I sent off a manuscript to a chemical journal not long ago. There's an initial flurry of e-mail activity when you do that - we've received your manuscript, we've sent your manuscript out to reviewers - and then a more or less prolonged period of silence. The next thing you hear is whether the paper's been accepted or not, along with the referee comments.
Mine were the usual mix of helpful suggestions and things that make you roll your eyes. One of the latter was a comment that immediately pegged the reviewer as someone from academia. They noticed that the data from our primary assay, against a human enzyme, didn't always match up well with the secondary assay, which was against a rodent cell line, and wanted some more explanation for why some groups of compounds weren't active.
To which I could only reply "You and me both!" That's a constant problem in medicinal chemistry. A majority of projects are set up in that format, with a cell-free assay as the first filter, then cells expressing the target as the next hurdle. And it's just about inevitable that there will be whole groups of compounds that work fine in the first assay, but just wipe out in the second one.
Why should that be? As far as we know, there are two general ways that compounds can get into cells: passive transport and active transport. The passive route is just diffusion across the cell membrane: "Wonder drug? You're soaking in it!" It's affected by broad trends in molecular size, polarity, and so on. The second route is when your compound hitches a ride on some transport protein.
There are hundreds of these things involved in opening up channels into and out of the cell. Some of the famous ones move ions (calcium, potassium and the like), which makes sense. Those are small and electrically charged, so they're not going to just wander across the membrane on their own, and the cellular machinery depends on keeping such membrane potentials tightly controlled. Then there are transporters for large proteins, which are too huge to diffuse by themselves, and for essential classes of small molecules like fatty acids.
No one's sure how many of these things exist. Just in the last few years, there's been a whole new class discovered, the aquaporins, which (as the name implies) move water itself across the cell membrane. You wouldn't think that you'd need an active transport system for that (at least a lot of people didn't think so) but the things turn out to be ubiquitous. If there's a transporter for water, there can be one for anything.
The efflux pumps I spoke of the other day in antibiotic resistance are active transport proteins, too, naturally. Those complicate things by taking compounds that diffuse perfectly nicely into cells and making them look like they're bouncing off a layer of armor plate instead. You'll also get that effect when your standard project compounds ride in on some transport system, then you make some small structural change which causes your drug to lose its train ticket.
It's a lot of work to figure out what's going on, and often you can't get a handle on it, anyway. Many of these transport systems don't have specific inhibitors, so it's not like you can switch them off one by one to see which one is the problem. If you have a good way to monitor your compound on a cellular level (like a fluorescent probe), you can actually see the things going in and being pumped back out sometimes, or you can see if the transport system can be saturated as you load up on drug. But there's no way you can do this for hundreds of drug candidates on every project.
So, it's just one of those things. I'm on a project right now that has the same thing going on. We make tiny changes to our molecules, and the cell activity suddenly gets a hundred times better, or a thousand times worse. But are these trends going to translate to the cells inside a real animal? And if they do, will they be relevant to the active transport systems in humans? Bite your tongue.
