About this Author
Derek Lowe, an Arkansan by birth, got his BA from Hendrix College and his PhD in organic chemistry from Duke before spending time in Germany on a Humboldt Fellowship on his post-doc. He's worked for several major pharmaceutical companies since 1989 on drug discovery projects against schizophrenia, Alzheimer's, diabetes, osteoporosis and other diseases.
To contact Derek email him directly: derekb.lowe@gmail.com
Twitter: Dereklowe
|
Category Archives
March 6, 2009
Posted by Derek
There are a huge number of techniques in the protein world that relay on tying down some binding partner onto some kind of solid support. When you’re talking about immobilizing proteins, that’s one thing – they’re large beasts, and presumably there’s some tether that can be bonded to them to string off to a solid bead or chip. It’s certainly not always easy, but generally can be done, often after some experimentation with the length of the linker, its composition, and the chemistry used to attach it.
But there are also plenty of ideas out there that call for doing the same sort of thing to small molecules. The first thing that comes to mind is affinity chromatography – take some small molecule that you know binds to a given protein or class of proteins well, attach it to some solid resin or the like, and then pour a bunch of mixed proteins over it. In theory, the binding partner will stick to its ligand as it finds it, everything else will wash off, and now you’ve got pure protein (or a pure group of related proteins) isolated and ready to be analyzed. Well, maybe after you find a way to get them off the solid support as well.
That illustrates one experimental consideration with these ideas. You want the association between the binding partners to be strong enough to be useful, but (in many cases) not so incredibly strong that it can never be broken up again. There are a lot of biomolecule purification methods that rely on just these sorts of interactions, but those often use some well-worked-out binding pair that you introduce into the proteins artificially. Doing it on native proteins, with small molecules that you just dreamed up, is quite another thing.
But that would be very useful indeed, if you could get it work reliably. There are techniques available like surface plasmon resonance, which can tell with great sensitivity if something is sticking close to a solid surface. At least one whole company (Graffinity) has been trying to make a living by (among other things) attaching screening libraries of small molecules to SPR chips, and flowing proteins of interest over them to look for structural lead ideas.
And Stuart Schreiber and his collaborators at the Broad Institute have been working on the immobilized-small-molecule idea as well, trying different methods of attaching compound libraries to various solid supports. They’re looking for molecules that disrupt some very tough (but very interesting) biological processes, and have reported some successes in protein-protein interactions, a notoriously tempting (and notoriously hard) area for small-molecule drug discovery.
The big problem that people tend to have with all these ideas – and I’m one of those people, in the end – is that it’s hard to see how you can rope small molecules to a solid support without changing their character. After all, we don’t have anything smaller than atoms to make the ropes out of. It’s one thing to do this to a protein – that’ll look like a tangle of yarn with a small length of it stretching out to the side. But on the small molecule scale, it’s a bit like putting a hamster on a collar and leash designed for a Doberman. Mr. Hamster is not going to be able to enjoy his former freedom of movement, and a blindfolded person might, on picking him up, have difficulty recognizing his essential hamsterhood.
There's also the problem of how you attach that leash and collar, even if you decide that you can put up with it once it's on. Making an array of peptides on a solid support is all well and good - peptides have convenient handles at both ends, and there are a lot of well-worked-out reactions to attach things to them. But small molecules come in all sorts of shapes, sizes, and combinations of functional groups (at least, they'd better if you're hoping to see some screening hits with them). Trying to attach such a heterogeneous lot of stuff through a defined chemical ligation is challenging, and I think that the challenge is too often met by making the compound set less diverse. And after seeing how much my molecules can be affected by adding just one methyl group in the right (or wrong) place, I’m not so sure that I understand the best way to attach them to beads.
So I’m going to keep reading the tethered-small-molecule-library literature, and keep an eye on its progress. But I worry that I’m just reading about the successes, and not hearing as much about the dead ends. (That’s how the rest of the literature tends to work, anyway). For those who want to catch up with this area, here's a Royal Society review from Angela Koehler and co-workers at the Broad that'll get you up to speed. It's a high-risk, high-reward research area, for sure, so I'll always have some sympathy for it.
Comments (12)
+ TrackBacks (0) | Category: Analytical Chemistry | Drug Assays | General Scientific News
January 22, 2009
Posted by Derek
Now here’s a news item that I’m pretty sure you haven’t heard about unless you work in or near a laboratory. We’re in the middle of an extreme shortage of acetonitrile, a common solvent. This has been going on since back in the fall, but instead of gradually getting better, it’s been gradually getting worse: major suppliers are sending out letters like this one (PDF).
What’s the stuff good for? Well, it’s used on a manufacturing scale in some processes, so they’re in trouble for sure. Acetonitrile is a good solvent, since it’s fairly powerful at dissolving things but still reasonable low-boiling. (That’s the nitrile functional group for you; there’s nothing else quite like it). It’s no DMSO, but then again, DMSO’s boiling point is three times a lot higher, and compared to acetonitrile it pours like pancake syrup. Nobody does industrial-scale chemistry in DMSO if they can possibly help it.
Those properties mean that acetonitrile/water mixtures are ubiquitous in analytical and prep-sized chromatography systems. This is surely its most widespread use, and is causing the most widespread consternation as the shortage becomes more acute. Many people are switching to methanol/water, which usually works, but can be a bit jumpier. But that’s not always an option. Labs working under regulatory-agency controls (GLP / GMP) have a very hard time changing analytical methods without triggering a blizzard of paperwork and major delays. In many companies, it’s those people who are first in line for what acetonitrile may turn up.
So why are we going dry on the stuff? There seem to be several reasons, one of which, interestingly, is the summer Olympics. The industrial production that the Chinese government shut down to improve Beijing’s air quality seems to have included a disproportionate amount of the country’s acetonitrile production (for example). A US facility on the Gulf Coast was shut down during Hurricane Ike as well. But on top of these acute reasons, there's a secular one: yep, the global economic slowdown. A lot of acetonitrile comes as a byproduct of acrylonitrile production, which is used in a lot of industrial resins and plastics. Those go into making car parts, electronic housings, all sorts of things that are piling up in inventory and thus not being turned out at the rates of a year ago.
So taken together, there’s not much acetonitrile to be had out there. We’ve seen some glitches like this in the past, naturally, since chemical production can depend on a limited number of plants and on raw material prices. When I was an undergraduate, I remember professors complaining aboiut the price of silver reagents during the attempted Hunt brothers corner of that market, for example. But this one will definitely be near the top of the list, and it could be months before the Great Acetonitrile Drought lifts. If you've been saving some in your basement, it’s time to break it out.
Comments (101)
+ TrackBacks (0) | Category: Analytical Chemistry | Chemical News
October 9, 2008
Posted by Derek
I’ve spoken before about the acetylene-azide “click” reaction popularized by Barry Sharpless and his co-workers out at Scripps. This has been taken up by the chemical biology field in a big way, and all sorts of ingenious applications are starting to emerge. The tight, specific ligation reaction that forms the triazole lets you modify biomolecules with minimal disruption (by hanging an azide or acetylene from them, both rather small groups), and tag them later on in a very controlled way.
Adrian Salic and co-worker Cindy Yao have just reported an impressive example. They’ve been looking at ethynyluracil (EU), the acetylene-modified form of the ubiquitous nucleotide found in RNA. If you feed this to living organisms, they take it up just as if it were uracil, and incorporate it into their RNA. (It’s uracil-like enough to not be taken up into DNA, as they’ve shown by control experiments). Exposing cells or tissue samples later on to a fluorescent-tagged azide (and the copper catalyst needed for quick triazole formation) lets you light up all the RNA in sight. You can choose the timing, the tissue, and your other parameters as you wish.
For example, Salic and Yao have exposed cultured cells to EU for varying lengths of time, and watched the time course of transcription. Even ten minutes of EU exposure is enough to see the nuclei start to light up, and a half hour clearly shows plenty of incoporation into RNA, with the cytoplasm starting to show as well. (The signal increases strongly over the first three hours or so, and then more slowly).
Isolating the RNA and looking at it with LC/MS lets you calibrate your fluorescence assays, and also check to see just how much EU is getting taken up. Overall, after a 24-hour exposure to the acetylene uracil, it looks like about one out of every 35 uracils in the total RNA content has been replaced with the label. There’s a bit less in the RNA species produced by the RNAPol1 enzyme as compared to the others, interestingly.
There are some other tricks you can run with this system. If you expose the cells for 3 hours, then wash the EU out of the medium and let them continue growing under normal conditions, you can watch the labeled RNA disappear as it turns over. As it turns out, most of it drops out of the nucleus during the first hour, while the cytoplasmic RNA seems to have a longer lifetime. If you expose the cells to EU for 24 hours, though, the nuclear fluorescence is still visible – barely – after 24 hours of washout, but the cytoplasmic RNA fluorescence never really goes away at all. There seems to be some stable RNA species out there – what exactly that is, we don’t know yet.
Finally, the authors tried this out on whole animals. Injecting a mouse with EU and harvesting organs five hours later gave some very interesting results. It worked wonderfully - whole tissue slices could be examined, as well as individual cells. Every organ they checked showed nuclear staining, at the very least. Some of the really transcriptionally active populations (hepatocytes, kidney tubules, and the crypt cells in the small intestine) were lit up very brightly indeed. Oddly, the most intense staining was in the spleen. What appear to be lymphocytes glowed powerfully, but other areas next to them were almost completely dark. The reason for this is unknown, and that’s very good news indeed.
That’s because when you come up with a new technique, you want it to tell you things that you didn’t know before. If it just does a better or more convenient job of telling you what you could have found out, that’s still OK, but it’s definitely second best. (And, naturally, if it just tells you what you already knew with the same amount of work, you’ve wasted your time). Clearly, this click-RNA method is telling us a lot of things that we don’t understand yet, and the variety of experiments that can be done with it has barely been sampled.
Closely related to this work is what’s going on in Carolyn Bertozzi’s lab in Berkeley. She’s gone a step further, getting rid of the copper catalyst for the triazole-forming reaction by ingeniously making strained, reactive acetylenes. They’ll spontaneously react if they see a nearby azide, but they’re still inert enough to be compatible with biomolecules. In a recent Science paper, her group reports feeded azide-substituted galactosamine to developing zebrafish. That amino sugar is well known to be used in the synthesis of glycoproteins, and the zebrafish embryos seemed to have no problem accepting the azide variant as a building block.
And they were able to run these same sorts of experiments – exposing the embryos to different concentrations of azido sugar, for different times, with different washout periods before labeling all gave a wealth of information about the development of mucin-type glycans. Using differently labled fluorescent acetylene reagents, they could stain different populations of glycan, and watch time courses and developmental trafficking – that’s the source of the spectacular images shown.
Losing the copper step is convenient, and also opens up possibilities for doing these reactions inside living cells (which is definitely something that Bertozzi’s lab is working on). The number of experiments you can imagine is staggering – here, I’ll do one off the top of my head to give you the idea. Azide-containing amino acids can be incorporated at specific places in bacterial proteins – here’s one where they replaced a phenylalanine in urate oxidase with para-azidophenylalanine. Can that be done in larger, more tractable cells? If so, why not try that on some proteins of interest – there are thousands of possibilities – then micro-inject one of the Bertozzi acetylene fluorescence reagents? Watching that diffuse through the cell, lighting things up as it found azide to react with would surely be of interest – wouldn’t it?
I’m writing about this the day after the green fluorescent protein Nobel for a reason, of course. This is a similar approach, but taken down to the size of individual molecules – you can’t label uracil with GFP and expect it to be taken up into RNA, that’s for sure. Advances in labeling and detection are one of the main things driving biology these days, and this will just accelerate things. (It’s also killing off a lot of traditional radioactive isotope labeling work, too, not that anyone’s going to miss it). For the foreseeable future, we’re going to be bombarded with more information than we know what to do with. It’ll be great – enjoy it!
Comments (7)
+ TrackBacks (0) | Category: Analytical Chemistry | Biological News
September 23, 2008
Posted by Derek
Over the years, when some puzzling feature of a drug candidate’s binding to a target came up, I’ve often said “Well, we’re not going to know what’s happening until some lunatic builds a femtosecond X-ray laser”. Various lunatics are now pitching in to build some. I’m going to have to revise my lines.
The reason I’d say such a mouthful is that we already, of course, get a lot of structural information from X-ray beams. Shining them through crystals of various substances can, after a good deal of number-crunching in the background, give you a three-dimensional picture of how the unit molecules have packed together. Proteins can be crystallized, too, although it can be something of a black art, and they can be either crystallized with or soaked with our small molecules, giving us a picture of how they’re actually binding.
There are, as mentioned earlier around here, plenty of ways for this process to go wrong. For starters, a lot of things – many of them especially interesting – just don’t crystallize. And the crystals themselves may or may not be showing you a structure that’s relevant to the question you’re trying to answer – that’s particularly true in the case of those ligand-bound protein structures. And the whole process is only good for static pictures of things that aren’t moving around. It used to take many days to collect enough data for a good crystal structure. That moved down to hours as X-ray sources got brighter and detectors got better, and now X-ray synchrotrons will blast away at your crystals and give you enough reflections inside of twenty minutes. And that’s great, but molecules move around a trillion times faster than that, so we’re necessarily seeing an average of where they hang out the most.
Enter the femtosecond X-ray laser. A laser will put out the cleanest X-ray beam that anyone’s ever seen, a completely coherent one at an exact (and short) wavelength which should give wonderful reflection data. The only ways we know how to do that are on large scale, too, so it’s going to be a relatively bright source as well. The data should come so quickly, in fact, that several things which are now impossible are within reach: X-ray structures of single molecules, for one. X-rays of things that aren’t in a crystalline state at all, for another. And femtosecond-scale sequential X-ray structures – in effect, well-resolved high-speed movies of molecular motions.
Now that will be something to see. Getting all that to work is going to be quite a job, not least because X-ray bursts of this sort will probably destroy the sample that they're analyzing. But there are two free-electron X-ray lasers under construction – one set to complete next year at Stanford’s SLAC facility and a larger one that will be built in Hamburg. “Large” is the word here. The smaller SLAC instrument is already two kilometerslong. According to an article in Nature, though, a Japanese group have proposed some ways to make future instruments smaller and more efficient – all the way down, to, um, the size of a couple of football fields. But there’s another completely different technology coming along (laser-plasma wakefield instruments) that could produce far shorter X-rays in one hundredth the space, which is more like it.
I don’t think we’re going to see a benchtop-sized X-ray laser any time soon, especially since these things are going to need to be large just to get up to the brightness that will be needed. But I’m very interested to see what even the first generation machine at Stanford will be able to do. There are a lot of mysteries in the way that molecules move and interact, and we may finally be about to get a look at some of them.
Comments (12)
+ TrackBacks (0) | Category: Analytical Chemistry
September 4, 2008
Posted by Derek
X-ray crystallography is wonderful stuff – I think you’ll get chemists to generally agree on that. There’s no other technique that can provide such certainty about the structure of a compound – and for medicinal chemists, it has the invaluable ability to show you a snapshot of your drug candidate bound to its protein target. Of course, not all proteins can be crystallized, and not all of them can be crystallized with drug ligands in them. But an X-ray structure is usually considered the last word, when you can get one – and thanks to automation, computing power, and to brighter X-ray sources, we get more of them than ever.
But there are a surprising number of ways that X-ray data can mislead you. For an excellent treatment of these, complete with plenty of references to the recent literature, see an excellent paper coming out in Drug Discovery Today from researchers at Astra-Zeneca (Andy Davis and Stephen St.-Gallay) and Uppsala University (Gerard Kleywegt). These folks all know their computational and structural biology, and they’re willing to tell you how much they don’t know, either.
For starters, a small (but significant) number of protein structures derived from X-ray data are just plain wrong. Medicinal chemists should always look first at the resolution of an X-ray structure, since the tighter the data, the better the chance there is of things being as they seem. The authors make the important point that there’s some subjective judgment involved on the part of a crystallographer interpreting raw electron-density maps, and the poorer the resolution, the more judgment calls there are to be made:
Nevertheless, most chemists who undertake structure-based design treat a protein crystal structure reverently as if it was determined at very high resolution, regardless of the resolution at which the structure was actually determined (admittedly, crystallographers themselves are not immune to this practice either). Also, the fact that the crystallographer is bound to have made certain assumptions, to have had certain biases and perhaps even to have made mistakes is usually ignored. Assumptions, biases, ambiguities and mistakes may manifest themselves (even in high-resolution structures) at the level of individual atoms, of residues (e.g. sidechain conformations) and beyond.
Then there’s the problem of interpreting how your drug candidate interacts with the protein. The ability to get an X-ray structure doesn’t always correlate well with the binding potency of a given compound, so it’s not like you can necessarily count on a lot of clear signals about why the compound is binding. Hydrogen bonds may be perfectly obvious, or they can be rather hard to interpret. Binding through (or through displacement of) water molecules is extremely important, too, and that can be hard to get a handle on as well.
And not least, there’s the assumption that your structure is going to do you good once you’ve got it nailed down:
It is usually tacitly assumed that the conditions under which the complex was crystallised are relevant, that the observed protein conformation is relevant for interaction with the ligand (i.e. no flexibility in the active-site residues) and that the structure actually contributes insights that will lead to the design of better compounds. While these assumptions seem perfectly reasonable at first sight, they are not all necessarily true. . .
That’s a key point, because that’s the sort of error that can really lead you into trouble. After all, everything looks good, and you can start to think that you really understand the system, that is until none of your wonderful X-ray-based analogs work out they way you thought they would. The authors make the point that when your X-ray data and your structure-activity data seem to diverge, it’s often a sign that you don’t understand some key points about the thermodynamics of binding. (An X-ray is a static picture, and says nothing about what energetic tradeoffs were made along the way). Instead of an irritating disconnect or distraction, it should be looked at as a chance to find out what’s really going on. . .
Comments (15)
+ TrackBacks (0) | Category: Analytical Chemistry | Drug Assays | In Silico
March 27, 2008
Posted by Derek
There’s an excellent paper in the most recent issue of Chemistry and Biology that illustrates some of what fragment-based drug discovery is all about. The authors (the van Aalten group at Dundee) are looking at a known inhibitor of the enzyme chitinase, a natural product called argifin. It’s an odd-looking thing – five amino acids bonded together into a ring, with one of them (an arginine) further functionalized with a urea into a sort of side-chain tail. It’s about a 27 nM inhibitor of the enzyme.
(For the non-chemists, that number is a binding affinity, a measure of what concentration of the compound is needed to shut down the enzyme. The lower, the better, other things being equal. Most drugs are down in the nanomolar range – below that are the ulta-potent picomolar and femtomolar ranges, where few compounds venture. And above that, once you get up to 1000 nanomolar, is micromolar, and then 1000 micromolar is one millimolar. By traditional med-chem standards, single-digit nanomolar = good, double-digit nanomolar = not bad, triple-digit nanomolar or low micromolar = starting point to make something better, high micromolar = ignore, and millimolar = can do better with stuff off the bottom of your shoe.
What the authors did was break this argifin beast up, piece by piece, measuring what that did to the chitinase affinity. And each time they were able to get an X-ray structure of the truncated versions, which turned out to be a key part of the story. Taking one amino acid out of the ring (and thus breaking it open) lowered the binding by about 200-fold – but you wouldn’t have guessed that from the X-ray structure. It looks to be fitting into the enzyme in almost exactly the same way as the parent.
And that brings up a good point about X-ray crystal structures. You can’t really tell how well something binds by looking at one. For one thing, it can be hard to see how favorable the various visible interactions might actually be. And for another, you don’t get any information at all about what the compound had to pay, energetically, to get there.
In the broken argifin case, a lot of the affinity loss can probably be put down to entropy: the molecule now has a lot more freedom of movement, which has to be overcome in order to bind in the right spot. The cyclic natural product, on the other hand, was already pretty much there. This fits in with the classic med-chem trick of tying back side chains and cyclizing structures. Often you’ll kill activity completely by doing that (because you narrowed down on the wrong shape for the final molecule), but when you hit, you hit big.
The structure was chopped down further. Losing another amino acid only hurt the activity a bit more, and losing still another one gave a dipeptide that was still only about three times less potent than the first cut-down compound. Slicing that down to a monopeptide, basically just a well-decorated arginine, sent the activity down another sixfold or so – but by now we’re up to about 80 micromolar, which most medicinal chemists would regard as the amount of activity you could get by testing the lint in your pocket.
But they went further, making just the little dimethylguanylurea that’s hanging off the far end. That thing is around 500 micromolar, a level of potency that would normally get you laughed at. But wait. . .they have the X-ray structures all along the way, and what becomes clear is that this guanylurea piece is binding to the same site on the protein, in the same manner, all the way down. So if you’re wondering if you can get an X-ray structure of some 500 micromolar dust bunny, the answer is that you sure can, if it has a defined binding site.
And the value of these various derivatives almost completely inverts if you look at them from a binding efficiency standpoint. (One common way to measure that is to take the minus log of the binding constant and divide by the molecular weight in kilodaltons). That’s a “bang for the buck” index, a test of how much affinity you’re getting for the weight of your molecule. As it turns out, argifin – 27 nanomolar though it be – isn’t that efficient a binder, because it weighs a hefty 676. The binding efficiency index comes out to just under 12, which is nothing to get revved up about. The truncated analogs, for the most part, aren’t much better, ranging from 9 to 15.
But that guanylurea piece is another story. It doesn’t bind very tightly, but it bats way above its scrawny size, with a BEI of nearly 28. That’s much more impressive. If the whole argifin molecule bound that efficiently, it would be down in the ten-to-the-minus nineteenth range, and I don’t even know the name of that order of magnitude. If you wanted to make a more reasonably sized molecule, and you should, a compound of MW 400 would be about ten femtomolar with a binding efficiency like that. There’s plenty of room to do better than argifin.
So the thing to do, clearly, is to start from the guanylurea and build out, checking the binding efficiency along the way to make sure that you’re getting the most out of your additions. And that is exactly the point of fragment-based drug discovery. You can do it this way, cutting down a larger molecule to find what parts of it are worth the most, or you can screen to find small fragments which, though not very potent in the absolute sense, bind very efficiently. Either way, you take that small, efficient piece as your anchor and work from there. And either way, some sort of structural read on your compounds (X-ray or NMR) is very useful. That’ll give you confidence that your important binding piece really is acting the same way as you go forward, and give you some clues about where to build out in the next round of analogs.
This particular story may be about as good an illustration as one could possibly find - here's hoping that there are more that can work out this way. Congratulations to van Aalten and his co-workers at Dundee and Bath for one of the best papers I've read in quite a while.
Comments (12)
+ TrackBacks (0) | Category: Analytical Chemistry | Drug Assays | In Silico
January 22, 2008
Posted by Derek
There’s been a big trend the last few years in the industry to try to build our molecules up from much smaller pieces than usual. “Fragment-based” drug discovery is the subject of many conferences and review articles these days, and I’d guess that most decent-sized companies have some sort of fragment effort going on. (Recent reviews on the topic, for those who want them).
Many different approaches come under that heading, though. Generally, the theme is to screen a collection of small molecules, half the size or less of what you’d consider a reasonable molecular weight for a final compound, and look for something that binds. At those sizes, you’re not going to find the high affinities that you usually look for, though. We usually want our clinical candidates to be down in the single-digit nanomolar range for binding constants, and our screening hits to be as far under one micromolar as we can get. In the fragment world, though, from what I can see, people regard micromolar compounds as pretty hot stuff, and are just glad not to be up in the millimolar range. (For people outside the field, it’s worth noting that a nanomolar compound binds about a million times better than a millimolar one).
Not all the traditional methods of screening molecules will pick up weak binders like that. (Some assays are actually designed not to read out at those levels, but to only tell you about the really hot compounds). For the others, you’d think you could just run things like you usually do, just by loading up on the test compounds, but that’s problematic. For one thing, you’ll start to chew up a lot of compound supplies at that rate. Another problem is that not everything stays in solution for the assay when you try to run things at that concentration. And if you try to compensate by using more DMSO or whatever to dissolve your compounds, you can kill your protein targets with the stuff when it goes in. Proteins are happy in water (well, not pure distilled water, but water with lots of buffer and salts and junk like the inside of a cell has). They can take some DMSO, but it’ll eventually make even the sturdiest of them unhappy at some point. (More literature on fragment screening).
And once you’ve got your weak-binding low-molecular weight stuff, what then? First, you have to overcome the feeling, natural among experienced chemists, that you’re working on stuff you should be throwing away. Traditional medicinal chemistry – analog this part, add to that part, keep plugging away – may not be the appropriate thing to do for these leads. There are just too many possibilities – you could easily spend years wandering around. So many companies depend on structural information about the protein target and the fragments themselves to tell them where these little guys are binding and where the best places to build from might be. That can come from NMR studies or X-ray crystal determinations, most commonly.
Another hope, for some time now, has been that if you could discover two fragments that bound to different sites, but not that far from each other, that you could then stitch them together to make a far better compound. (See here for more on this idea). That’s been very hard to realize in practice, though. Finding suitable pairs of compounds is not easy, for starters. And getting them linked, as far as I can see, can be a real nightmare. A lot of the linking groups you can try will alter the binding of the fragments themselves – so instead of going from two weak compounds to one strong one, you go from two weak ones to something that’s worse than ever. Rather than linking two things up, a lot of fragment work seems to involve building out from a single piece.
But that brings up another problem, exemplified by this paper. These folks took a known beta-lactamase inhibitor, a fine nanomolar compound, and broke it up into plausible-looking fragments, to see if it could have been discovered that way. But what they found, each time they checked the individual pieces, was that each of them bound in a completely different way than it did when it was part of the finished molecule. The binding mode was emergent, not additive, and it seems clear that most (all?) of the current fragment approaches would have been unable to arrive at the final structure. The authors admit that this may be a special case, but there’s no reason to assume that it’s all that special.
So fragment approaches, although they seem to be working out in some cases, are probably always going to miss things. But hey, we miss plenty of things with the traditional methods, too. Overall, I’m for trying out all kinds of odd things, because we need all the help we can get. Good luck to the fragment folks.
Comments (6)
+ TrackBacks (0) | Category: Analytical Chemistry | Drug Assays
October 25, 2006
Posted by Derek
There's an interesting analytical chemistry paper in the preprint section of PNAS (open access if you want to read it) that may reopen an old controversy. It's from a large multinational team (Mexico, Spain, France, NASA-Ames) investigating the GC-mass spec instrumentation that was flown to Mars on the Viking landers in 1976. That's a key instrument in the life-on-Mars debate, so an attack on it is significant. First, though, some background - it's a tangled story.
The Viking landers each had three biology experiments to look for possible signs of Martian life, whose results were famously difficult to interpret. They produced both excitement and confusion at the time (scroll down in that NASA history page) and they've been fuel for arguments ever since.
There was the pyrolytic release experiment, which incubated Martian soil with 14C-labled carbon monoxide and carbon dioxide. After several days, the sample was purged, then heated to 650C and analyzed for the release of any labeled carbon compounds that might have been formed by living organisms. A control sample was heated before incubation, to kill off any such life forms. Seven out of the nine runs of this experiment seemed to produce positive results - that is, volatile labeled carbon was produced after pyrolysis.
The gas-exchange experiment used the same sort of apparatus, exposing the soil to either water vapor or nutrient solution under a mixed atmosphere of gases. The headspace was analyzed for changes in the concentrations of the various components, which could be due to biological uptake or release. This one showed a strong release of oxygen and carbon dioxide from the samples once moisture was added, but the amount decreased over time, leading to theories that this was the product of an inorganic reaction rather than a signature of life.
The labeled release experiment put Martian soil into a dilute nutrient broth, with several small organic compounds which were all labeled with 14C. After incubation, the headspace of the experimental cell was analyzed for any released labeled gases and again, a control experiment was done with pre-heated soil. This one produced exciting data, with release of labeled gas in the experimental samples well over those in the controls. One odd result, though, was that the subsequent injection(s) of nutrient solution did not produce a further spike of released gas. The final curves ended up looking neither like what you'd have expected from a classic bacterial positive, nor from a simple chemical reaction. This ambiguity has meant that the LR results have been re-analyzed and re-fought ever since the 1970s, with the experiment's designer, Gilbert Levin, leading the effort to rescue the data as a case for Martian life.
But then there were the GC-MS data, from an experiment considered to be the backstop test in case the biology experiments were difficult to interpret. Since they certainly were that, from beginning to end, this experiment became for many people the most important one on the landers. (It already had been for the people - a not insignificant group - who thought from the start that the biology tests were unlikely to provide a conclusive answer). This one heated soil samples directly and looked for volatile organics. Heating to 200C showed little or nothing in the way of carbon compounds, and very little water besides. By contrast, another sample taken up to 500 degrees released a comparative flood of water, but still showed no evidence of organic molecules.
And that, for most observers, was that. No organic molecules, no life. Explanations after the GC-MS results mainly turned to what sorts of inorganic chemistry might have given the behavior seen in the three other experiments. Martian soil was thus hypothesized to be a sterile mixture of interesting chemicals (iron peroxides? carbon suboxide polymers?) that had fooled the biology test packages, but couldn't fool the GC-MS.
There's always been an underground, though, that has held that the results were indeed the result of life. Gilbert Levin has never given up. In 1981, he pointed out that tests of a Viking-style GC-MS instrument had shown that it was insensitive to organics in a particular Antarctic soil sample, but that this same soil nonetheless gave a positive result in the LR experiment. And he really put his opinions out in the store window in 1997, with a paper that flatly concluded that the 1976 LR experiments had indeed detected Martian life.
In the last few years, others have joined the battle. Steven Benner at Florida, whose work I wrote about here, published a PNAS paper in 2000 which maintained that organic molecules on Mars would likely be retained as higher molecular weight carboxylates, which would not have been volatile enough for the Viking GC/MS instrument to detect. And now this latest group has weighed in.
They've also analyzed various Antarctic and temperate desert samples, and found that all of them contain organic matter that cannot be detected by thermal GC-MS analysis. And the ones that contain iron, including the NASA reference simulated Mars soil (a weathered basalt sample from near Mauna Kea), tend to oxidize their organics quickly under heating. The conclusion is that while much of the water and carbon dioxide produced in the Viking experiment from heating the Martian soil was surely inorganic, some of it could have been from the oxidation of organic material. The paper concludes that the Viking GC-MS results are. . .inconclusive, and should not be taken as evidence either way for the presence of organic molecules or life. The question, they feel, is still completely open.
The good news is that future missions are relying on other technologies. In addition to good ol' thermal volatilization/GC-MS, there are also plans for solvent extractions, laser desorption mass spec, short-path sublimation, and other nifty ideas. If these various US and European missions get off the ground (and on the Martian ground), we're going to have some very interesting data to look at. And argue about.
Comments (6)
+ TrackBacks (0) | Category: Analytical Chemistry | Life As We (Don't) Know It
August 27, 2006
Posted by Derek
After my article on the role of carbon isotope testing in the Floyd Landis case, a question has come up several times in the comments and in my e-mail: since it's well-known that Landis was taking cortisone for his hip, could this have skewed the isotope ratios in his testosterone?
I doubt it very much, and here's why: first off, around 95% of the circulating testosterone in the male body is produced in the testes. For Landis's isotope ratios to be off a significant amount through something involving his own metabolic pathways, this is the only place that's worth looking.Testosterone and the other steroids are produced from cholesterol. The testes and other steroidogenic tissues have a stockpile of cholesteryl esters ready to be used for steroid synthesis, so it's going to be an uphill fight to alter things by any route, given that reserve.
Now it's time to dive into some biochemistry for the next few paragraphs - follow along if you like, or jump down to near the end if you don't want to see a lot of structures. OK, in steroid synthesis the first thing that happens is the chewing off of a side chain on the D ring to form pregnenolone, which is then turned into progesterone. That's the starting material for both testosterone and cortisol/cortisone. (Note that those last two are interconverted in the body by the 11-HSD enzymes).
Going down these different pathways, testosterone and cortisol end up with rather different structures. Cortisol's more complex. If you flip back and forth between those links in the previous paragraph, you'll see that the A and B rings are the same in both, but the C ring of cortisol has an extra hydroxyl group at C11, and it also has some oxidized side chain left at C17, which has been completely chopped off in testosterone. The question is, can you get from cortisol back to something that could be used to make testosterone?
I can believe the side-chain transformation much easier than the C-11 deoxygenation. Here's the metabolic fate of cortisol. Note that all these metabolites still have an oxidized C-11 - if anything is going to be recycled into testosterone, that C-11 is going to have to be reduced back down. And if there's a metabolic pathway that does that to any degree, I can't seem to find out anything about it. If it's a feasible pathway at all, it must be very minor indeed. If any steroid experts can shed light on this, I'd be glad to hear the details. (There's also the question of how long such intermediates would be available, versus their half-life before further metabolism and excretion, but that's a whole other issue).
No, if Landis's carbon isotope ratios are off significantly - and we haven't seen the official numbers yet - then it's hard for me to see how the cortisone injections could have much to do with it. We'll be stuck, in that case, with either conspiracy theories or with the conclusion that Landis used testosterone, and if it comes to that, I know which one I'm most likely to believe.
Comments (22)
+ TrackBacks (0) | Category: Analytical Chemistry | Current Events
August 1, 2006
Posted by Derek
The New York Times broke the story today that the testosterone found in Tour de France champion Floyd Landis's blood was not from a natural source. Just how do they know that, and how reliable is the test?
The first thing an anti-doping lab looks for in such a case is the ratio of testosterone to the isomeric epitestosterone - too high an imbalance is physiologically unlikely and arouses suspicion. Landis already is in trouble from that reading, but the subject of the Times scoop is the isotopic ratio of the testosterone itself. And that one is going to be hard to get away from, if it's true.
Update: people are asking me why athletes don't just take extra epistestosterone to even things out. That they do - that's the most basic form of masking, and if Landis's ratio was as far off as is being reported, it's one of the odd features about this case. But the isotope test will spot either one, if it's not the kind your body produces itself - read on.
Steroids, by weight, are mostly carbon atoms. Most of the carbon in the world is the C-12 isotope, six protons and six neutrons, but around one per cent of it has an extra neutron to make it C-13. Those are the only stable isotopes of carbon. You can find tiny bits of radioactive C-14, though, and you can also get C-11 if you have access to a particle accelerator. Work fast, though, because it's hot as a pistol.
So, testosterone has 19 carbon atoms, and if on average every one out of a hundred carbon atoms is a C-13, you can calculate the spread of molecular weights you could expect, and their relative abundance. One out of every ten thousand molecules would have two C-13 atoms in there somewhere, one out of every million or so would have three, and so on. A good mass spectrometer will lay this data out for you like a deck of cards.
But here's the kicker: those isotopic forms of the elements behave a bit differently in chemical reactions. The heavier ones do the same things as their lighter cousins, but if they're involved in or near key bond-breaking or bond-making steps, they do them more slowly. It's like having a heavier ball attached to the other end of a spring. This is called a kinetic isotope effect, and chemists have found all sorts of weird and ingenious ways to expoit it. But it's been showing up for a lot longer than we've been around.
The enzymatic reactions that plants and bacteria use when they take up or form carbon dioxide have been slowly and relentlessly messing with the isotope ratios of carbon for hundreds of millions of years. And since decayed plants are food for other plants, and the living plants are food for animals, which are food for other animals and fertilizer for still more plants. . .over all this time, biological systems have become enriched in the lighter, faster-reacting C-12 isotope, while the rest of the nonliving world has become a bit heavier in C-13. You can sample the air next to a bunch of plants and watch as they switch from daytime photosynthesis to nighttime respiration, just based on the carbon isotope ratios. Ridiculously tiny variations in these things can now be observed, which have led to all sorts of unlikely applications, from determining where particular batches of cocaine came from to figuring out the dietary preferences of extinct herbivores.
So, if your body is just naturally cranking out the testosterone, it's going to have a particular isotopic signature. But if you're taking the synthetic stuff, which has been partly worked on with abiotic forms of carbon derived from a different source (see below), the fingerprints will show. (Update: yes, this means that the difference between commercial testosterone and the body's own supply isn't as large as it would be otherwise, since the commercial synthesis generally starts from plant-derived steroid backbones. But it's still nothing that a good mass spec lab would miss). If the news reports are right, that's what Landis's blood samples have shown. And if they have, there seems only one unfortunate conclusion to be drawn.
Chem-Geek Supplemental Update: for the folks who have been wondering where exactly the isotopic difference comes in, here's the story: synthetic testosterone is made from phytosterol percursors, typically derived from wild yams or soy. Those are both warm-climate C3 plants, which take up atmospheric carbon dioxide by a different route than temperate-zone C4 plants, leading to noticeably different isotope ratios. That's where all the isotope-driven studies of diet start from. The typical Western industrial-country diet is derived from a mixture of C3 and C4 stocks, so the appearance of testosterone with a C3-plant isotopic profile is diagnostic.
Comments (295)
+ TrackBacks (0) | Category: Analytical Chemistry | Current Events
June 3, 2004
Posted by Derek
Another day spent rooting around in the archives, trying to appease the rapacious Taiwanese patent office. One more day should about do it, and not a moment too soon. I'm now unearthing NMR spectral data for compounds, and translating those to print is not enjoyable.
For those outside the field, an NMR spectrum of a typical organic molecule is a rather complex linear plot of multiple lines and peaks. After staring at it a while, it gets rendered into text as something like "1.63, t, 3H; 2.34, s, 3H; 3.1 - 3.39, m, 4H. . ." In plain text, that's "At 1.63 and 2.34, there are a triplet signals that represent three protons each, and between 3.1 and 3.39 there's a messy multiplet that adds up to four protons' worth. . ."
If you really want to get into it, you list the coupling constants, the spacings between the individual peaks of those triplets and etc. No thanks. A typical spectrum will go on for a reasonable paragraph in this way, and the Taiwanese would like nothing better than several pages of this sort of thing, or so they maintain. What they'll is get as much as I can stand.
I'll try to lead off next week with a discussion of today's news about everyone's pal, Elliot Spitzer, and his suit against GSK. It's a wide-ranging topic, and there wasn't enough time to wrestle it to the ground today.
Comments (2)
+ TrackBacks (0) | Category: Analytical Chemistry | Patents and IP
February 1, 2002
Posted by Derek
I can't talk about rain forest drug discovery without mentioning the (pretty bad) 1992 Sean Connery movie Medicine Man. He plays an alleged biochemist who comes up with a Miracle Drug, more or less by finding it under a leaf.
Plenty of large and small stuff is misportrayed, but I did say it was a movie. (One of these days someone will have to make a list of jobs that movies actually get right.) The part of this one that drug discovery people particularly enjoyed, though, was when some crude extract is fed into an impressive device that immediately displays the structure of the active compound. "I want one of those!" was the universal reaction.
I believe that this was supposed to be the one active component of a complex mixture, the plot hinging on being able to find it and isolate it. Of course, Shaman's business model (see previous post) depended on being able to do this sort of thing, and you see where it got them.
I only wish we could find things out as suddenly and dramatically as they do in films like these. As is true in most areas of research, medicinal chemists spend a fair amount of time looking at printouts (or up at the ceiling tiles,) wondering just what the heck happened in the last experiment. Determining chemical structures is easier than it's ever been (read: in most cases, it's possible to do it), but for natural products it still isn't trivial. My Connery-ometer remains on back-order.
Comments (0)
+ TrackBacks (0) | Category: Analytical Chemistry
|
|
