it's more stable you can carry anywhere what you wanted yeah. So, this is a tray which holds your glass capillaries. So, it's like labeled one to 16. So, like whatever you are taking it you just try to put into this and then do it. And we have a small magnetic strip to arrest your glass capillaries. It's like you know when you are loading it they might not fell down. So, it's just a magnetic strip to arrest the glass capillaries when you are loading it. That's all. So, maybe we'll take two more minutes to load and after this we can just start. Yeah, yeah as somebody want to load they can come and load you can load it no problem. Or a protein purification by columns or anything like that. So, how you maintain the quality how you check the quality that is nothing but the stability or the functionality. So, we have recently launched this instrument only not even a one month old is this. So, we can quickly check the protein quality. How, what is the basic principle of this and what kind of applications she'll try to summarize and give it in just 15 minutes of time. And after that that instrument also we can give a demo. It takes in principle only three minutes to run one like run six samples. We got some controls we can quickly run that also. Sachi please go ahead. So, have you ever asked to yourself that whatever protein you are purifying whether it's pure or not. I mean whether that can be used for your future essay experiments or not. So, that's a very big question that always comes in the mind of any researcher. So, before going for any essay what you do is like you run a column you do a SDS page or you do spectrophotometric analysis. So, these are like very standard goal methods you're going for a spectrophotometry or electrophoresis or a column chromatography. But still you're not aware whether your protein is a functional protein or not. So, we have this newly launched product which is known as Tycho. And in one go it takes six samples and you can actually check the quality of the protein. So, it will come to know whether the protein what you have you know purified is functional or not. So, what's the basic principle of this machine is that it can work with any kind of protein whether it's antibody or whether it's a normal protein, membrane protein, receptor protein, any kind of protein and give a three minute measurement. So, within three minutes you're actually checking your quality and the purity of the protein. So, it's a label free measurement. Like here one of the partner is labeled you are not labeling here. It's the intrinsic protein fluorescence that we are checking which will give you a label free what you call inflection temperature which is very equivalent to the melting temperature. And you're similarly using capillaries and the amount of the volume that you're using is less than 10 micro liter. And in one go almost six samples you can analyze. Then the run rate here is little fast because you are analyzing you know in three minutes. So, the run rate here is 30 degree per minute and as of our all the instruments are maintenance free. There is no cleaning. You're just putting the capillaries in the instrument taking off once the run is complete. So, what are the basic benefits here is that it's a faster measurement technique and it checks the purification and the I mean it checks at the purification and the characterization levels. So, if you have like batch to batch samples or like one day old samples or 10 day old samples or like very old samples and you're free storing the samples all that samples when you take and check one by one you can check the actual purity whether it has denatured or not like that you can always check batch wise. Then you can analyze the samples in a wide range that is from five microgram per ml to 250 mg per ml. And as you can see it's a very small instrument you can hand carry the instrument here and there. So, it's a very easy to you know work with. So, when you see the characterization workflow where all the Tycho can be used. So, when you go for the initial purification after that when you get the protein you can check the purity at this level. After chromatography I mean ion exchange chromatography or column purification you can again check the purity of your protein. Once the protein is purified then you can again you have to store it at different conditions. So, at which buffer at which pH you want to store it you can always check that and then store it in that conditions. Once you have stored the proteins suppose you have kept it for 10 days or 20 days after that you want to again check the purity because for SA you need to have a pure protein. If it's not pure then definitely suppose you are going for SPR. One chip you can't waste if your protein is not pure. So, before the SA development you always have to check the protein how the protein is. If it's totally denatured and you are doing an SA definitely you are you losing your time as well as the chip what you are using for a SPR. So, all these are the steps where you can actually check your protein quality. Now, I have few applications where it will tell you where all tyco can be used. Now, in this when you see what is the basic principle that is applied here is that it is checking the intrinsic tryptophan fluorescence. So, protein from a folded to a unfolded state it will give you a particular T i value. So, the temperature the results that a tyco is going to give is the initial ratio. The initial ratio is the ratio where you know it will give you the purity of the protein I mean whether it's denatured or not. And the delta ratio is the ratio which is from the folded to the unfolded region. Like that when the protein folds when the protein unfolds you will get a T i value as well as the sample brightness. The sample brightness here will give you a relative concentration of your protein molecule. It will not give you actual concentration but a relative concentration that T i this much amount of protein is present. And then protein profile similarity is that if you have a reference sample of your protein that can be taken as a standard protein sample and then when you are going for batch to batch or like if you are doing a freeze thawing. At that time you can always take that as a reference sample and cross check the other batch to bias or you can freeze thawed once and then check how much it has denatured or how much it has changed. Now, here is one example where you have compared it with batch to batch you know screenings. So, here you can see the protein which is a pure protein and a protein that has been kept in minus 80 it is almost showing a similar profile. But when it is used after one week but kept at 4 degrees centigrade the profile has changed. So, you can see the shift has gone up that means the protein has slightly unfolded. So, that means you can definitely go for a say but still you can check the quality of the protein how it has varied. Now, here you can check the buffer screening as well as the pH stability measurements. So, you know like basically for SPR you need low pH. So, if you have a standardized form of how to do it you can check at pH 4 it is totally denatured and then at pH 5 which is like a basic pH for SPR measurements the curve is almost good. I mean when you compare it with different buffers you can always check which pH you can use which buffer you can use. Now, here you can also check the thermal shift assay. I mean a protein is there and if you want to check whether the protein can bind with a particular ligand or not. So, when you singly when you run the protein without the ligand you will see a particular profile and then with the in the presence of a ligand the shift will change. So, this shift can either be a positive shift or a negative shift. If it is a positive shift that means it is a true binder. If it is a negative shift that means it is not a true binder. We can actually go for a yes or no that it whether the shift is happening or not. I mean whether the interaction is possible or not. With this you can go for further assays like if you are going for SPR or if you are going for ITC you can say that oh this protein with this ligand it is good because there is a thermal shift because the Ti's are shifting here. I have few examples which will show you the data. Now this is a example in which the monoclonal antibodies was checked at different oxidation levels. So, you can see at the native state it has given a different profile and then 3 hour oxidation with hydrogen peroxide it has given a different profile and 18 hour oxidation it is giving a different profile. So, you can see there is a Ti shift in all the 3 samples and when you had gone for the MSD assay the same thing you have you can see. The shift is I mean the KD value is changing with respect to oxidation. If you want to go for a biosensor assay like the SPR. So, here you have to check the pH values. Definitely which pH is best for the SPR. So, you can always green different pH ranges and check which buffer is best. So, like here kinase is taken as a standard and at different pH it is tested. So, you can see at pH 7 and pH 4 and pH 4.5 it is just giving a planar line. That means it is already denatured. But the other in pH 7 and pH 5.5 it is giving a curve which is showing the stability of the protein. Now, this is a thermal shifter assay that we have done where we have taken a kinase and then the kinase is treated with ligand molecule. You can see the shift in the assay. You can see protein alone was giving a shift at 50, 55 and once it is added with the ligand the shift has gone to 62. So, that is how you can go check the thermal assays. Now, this is once the chromatography is run. I mean you get lot of fragments. So, you check all the fragments and see which is your best, which is the best fragment where your protein is there. So, this will give you a protein profile that this fragment or the first fragment or the second fragment is having the best protein. But if you have a standard reference along with it you can always validate that the protein is actually the pure protein. Now, here is another example that we have conducted where you have taken fresh samples, free-store cycle samples and fully denatured and partially denatured samples. So, you can see how the trend has changed when you are checking the unfolding profiles. So, the fully denatured sample is giving you a very planar profile where as the partially denatured is giving a different profile compared to the free store and the fresh samples. So, the same is analyzed with the MST. So, you can see the fully denatured sample is you know there is no KD value to it. But the other samples you can get a KD value, but the KD value will vary compared to the fresh sample. So, in all if you see it is a very quick tool to determine the purity as well as the you know stability and functionality of the protein. And when you want to characterize some kinases some receptors or some membrane proteins or transmembrane proteins like that. You can always check the purity of those proteins and like it will give you a yes or no signal when you want to do a thermal stability assays or you want to go for a SPR assay or a cryo EMSA or NMR studies. So, before that you want to check whether your protein is stable or not whether your protein is pure or not. So, yes or no will always this machine will always help you with the yes or no decisions. So, I can give you a small demo with one sample that is the BSA. There is one mg per ml I have taken. So, I can just give you it is a three minute run that will be there. And we can check the profile how it goes. Question. Do you have any example of this non-term reference proteins whether they change? Yes, I have. There is still many sample proteins the reference proteins that are used for example, as the reference proteins we do not have, but in yeah. But like if we do for you yes. So, like if we have a reference protein we can always you know if you give us a fresh sample if we have a standard protein we can always correlate with that. In principle no. Because in few of the demos we have seen this kind of shift where there is a positive shift as well as a negative shift. So, positive shift. Yeah. So, the positive shift will always say that it is a binder but the negative shift is destabilizing your protein and you can see the profile when it goes above 0.9 or something whether it is fully denatured or not. When we store a protein it will be denatured as a base plasma it will be some important it will be unfolded to not be a change. Basically the proteins. It depends upon your specific protein body. It is the same process. Exactly that is the main purpose. As long as like 10 percent is fine if your protein is unfolded within 30-40 percent then obviously like when you are working with the red protein which doesn't make any sense because suppose you are immobilizing your protein and you are changing a lot many profiles and you are trying to beat everything and if it is not working the problem is your protein is really not in a good state actually. A quick checkpoint can always be taken care of and when you are doing a purification like in all the fractions you need not go to the STSB and then check every time. So, quickly you can always check it It is basically like a quick checkpoint to know what is the quality of the protein what you are working with and till now you don't have any direct method to like you know measure this kind of quality through the quantitation by spectrometry you grind forward other things and you do a quality purification you run the STSB but there is no any checkpoint so if you know that the protein you work with is really stable and functional or not so when you work with a biosys unless your protein is not functional there is no point in like you know setting up the assay and spending a lot of time and since it is like you know a label free you don't require anything like you know just dip it and then do the measurement so that would be like a way to check the protein quality and many a times when you take the proteins out I mean you keep it in eyes but still taking in out in out so you know how whether the protein is stable or not so this is like a quick tour the percentage is unfolding like as you said after 10 days after 1 month what is the percentage that is unfolding see as long as 20-30% it's fine when it goes above 50% then sure like you know everything changes there is not even a true protein what you are actually working on so on the background we never know what is actually happening when you are taking from minus 20 or minus 80 and thawing it your protein might be spoiled but you are not sure but still you are going for the biosys it doesn't work then you think that when there is something else wrong it doesn't have to be like now unfolded while doing the fine work so at that temperature protein will not get to you see our basic principle here is see when you see a protein the nitrofans are actually buried inside right only the nitrofans are buried inside the protein and then now you are heating it up the nitrofans gets exposed once your nitrofans are exposed the fluorescence is changing so when completely all the nitrofans are exposed like you know the activity or like you know the company unfolding is happening so that is the basic principle what we are taking here and we are wrapping 30 minutes per minute from 30 to 90 we are reaching 3 minutes period because just you are heating the sample and once all the nitrofans are exposing your sample then you are getting a like you know what you call the transition like T I inflection point so that is the basic principle but we have other version of the instrument so you can get it 2 pms the welding temperature like now you can always control the gravity and can get the pms values also it is very similar to DLC but this one is something like for any bio or say any mass because like you know most of the intact proteins when you are heating the LC-MS like if the protein is already dead then for sure you do not get it you just go and throw your nano LC or something like that so here we quickly say A my protein is fine so you can go into the LC-MS so like this at a quick check point at a different experiments like NMR or COVEM any kind of things you can always use as a quick tool to know the protein quality that is the main purpose of this and first effect 2 and also like 3 minutes like now hardly matters like now 6 samples you are getting the data in 3 minutes just you are heating it and then pulling it back that is the end question do you want to do a DSC or a CV or this kind of thing it is called mass spectrometry fine so that is all we quickly 3 minutes at a sex you load 1 sample quickly then comes here the time you will be on there so you just load 1 sample it is fine just we are showing the previous data to get an understanding because we had only 1 BSA so like you know you can clearly see there is a change in the pms you know TI points like whenever there is an interaction generally you know it will always stabilize the complex so there will be a shift in the pms so typically you always get the curves with a clear pms and if you just look at the results you can always see you always get the temperatures at which they got unfolded and it is clearly showing a shift of somewhere 4 degrees so like this you can always see the differences in the pms so that is the biggest advantage in the stability of your protein for always I know like many times you prefer the same protein over some months or something like that so you can always compare like you know what is the profile similarity what is the percentage loss I am doing here so this kind of all things you can always integrate in your workflow to measure the protein stability so you always get you know the ratio which will tell you the percentage unfolding the sample brightness it is not exactly a one way new parameter it is kind of a relatively if I am taking one sample as a standard I can guess that one what is the brightness what do you have it like it will always give a really new quantitation figure also so it is very easy like you know as I said like you have to point where you can always assess the quality before going to any bio assay it can be SPR it can be like any ITC anything like that so you can always quickly know the quality of your protein whether it is really good or not and if it is good then compare it with your references how it actually works so that is the biggest advantage what you have to do because one sample is not make sense to show you all the data so we are truly the previous start so this is how typically you are getting it here the software automatically calculates and then show you this is the theme of it like this if you say I am absorbing two to three proteins in same sample then it all depends like you know when it is actually unfolding in some the tryptophan might be exposed some tryptophan are like already very inside so in principle you get always a very nonspecific data now it is a biophysical always work with a few proteins is recommended but yes many of our customers like with two proteins they have tried it so always they know like you know one protein Dm another one whatever it was getting from that another protein Dm it is always like RND kind of thing we can see that and you can always see the multiple Dms you can have the same protein have multiple domains which are like exposed at different temperatures like what will be maps if you see you get the Fab region and the FC region at different types so like this even if you have same proteins having multiple teams you can still get the data so this is what we can do so this is how you get the typical results so any questions or something I am sure like you know your proteomics means everybody will be doing all this kind of things so we should be very glad if you are interested we can evaluate and we can still do more insights in the technology the advent of new technologies both label based and label free have now started offering us new insights for lot of new functions for the proteins of interest for which there is no function defined earlier in this slide there is advent of new technologies which aims to do these experiments without taking the protein of interest and also provide the binding and the kinetics information in the slide there are new developments new technologies are coming forward and today we try to provide you interaction with one of the leading companies and their application scientist who talk to you about MST technologies and also the quality control assays we are done using tyco technology I hope these provide you some insight about how to do your experiments for the proteins to further characterize using latest label free biosensors thank you