By next summer, more than 40% of
Streptococcus pneumoniae strains in the United States will
resist both penicillin and erythromycin, according to a recent prediction from the Harvard
School of Public Health. The forecast, based on mathematical modeling, was published in the
spring of 2003. It's too early to tell whether that prediction is precisely on track,
according to the senior author on that paper, Marc Lipsitch. But no one doubts that
multidrug resistance in this common bug—responsible for diseases that range from sinus
trouble and ear infections to meningitis and pneumonia—is speeding up.
It is the certain fate of all antibacterials to be fought off eventually by the
pathogens they target. The fact that the process is accelerating has been alarming public
health officials for some time, especially in the United States. We need new ways to defeat
disease, and we will need them forever.
Tried and True—and Tired?
Antibiotics have traditionally been plucked from nature's battleground. For billions of
years, tiny organisms have engaged in an arms race, hurling toxic molecules at each other
in the struggle to prosper. Nearly all of today's antibiotics are versions of weapons long
wielded by microbes and fungi. Chemical synthesis of entirely human-created antibiotics has
so far yielded only fluoroquinolones, a group of broad-spectrum antibiotics that includes
Cipro, which became famously scarce during the 2001 anthrax scare, and linezolid
(trade-named Zyvox), which is effective against some resistant strains of
Staphylococcus ,
Streptococcus , and
Enterococcus .
The usual way to find a new antibiotic has been laborious screening of immense libraries
of compounds, natural and otherwise. Some argue that screening chemical libraries is
approaching a deadend. There may be diminishing returns from screening, but it's not quite
dead yet: in October, researchers at the University of Wisconsin at Madison reported a new
class of bacterial RNA polymerase inhibitors with antibiotic potential. They were found by
screening for molecules that prevent
Escherichia coli from transcribing RNA.
Christopher T. Walsh of Harvard Medical School says screening's problem may be simply
that libraries aren't good enough. Marine organisms have not been studied well, he points
out, and 90% of organisms in the biosphere can't be cultured in standard ways. He says,
“We're missing 90% of them every time we go and look in nature.”
Walsh is doing his bit to create new libraries. He and his colleagues have recently
employed combinatorial biosynthesis to learn how to use part of the machinery for
assembling cyclic peptide antibiotics to control their architecture. The result was a small
library of natural product analogs, some of which have improved antibiotic activity against
common bacterial pathogens. “There are dozens of such enzymatic domains that in principle
one could clone, express, and test with other substrates. I view that as the kind of thing
we should do,” he says. For example, Walsh suggests, it is a reasonable approach to
second-generation improvement of daptomycin, the antibiotic most recently approved for sale
in the United States.
Improving on Nature
Walsh collaborates with Chaitan Khosla of Stanford University on finding ways to make
existing antibiotics better. They are studying biosynthesis of rifamycin, an antibiotic
that is increasingly less effective against its prime target, tuberculosis (TB) (see Figure
1). “In the course of learning about that pathway, we've learned a few interesting things
lately about how that molecule is initiated, and we're trying to apply it in other
contexts, especially in the context of erythromycin biosynthesis,” Khosla says. The idea
would be to make a molecule that might be more effective against bacteria that are becoming
resistant to rifamycin—and are already naturally resistant to molecules like
erythromycin.
“Basically, what we do is to try and figure out new ways to hijack the biosynthesis of
antibiotics in nature so as to modify their structures with the goal of improving them,”
Khosla explains. He works with an important class of natural antibiotics called polyketides
that have generated dozens of drugs, including erythromycin.
Polyketides are secondary metabolites (which give their producers a competitive
advantage in their environment) produced mostly by bacteria and fungi and made by a complex
and structurally diverse family of enzymes called polyketide synthases (see the primer by
David Hopwood in this issue of
PLoS Biology ). Among them are the anthracyclines, a group of anticancer
drugs and antibacterials that includes tetracycline. In this issue of
PloS Biology , Khosla and his colleagues report that they can make
selective positional modifications in existing anthracycline antibiotics by starting in a
different way with a different starting molecule. The molecule came from a natural
anthracycline antibiotic, an estrogen receptor antagonist called R1128. R1128 is made via
two modules of enzymes that work sequentially; the first module starts the process, and the
second completes it. This division of labor permitted the researchers to tack the first
R1128 module onto two other enzyme systems, thus engineering completely new anthracyclines.
Some were more active in two types of assays than the natural parent molecule. “One setting
was an assay on an estrogen-sensitive cancer cell line. Another setting was an assay to
probe activity of an enzyme that's of particular interest in Type 2 diabetes, called
glucose-6-phosphate translocase.” The work also revealed fundamental mechanistic features
of the polyketide synthases, Khosla says.
The researchers didn't study the new anthracyclines' effects on bacteria, but Khosla
notes that the general principle should apply to other classes of compounds, although the
details of how it's implemented will vary from system to system. He says, “The upshot of
this paper is that it is now possible to modify a particular methyl group in just about any
anthracycline antibiotic.”
Finding New Targets
Instead of searching for new antibiotics by modifying existing ones, some researchers
are trying something completely different—first finding the most vulnerable targets in a
bacterium and then designing something that hits one or more of them hard. “You have to
understand a helluva lot more about how these little cells work. In fact, we think we
understand a lot, but I think we can understand almost everything now that we have all the
genomes,” says Lucy Shapiro of Stanford University School of Medicine. While having full
genome sequences—more than 100 microbe sequences have been completed—is essential, Shapiro
believes that knocking outs genes galore to find out which ones are necessary and going
after them all is not a sensible strategy. She observes, “People have been doing that for a
while with absolutely no success. That's really going after the problem with a Howitzer
instead of with an intelligent approach.”
So instead of screening libraries of existing compounds, Shapiro prefers using
structural information about drug targets or their natural ligands to create new drugs, an
approach known as rational drug design. And instead of looking at all essential genes in a
bacterium and choosing one to target, she and her colleagues look at genetic circuitry that
controls the cell cycle, the pathway that coordinates cell growth and differentiation. They
have identified key control points, or nodes, in the circuitry for their favorite study
subject,
Caulobacter crescentus . Thus, they have found critical genes
encoding proteins that control several critical functions in the cell. Their first
candidate was an essential enzyme, a methyltransferase called CcrM, that prevents a
particular piece of DNA from being expressed in a cell by tagging it with a methyl
group.
Antibiotic discovery is all chemistry, Shapiro says, which is why she joined with
biochemist Stephen J. Benkovic of Pennsylvania State University. They didn't know the
structure of CcrM, Benkovic explains, but the literature about other methyltransferases
suggested that the adenine molecule, which is the substrate for CcrM within DNA, binds to a
specific region of the enzyme.
The researchers designed adenine-like molecules that would bind to CcrM and then
developed inhibitors. Benkovic says, “We already knew what kind of structure we wanted, and
we simply fine-tuned it.” They worked their way through 1,000 inhibitor candidates, ending
up with a small subset—no more than about 20—that not only inhibited CcrM, but also killed
Caulobacter very quickly.
And not only inoffensive
Caulobacter . The compounds knock out other gram-negative
bacteria, such as the pathogens
Brucella abortus and
Francisella tularensis . Some even killed off anthrax, a big
surprise because it is gram-positive and so has much thicker cell walls than gram-negative
bacteria. The researchers undertook an exhaustive series of experiments to identify which
gram-positive bacteria would be affected by which compounds. The list of sensitive
pathogens now includes multidrug-resistant
Streptococcus ,
Staphylococcus , and
Mycobacterium tuberculosis .
More recently, Shapiro reports, they have demonstrated efficacy against rats infected
with anthrax or multidrug-resistant
Staph , although the compounds save only about 60% of the rats
at present. She notes, “So we have a long way to go. But this has proven that if you go
after something using some rational approach instead of hit-and-miss, you'll probably have
more success than by the other method.”
Benkovic points out that theirs is an entirely new class of compounds, small molecular
weight compounds that can be made in a few steps. He says, “They don't look like the normal
antibiotic, so that's why I think they're fairly unique.” The basic research was done under
a grant from the Defense Advanced Research Projects Agency (DARPA), the United States
Department of Defense's (DOD) central research and development organization, and once the
researchers realized they wanted to develop drugs against three agents that have been
considered bioterrorism threats —
Brucella , tularensis, and anthrax — they established a separate
operation, Anacor Pharmaceuticals, which is developing them with DOD funding and without
Shapiro. In her Stanford lab, she continues her fundamental research to define the complete
genetic circuitry of
Caulobacter , hoping to identify additional nodes in the
circuit. She says, “I am not doing it to develop antibiotics; that's what comes out of the
work. My goal is to understand how the cell works. I think a lot of studies in pathogenesis
should not be just to understand pathogenic organisms, but to understand the complete
network of regulatory mechanisms that controls the bacterial cell.”
Phage Therapy
The most radical approach to new antibiotics may be the resurrection of an old idea:
bacteriophage therapy (see Figure 2). Late in the 19th century, a researcher noticed that
water from some of India's sacred rivers combated cholera. Some years later, the active
agents were identified as viruses that infected bacteria. Such viruses are called
bacteriophage, or phage for short. There were reports of phage success against dysentery,
typhoid, and plague, and bacteriophage therapy had a brief heyday, especially in the 1920s.
Results on other diseases were mixed, and with the appearance of antibiotics, phage therapy
became unfashionable in the United States, although it has continued in Russia and Eastern
Europe.
Phage were the model organisms of choice for genetics research in the 1930s and 1940s,
but became less fashionable as research tools when investigators moved on to eukaryotes. A
few held on, like Ry Young of Texas A&M University, who has made phage-induced cell
lysis his life's work. “The cell is basically genetically dead as soon as the phage goes in
there, but it will keep living as sort of an infected zombie for as long as the phage wants
it to, with virus particles accumulating inside the cell,” he explains. “Only when the
phage is ready and has decided that it's the right time will it pull the trigger. And the
cell blows up.” The freed phage then spew forth to infect new cells.
Antibiotic resistance has led to new interest in phage therapy by several small biotech
companies. Young continues basic research at Texas A&M, but has also joined one of
them, GangaGen, providing bacteriophage expertise to its labs.
Phage do kill pathogenic bacteria effectively, and they do it without penetrating human
cells, which they can't even recognize. So what is keeping phage therapy out of the clinic?
Problems that some doubt can be overcome.
Because bacteria develop resistance to phage rapidly, phage therapy companies will need
to direct cocktails against a single pathogen, according to Vincent Fischetti at The
Rockefeller University. Phage are also antigenic, and the antibodies they stimulate will
neutralize their effects during subsequent treatment, he says. But the chief problem
appears to be regulatory—regulatory in the political, rather than the genetic, sense. When
bacteriophage package their DNA, they occasionally include varying amounts of their hosts'
DNA, too. This miscellany, Fischetti points out, is likely to make the Food and Drug
Administration unhappy. “Phage normally are very fragile, their tails break, so lot-to-lot
homogeneity could be a problem too,” he adds. “So even though it will work, I think they'll
have an uphill battle.” Phage may well enter agricultural or veterinary use, he predicts,
but are probably not going to be available to patients in the United States any time
soon.
Fischetti chose a different approach to phage therapy. It does not rely on phage
themselves, but on enzymes that phage produce to smash their way out of their host bacteria
so they can infect new hosts. He and his colleagues employ these enzymes externally to kill
bacteria. He reports, “We now have enzymes that will kill
Strep pyogenes , pneumococci,
Strep pneumoniae ,
Bacillus anthracis ,
Enterococcus faecalis , and group B
Strep . The beauty of these enzymes is that they are targeted
killing. You only kill the organism you intend to kill, without destroying or affecting the
surrounding organisms that are necessary for health.”
The enzymes can be loaded into a nasal spray that wipes out pathogens such as
Pneumococcus ,
Staphylococcus , and group A
Strep on contact with mucous membranes. The strategy might
prevent bacterial infections from spreading in close quarters like hospitals, nursing
homes, and daycare centers. Fischetti says, “Clinical trials would tell us how often we had
to treat, but more important, we'd have a reagent that could treat people who walk out the
door of the hospital to eliminate or reduce the transmission of resistant organisms into
the community. We don't have that capability right now.”
Fischetti and his colleagues have moved on to using the enzymes systemically to wipe out
Bacillus anthracis spores, preventing them from germinating and
seething through the bloodstream, producing deadly toxins. An IV drip would be started
after exposure to the spores. The method, Fischetti reports, is already successful in mice;
clinical trials will determine how long treatment must be continued, perhaps a week or so.
They have also eliminated septicemia from pneumocci with the same intravenous method.
Up to now the enzymes must make contact with bacteria to kill, but Fischetti is hoping
that a new generation of engineered enzymes will be able to kill pathogens inside cells
too. A second disadvantage is that they are effective only against gram-positive bacteria,
although that group includes many vicious pathogens.
But phage enzymes seem to offer one very big advantage: resistance to them has yet to
develop. Fischetti says, “We've tried very hard to identify resistant bacteria, but so far
we haven't found resistant organisms in all three of the enzymes we're working with. It
appears to be a very rare event, much rarer than resistance to antibiotics.” Fischetti
cautions against expecting that gladsome state to last forever, but he points out that even
if widespread resistance takes the same 40 or 50 years that antibiotics required to become
significantly resistant, phage enzymes could buy researchers decades for inventing other
approaches.
Antibiotics in the 21st Century
There is no shortage of ideas for unearthing new antibiotic candidates. Why are they so
slow to enter medical practice? The bottleneck, researchers agree, lies in the development
process of turning them into effective therapies. Several researchers blame the big
pharmaceutical companies that got so big by leading the way to new drugs for battling
infectious disease, but in recent years have dropped out. Fischetti complains, “These are
the big companies that have the money to develop antiinfectives, but they leave it to small
biotech companies, and it's not going to happen as rapidly as it should. I think it's
really unconscionable for these big companies to drop the ball because it's not going to be
a billion-dollar market for them and that's what they're looking for.”
Half a billion at least, says Francis Tally, a big pharmaceuticals veteran who is now
chief scientific officer at Cubist Pharmaceuticals, a biotech company located in Lexington,
Massachusetts. According to Tally, Cubist produced daptomycin, approved in September 2003,
by licensing it from Eli Lilly, which shelved the new compound after concluding its
potential market was only $250 million.
But, Tally argues, the size of the market is not the only barrier to new antibiotics.
Combinatorial chemistry and the genomics revolution have simply not delivered on their
early promise. “The pipeline is very dry,” he says. “There's been a real lag at the basic
research level.”
“Antibiotic discovery is hard,” Shapiro says. “It's a huge long process to get a decent
antibiotic.” Walsh agrees. “It's easier to find inhibitors of particular enzymes for
particular processes—and a very long road to convert that into something for
development.”
In the meantime, there is a rising clamor to slow down the rate at which bacteria
develop resistance. Doctors are exhorted to cut back on prescribing antibiotics and decline
to prescribe for viral diseases, which antibiotics can't combat, even when their patients
badger them.
But even if antibiotic consumption slowed, we will still need new antibiotics. “I always
say it's not a matter of if, it's only a matter of when,” says Walsh. “There will always be
a need for new antibiotics because the clock starts ticking on the useful lifetime of any
antibiotic once you start to use it. That cannot be argued.”
