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About this Author
Derek Lowe
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

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March 2, 2010

Why You Don't Want to Make Death-Star-Sized Drugs

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Posted by Derek

I was just talking about greasy compounds the other day, and reasons to avoid them. Right on cue, there's a review article in Expert Opinion in Drug Discovery on lipophilicity. It has some nice data in it, and I wanted to share a bit of it here. It's worth noting that you can make your compounds too polar, as well as too greasy. Check these out - the med-chem readers will find them interesting, and who knows, others might, too:
MW350%20graph%20jpeg.jpg
MW500%20graph%20jpeg.jpg
So, what are these graphs? They show how well compound cross the membranes of Caco-2 cells, a standard assay for permeability. These cells (derived from human colon tissue) have various active-transport pumps going (in both directions), and you can grow them in a monolayer, expose one side to a solution of drug substance, and see how much compound appears on the other side and how quickly. (Of course, good old passive diffusion is also operating, too - a lot of compounds cross membranes by just soaked on through them).

Now, I have problems with extrapolating Caco-2 data too vigorously to the real world - if you have five drug candidates from the same series and want to rank order them, I'd suggest getting real animal data rather than rely on the cell assay. The array of active transport systems (and their intrinsic activity) may well not match up closely enough to help you - as usual, cultured cell lines don't necessarily match reality. But as a broad measure of whether a large set of compounds has a reasonable chance of getting through cell membranes, the assay's not so bad.

First, we have a bunch of compounds with molecular weights between 350 and 400 (a very desirable space to occupy). The Y axis is the partitioning between the two sides of the cells, and X axis is LogD, a standard measure of compound greasiness. That thin blue line is the cutoff for 100 nanomoles/sec of compound transport, so the green compounds above it travel across the membrane well, and the red ones below it don't cross so readily. You'll note that as you go to the left (more and more polar, as measured by logD), the proportion of green compounds gets smaller and smaller. They're rather hang out in the water than dive through any cell membranes, thanks.

So if you want a 50% chance of hitting that 100 nm/sec transport level, then you don't want to go much more polar than a LogD of 2. But that's for compounds in the 350-400 weight range - how about the big heavyweights? Those are shown in the second graph, for compounds greater than 500. Note that the distribution has scrunched disturbingly. Now almost everything is lousy, and if you want that 50% chance of good penetration, you're going to have to get up to a logD of at least 4.5.

That's not too good, because you're always fighting a two-front war here. If you make your compounds that greasy (or more) to try to improve their membrane-crossing behavior, you're opening yourself up (as I said the other day) to more metabolic clearance and more nonspecific tox, as your sticky compounds glop onto all sorts of things in vivo. (They'll be fun to formulate, too). Meanwhile, if you dip down too far into that really-polar left-hand side, crossing your fingers for membrane crossing, you can slide into the land of renal clearance, as the kidneys vacuum out your water-soluble wonder drug and give your customers very expensive urine.

But in general, you have more room to maneuver in the lower molecular weight range. The humungous compounds tend to not get through membranes at reasonable LogD values. And if you try to fix that by moving to higher LogD, they tend to get chewed up or do unexpectedly nasty things in tox. Stay low and stay happy.

Comments (24) + TrackBacks (0) | Category: Drug Assays | Pharma 101 | Pharmacokinetics

November 19, 2009

What Are the Best Med-Chem Books?

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Posted by Derek

I get regular requests to recommend books on various aspects of medicinal chemistry and drug development. And while I have a few things on my list, I'm sure that I'm missing many more. So I wanted to throw this out to the readership: what do you think are the best places to turn? This way I can be more sure of pointing people in the right directions.

I'm interested in hearing about things in several categories - best introductions and overviews of the field (for people just starting out), as well as the best one-stop references for specific aspects of drug discovery (PK, toxicology, formulations, prodrugs, animal models, patent issues, etc.)

Feel free to add your suggestions in the comments, or e-mail them to me. I'll assemble the highest-recommended volumes into a master list and post that. Just in time for the holidays, y'know. . .

Comments (35) + TrackBacks (0) | Category: Life in the Drug Labs | Pharma 101

August 13, 2009

Animal Testing: A View From the Labs

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Posted by Derek

Why do we test drugs on animals, anyway? This question showed up in the comments section from a lay reader. It's definitely a fair thing to ask, and you'd expect that we in the business would have a good answer. So here it is: because for all we know about biochemistry, about physiology and about biology in general, living systems are still far too complex for us to model. We're more ignorant than we seem to be. The only way we can find out what will happen if we give a new compound to a living creature is to give it to some of them and watch carefully.

That sounds primitive, and I suppose it is. We don't do it in a primitive manner, though. We watch with all the tools of our trade - remote-control physiological radio transmitters, motion-sensing software hooked up to video cameras, sensitive mass spectrometry analysis of blood, of urine, and whatever else, painstaking microscopic inspection of tissue samples, whatever we can bring to bear. But in the end, it all comes down to dosing animals and waiting to see what happens. That principle hasn't changed in decades, just the technology we use to do it.

No isolated enzymes can yet serve as a model for what can happen in a single real cell. And no culture of cells can recapitulate what goes on in a real organism. The signaling, the feedback loops, the interconnectedness of these systems is (so far) too much for us to handle. We keep discovering new pathways all the time, things that no model would have included because we didn't even know that they were there. The end is not yet in sight, occasional newspaper headlines to the contrary.

We do use all those things as filters before a compound even sees its first rodent. In a target-driven approach, which is the great majority of the industry, if a compound doesn't work on an isolated protein, it doesn't go on to the cell assay. If it doesn't work on the cells, it doesn't go on to animals. (And if it kills cells, it most certainly doesn't go on to the animals, unless it's some blunderbuss oncology agent of the old school). The great majority of compounds made in this business have never been given to so much as one mouse, and never will.

So what are we looking for when we finally do dose animals? We're waiting to see if the compound has the effect we're hoping for, first off. Does it lower blood pressure, slow or stop the growth of tumors, or cure viral infections? Doing these things requires having sick animals, of course. But we also give the drug candidates to healthy ones, at higher doses and for longer periods of time, in order to see what else the compounds might do that we don't expect. Most of those effects are bad - I'd casually estimate 99% of the time, anyway - and many of them will stop a drug candidate from ever being developed. The more severe the toxic effect, the greater the chance that it's based on some fundamental mechanism that will be common to all animals. In some cases we can identify what's causing the trouble, once we've seen it, and once in a great while we can use that information to argue that we can keep going, that humans wouldn't be at the same risk. But this is very rare - we generally don't know enough to make a persuasive case. If your compound kills mice or kills rats, your compound is dead, too.

I've lost count of the number of compounds I've worked on that have been pulled due to toxicity concerns; suffice it to say that it's a very common thing. Every time it's been something different, and it's often not for any of the reasons I feared beforehand. I've often said here that if you don't hold your breath when your drug candidate goes into its first two-week tox testing, then you haven't been doing this stuff long enough.

Here's the problem: giving new chemicals to animals to see if they get sick (and making animals sick so that we can see if they get better) are not things that are directly compatible with trying to keep animals from suffering. Ideally, we would want to do neither of those things. Fortunately, several factors all line up in the same direction to keep things moving toward that.

For one thing, animal testing is quite expensive. Only human testing is costlier. In this case, ethical concerns and capitalist principles manage to line up very well indeed. Doing assays in vitro is almost invariably faster and cheaper, so whenever we can confidently replace a direct animal observation with an assay on a dish, plate, or chip, we do. All that equipment I mentioned above has also cut down on the number of animals needed, and that trend is expected to continue as our measurements become more sensitive.

So things are lined up in the right direction. Any company that found a reliable way to eliminate any significant part of its animal testing would immediately find itself in a better competitive position.

And for the existing tests, it's also fortunate that unhappy animals give poor data. We want to observe them under the most normal conditions possible, not with stress hormones running through their systems, and a great deal of time and trouble (and money) goes toward that end. (In this case, it's scientific principles that line up with ethical ones). Diseased animals are clearly going to be in worse shape than normal ones, but in these situation, too, we try to minimize all the other factors so we're getting as clear a read as possible on changes in the disease itself.

So that's my answer: we use animals because we have (as yet) no alternative. And our animal assays prove that to us over and over by surprising us with things we didn't know, and that we would have had no other opportunity to learn. We'd very much like to be able to do things differently, since "differently" would surely mean "faster and more cheaply". None of us enjoy it when our compounds sicken healthy animals, or have no effect on sick ones. Just the wasted time and effort alone is enough to make any drug discoverer think so. There are billions of dollars waiting to be picked up by anyone who finds a better way.

Comments (79) + TrackBacks (0) | Category: Animal Testing | Pharma 101

February 6, 2009

Prep TLC: The Good Old Days Live On

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Posted by Derek

I did something in the lab the other day that I hadn’t done in several years: run some preparative TLC plates. I had some small reactions that needed to be cleaned up, and the HPLC systems were all in use, so I thought “Why not?” (I wrote here about the decline of analytical TLC in general in some labs, and I think it's fair to say that the larger-scale prep version has seen an even steeper drop in use over the years).

Prep TLC, for those of you not in the business, is a pretty simple technique. You take a square glass plate that’s been coated with a dry layer of ground silica, a white slurry that for this application is about the grittiness of flour or ground sugar. You then take your mixture of gunk, dissolve it up in a small volume of solvent, and deposit it in a line across the bottom of the plate, an inch or so up from one side and parallel to it. Then you take a large glass container and add some solvent to the bottom of it, and put your plate in so that the streaked line of material is near the bottom. Here's one running.

The solvent soaks into the layer of silica, and after it gets up an inch or so it hits your line of stuff. As it continues to move up, soaking further and further up the glass plate, the different components of the mixture will be carried along at different rates. The compounds that stick to silica gel (for one reason or another) will lag behind, while the ones that don’t will move out into the lead. After an hour or so, the solvent line will be up near the top of the plate, and your mixture will now be spread out across it into a series of bands. (The TLC page at Wikipedia has some useful images of this). Up at the top, running with the solvent, will the the nonpolar stuff that didn’t have anything to slow it down. Right down near the bottom, not far up from your original streak, will be the most polar stuff, especially any basic amines – silica gel is mildly acidic, so the amines will stick to it very tightly indeed. And in between will be the other components, divided out according to how they balanced out the pull of the silica gel support with the attraction of the solvent moving them along. Sometimes you can see them as colored bands on the silica plate, but more often you shine a UV light on the whole plate to see them. The silica we use has an ingredient that makes it fluoresce green under ultraviolet, and our compounds usually show up as dark blue or purple bands against the green. It’s a color combination known to every working synthetic organic chemist.

You can see that picking different solvents for this process can change things a great deal. A weak solvent (like hexane) will allow almost everything to stick to the silica. (A compound has to be mighty greasy to be swept along by just hexane; I doubt if there’s a drug in the business that you’d be able to clean up that way). A standard mix is some proportion of ethyl acetate mixed with hexane. You can go up to straight ethyl acetate, or even further by mixing in methanol or the like. And if you’re desperate, you can go to most any solvent mixture you like – three-solvent brews, toluene, acetonitrile, acetone, whatever works.

So how do you get the things off? By the lowest-tech method you can imagine. You mark the position of the band (or bands) you want, and then take a metal spatula and scrape the silica there off the plate. You them dump that into a flask and stir it with a strong solvent, then filter off the silica and wash it some more to rinse your compound out.

This used to be much more of an everyday technique, but automated column chromatography (same principle, pumped through a tube) has taken over. But prep TLC still has its appeal. Done with skill, it can provide very clean compounds, with quite good recovery. In fact, its low cost and power have made it a favorite technique at places like WuXi, the outsourcing powerhouse in China. I've had several first-hand descriptions of their prep TLC room, with rows of plates being run, marked, and scraped in assembly-line fashion. It's the sort of thing you'd only do in a cheap-labor market, because of the unavoidable hand work involved, but it is effective.

I don't know where WuXi gets its plates, but if you make your own, it's an even cheaper technique (discounting labor costs, naturally). You take up the silica gel powder in water, make a thick, well-mixed slurry out of it, and spread it across a square of glass, shaking and tapping it to get the air bubbles out. Back when I was doing summer undergraduate work, I poured a number of these things, although it's certainly nothing I've had experience with since the first Reagan administration. For all I know, that's how WuXi does it now. Perhaps they've found a low-cost supplier of their own, but the idea of a cheap supplier for a Chinese outsourcing company is an interesting one all by itself. . .

Comments (31) + TrackBacks (0) | Category: Life in the Drug Labs | Pharma 101

December 10, 2008

Floppiness Is Not Your Friend: Who Knew?

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Posted by Derek

There’s a trick that every medicinal chemist learns very early, and continues to apply every time its feasible: take two parts of your compound, and tie them together into a ring.

The reason that works so well may not be immediately obvious if you’re not a medicinal chemist, so let me expand on them a bit. The first thing to know is that this method tends to work either really well or not at all – it’s a “death or glory” move. And that gives you a clue as to what’s going on. The idea is that the rotatable bonds in your molecule are, under normal conditions, doing just that: rotating. Any molecule the size of a normal drug has all kinds of possible shapes and rotational isomers, and room temperature is an energetic enough environment to populate a lot of them.

But there’s only one of them that’s the best for fitting into your drug target, most likely. So what are the odds? As your molecule approaches its binding pocket, there’s a complicated energetic dance going on. Different parts of your drug candidate will start interacting with the target (usually a protein), and that starts to tie down all that floppy rotation. The question is, does the gain resulting from these interactions cancel out the energetic price that has to be paid for them? Is there a pathway that leads to a favorable tight-binding situation, or is your molecule going to approach, flop around a bit, and dance away?

Several things are at work during that shall-we-dance period. The different conformations of your compound vary in energy, depending on how much its parts are starting to bang into each other, and how much you’re asking the bonds to twist around. The closer that desired drug-binding shape is to the shape your molecule wants to be in anyway, the better off you are, from that perspective. So tying back the molecule and making a ring in the structure does one thing immediately: it cuts down on the range of conformations it can take, in the same way that tying a rope between your ankles cuts down on your ability to dance. You’ve handcuffed your molecule, which would probably be cruel if they were sentient, but then, a lot of organic chemistry would be pretty unspeakable if molecules had feelings.

That’s why this method tends to be either a big winner or a big loser. If the preferred binding mode of your compound is close to the shape it takes when you tie it down, then you’ve suddenly zeroed in on just the thing you want, and the binding affinity is going to take a big leap. But if it’s not, well, you’ve now probably made it impossible for the thing to adopt the conformation it needs, and the binding affinity is going to take a big leap over a cliff.

There’s another effect to reducing the flexibility of your compound, and that has to do with entropy. All that favorable-interaction business is one component of the energy involved, namely the enthalpy, but entropy is the other. Loosely speaking, the more disordered a system, the higher its entropy. A floppy molecule, when it binds to a drug target, has to settle down into a much tighter fit, and entropically, that’s unfavorable. Energetically, you’re paying to do that. But if your molecule is already much less flexible, there’s not much of a toll as it fits into the pocket. If loss-of-floppiness is a bad thing, then don’t start out with so much of it.

So, how much do I and my medicinal chemistry colleagues think about this stuff, day to day? A fair amount, but there are parts of it that we probably don’t pay enough attention to. Entropy gets less respect from us than it deserves, I think. It’s easy to imagine molecules bumping into each other, sticking and unsticking, but the more nebulous change-in-disorder part of the equation is just as important. And it doesn’t just apply to our drug molecules – proteins get less disordered as they bind those molecules (or more disordered, in some cases), and those entropic changes can mean a lot, too.

I also mentioned molecules finding a pathway to binding, and that’s something that we don’t think about as much, either. We probably make things all the time that would be potent binders, if they just could get past some energetic hump and wedge themselves into place. But there are no crowbars available; our drug candidates have to be able to work their way in on their own. The can’t-get-there-from-here cases come back from the assays as inactive. The tendency is to imagine these in the binding site already, and to try to think of what could be going wrong in there – but it may be that they’d be fine, but that their structures won’t allow them to come in for a landing.

Picturing this accurately is very hard indeed. We have enough trouble with good representations of static pictures of our molecules bound to their targets, so making a movie of the process is a whole different story. Each frame is on a femtosecond scale – molecules flip around rather quickly – and every frame would have to be computed accurately (drug structure, protein structure, and the energetics of the whole system) for the resulting video clip to make sense. It’s been done, but not all that often, and we’re not good at it.

Comments (13) + TrackBacks (0) | Category: In Silico | Pharma 101

August 30, 2007

Elbow Room

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Posted by Derek

I hope that in decades to come that our current drugs look as crude as I think they will. For all of our knowledge and all our equipment, we still don't have much of an idea of what we're doing around this industry, not compared to the sum of what there is to know.

Most of our drugs (by "most", I mean way over 95%) bind to proteins. And that's fine, as far as it goes, because proteins sure are important things. We love them because many of them have pockets and cavities that fit small molecules, of course, giving us a tremendous leg up. But it's not that we've figured out how to attack them reliably, though, when you consider that there are many entire classes that have never been successfully targeted (phosphatases, to pick an outstanding example).

Once you get out of the small-molecule-binding zone, you're out in the wild, wide open prairie of protein-protein interactions. So far, we can't really affect those with small molecules, not worth squat. It's a shame, because the number of potential targets goes up by orders of magnitude when you take these interactions into account - well, assuming that we figure out what these zillions of interactions are actually doing, which is quite another problem in itself. But they're doing something, that's for sure, and we'd love to be able to step in for our own purposes.

But protein-protein interactions are only the beginning. If you want to go upstream and alter protein production at the source, then you're going to be targeting protein-DNA and protein-RNA) interactions. The list of known drug-like molecules which can do that is pretty short, and the success rate has been pretty small (more on the reasons for that in another post). And this is another area where only small regions of interaction space have been mapped out and understood, so there's room to work in - if you can find a way to make things work.

Don't stop there, though. We really don't pay enough attention to carbohydrates in all their forms, but they've got some crucial roles, too. Contacts involving complex polysaccharides are key to immune function, and small molecules that can affect them are rare indeed. A whole landscape of inflammation targets is waiting for someone who can get a handle on this stuff. And I haven't even talked about lipids, because frankly, we don't understand a lot of what they're doing. Protein-lipid interactions have been targeted, but can be a hard row to hoe, since the small molecules that work tend to look awfully greasy themselves. But there may also be lipid-lipid interactions that no one has ever noticed, and how you'd target those therapeutically is a real stumper.

There are even more exotic combinations, but you get the idea. When you look at the whole medicinally active universe, it's clear that we've only done successful work in a few small parts of it. An interesting and rewarding time awaits those who can extend those holdings. . .

Comments (17) + TrackBacks (0) | Category: Drug Development | Pharma 101

August 28, 2007

Like Clockwork

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Posted by Derek

There are a lot of drug development issues that people outside the field (and beginning medicinal chemists) don't think about. A significant one that sounds trivial is how often your wonder drug is going to be taken.

Once a day is the standard, and it's generally what we shoot for unless there's some reason to associate the drug with meals, sleep/wake cycles, or the like. People can remember to take something once a day - well, they remember it better than most of the other dosing schedules, anyway. That's why you actually want your compounds to be metabolized and cleared - everything has to be ready for the next dose tomorrow.

If your compound has a long half-life in the body after dosing, you'll step on the tail end of the last dose and you can see gradual accumulation of the drug in plasma or other tissues. And that's almost always a bad thing, because eventually every drug in the world is going to do something that you don't want. All you have to do is get the concentration up too high for too long (and figuring out what's too high and what's too long is the one-sentence job description of a toxicologist). If you stairstep your way up with accumulating doses, you'll get there in the end.

Ah, you might say, then just take the drug every other day. Simple! Sorry. Every other day (or every three, or four) is a complete nightmare for patient compliance. People lose track, and doctors know it. You'd better have a really compelling reason to go ahead with a weird regiment like that, and if you do, someone's going to seize the chance to come into your market with a once-a-day as soon as they can find one. (The exceptions to this are drugs given in a clinic, like many courses of chemotherapy - but in those cases, someone else is keeping track).

How about more often than once a day (q.d., in the Latin lingo). Well, twice a day (b.i.d. can work if it's morning/night. Three times a day can go with meals, presumably, but people are going to get tired of seeing your pills. More than three times a day? There'd better be a reason, and it had better be good.

So don't be scared as you watch your compounds disappear after giving them to the animals. You want that. Just not too quickly, and not too slowly, either.

Comments (19) + TrackBacks (0) | Category: Drug Development | Pharma 101 | Pharmacokinetics

July 15, 2007

Proteomics 101

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Posted by Derek

Over at the entertaining culture-blog 2Blowhards, the comments to this post (on people who feel deficient in math ability) include a mention of proteomics, which prompted Michael Blowhard to say:

"Proteomics" -- even the word is scary. I wonder how people in the field are going to communicate the substance and importance of what they're up to to civilians ... A challenge, I guess."

A challenge that I'm willing to take up! It's not my exact field, of course, but close enough. I'm starting a new category for posts like this, when I (and the readership here, in the comments) try to explain some technical buzzword-laden area in language that intelligent non-scientists can profit from. So. . .proteomics.

The place to start, most likely, is where the word came from. It's a direct steal from "genomics", the study of genomes, which are the total DNA sequences of a species (or individuals of a species). Back a few years ago when the human genome was being sequenced for the first time (all the individual A T C G letters being read off), it became clear that the number of genes that humans carry around was very much on the low side of what most people expected. (The human genome, as we have it today, is a composite - the number of people in the world who have their complete genome read can be counted on one hand. That's going to change drastically in the years to come as the process gets cheaper, faster, and more useful).

The reason why people expected more genes relates to what a gene is: a stretch of DNA that's read off (transcribed) and turned into a specific protein. That's DNA's job; it's a set of coded instructions to make proteins. But, as it happens, we have a lot more different proteins than we have genes. Clearly, something more happens downstream of the DNA part of the process.

A lot of things happen, actually. Those first-made proteins get altered in all sorts of ways. The same protein can be folded into different shapes, for starters (we're just now recognizing how important a process this is in some diseases). Proteins can also be clipped into smaller ones by many different routes, and at any stage they'll be decorated with molecular tinsel like sugars and lipids and phosphates. All of those can totally change a protein's function. This gives you some idea of where all that diversity is coming from - and why sequencing the human genome, huge and necessary accomplishment though it was, was nowhere near the end of the story.

Proteins spend their time interacting with other proteins. If you think of a cell in your body as a large irregularly shaped bag, full of intricate (and somewhat squishy) 3-D jigsaw pieces which are constantly sluicing around assembling or sliding past each other, you'll have a pretty reasonable idea of what it's like in there. Any given cell will contain thousands upon thousands of different proteins, many of which are doing multiple jobs depending on the time and place. Proteomics is the attempt to understand which proteins are doing what, when, with whom, and why.

It hardly needs saying, but we're just at the very beginning of that study. We have some tools to track these interactions, and they're far better than anything people had twenty or thirty years ago, but they're still rather crude compared to what we need. Huge signaling networks get uncovered and extended, and are found to touch upon others for reasons that are unclear. All sorts of feedback loops and backup systems are sketched in, and many pathways have been missed (or, alternatively, assigned too much importance) because they only operate under certain special conditions that our assays may overemphasize or skip entirely.

This project is much harder than the deciphering of the genome, and will take much longer. But that's because it's much closer to the real-time workings of a living organism, which means that comprehension, when it comes, will be still more valuable. Really substantial sums are being spent on this stuff, along with serious brainpower and computing resources. Progress will be jerky, irregular, infuriating, and of very great interest indeed.

Comments (20) + TrackBacks (0) | Category: Pharma 101