How to reel in high-throughput results using worms and fish.
In the past few years, improvements in imaging and automation techniques have
made it easy for researchers to see hundreds of plates of cells partake in every
activity from differentiation to apoptosis. But in living and breathing animals,
we’re only just beginning to realize the potential of large-scale screens.
“To take a whole animal and be able to perform high-throughput
screening—that’s a new thing,” says Alan Mayer,
assistant professor of pediatrics and cell biology at the Medical College of
Wisconsin in Milwaukee. That’s worth the effort, he says, because you can
do tests that you can’t do in cell culture, such as determining how
changes in one gene can affect a living organism. What’s more, Mayer says,
the techniques are a boon to animal physiology studies; a cell assay can tell you
whether a drug is toxic, but only testing the drug in a living organism will show
you how that drug is metabolized. Organisms such as Caenorhabditis
elegans and zebrafish are a popular choice for such in vivo studies, because
they are easy to raise and manipulate and they share molecular pathways with those
of humans. Others, such as planarian flatworms, frog larvae, and Drosophila
melanogaster, offer their own distinct advantages.
But working in vivo raises its own challenges. First, establishing these
organisms as models for human disease isn’t always straightforward. Then,
doing in vivo studies in hundreds of animals at once rather than one by one entails
introducing automated components as well as making adjustments in everything from
the volume and chemical make-up of the organisms’ surroundings to the
brightness of the light in their living space. You’ve got to keep them
alive, and non-stressed, during the entire process.
The Scientist talked to some researchers who have blazed new
paths in whole-animal screens. Here’s what we found:
The microfluidic worm sorter uses suction to immobilize worms against a
flexible membrane, allowing researchers to automate their nerve regeneration screen.
The Yanik lab / MIT
User: Mehmet Fatih Yanik, assistant professor of electrical engineering
at the Massachusetts Institute of Technology, Cambridge
Project: Chemical and genetic screens of nerve regeneration in C.
elegans
Problem: Several years ago, Yanik developed a way to sever nerves in C. elegans using ultrashort laser pulses. The surgery, done
manually, took 10 minutes per worm. “I was sick and tired of doing it
manually,” Yanik says. What’s more, such surgery, and
high-resolution imaging to trace regenerating neurons, required using drugs to
anesthetize the animals, a process that isn’t conducive to subsequent
genetic and chemical screens since the anesthetic could alter gene expression or
the chemical environment. He needed a way to eliminate the need for anesthesia
and automate the surgery.
Solution: After months of brainstorming and experimenting, Yanik and his
students came up with a way to use a microfluidic chip to sort, isolate, and
immobilize individual worms. In 2007, they had a prototype of the chip, which
consists of a series of channels and suction devices that direct the flow of
worms, capturing individual worms and placing them into tiny chambers
(Proc Natl Acad Sci, 104:13891–95, 2007).
The crucial component is the tiny pump, which is used to briefly suck
living worms against a flexible membrane. This immobilizes the worm for a few
seconds, eliminating the need for anesthesia while allowing Yanik’s
team to take a high-resolution photo and cut select neuronal projections at a
precise distance from the cell body, which acts as a reference point.
“The first question was, ‘Are the zebrafish going to produce
enough acid for us to measure?’”
Finally, the pressure exerted by the pump is released, so the worm can
flow back into its chamber. There, they can treat the animal with chemicals and
perform the imaging process a few hours later to see how the nerve is regrowing.
“Once you can stop the animal, a whole world of opportunity
opens up,” he says. “Now we can screen tens of thousands,
looking for molecules that can enhance regeneration.”
Considerations: The technology is not commercially available, and the few
labs successful in creating similar set-ups have been armed with engineers,
Yanik says. Many have contacted his lab and asked for help. “The thing
that will make [the device] commercial is the demand on the user end,”
he says.
A 96-hour-old zebrafish embryo in a single well of a 96-well plate.
Alan Mayer
User: Alan Mayer, assistant professor of pediatrics and cell biology,
Medical College of Wisconsin, Milwaukee
Project: Assessing genetic programs of zebrafish metabolism during
development
Problem: For decades, researchers have studied metabolism by placing
small mammals into shoe box–sized chambers to measure oxygen consumption and
carbon dioxide production. Mayer's group wanted to find a high-throughput way to
assess metabolism in order to screen for genetic mutants that alter metabolic
rate or drugs that can accelerate metabolism.
Zebrafish embryos are good candidates for such screens: They can be
confined to fixed volumes, and ambient liquid can be analyzed for carbon
dioxide. But the tests can only be conducted on a few embryos at a time, using
single syringes to add and collect fluid.
Solution: About a year ago, Mayer's team began trying to scale up a
zebrafish metabolic assay from one that uses 1 to 10 fish to one that uses a
96-well plate. They decided to try measuring the acidity of the solution, which
would provide an indirect measure of carbon dioxide. "The first question was,
'Are the zebrafish going to produce enough acid for us to measure?'"
They had to settle on a volume that the embryo was confined in—small
enough for the acid concentration to affect the ambient pH, but large enough in
volume to physically accommodate the embryo. Then, they chose a simple indicator
dye—phenol red—as a pH readout.
Considerations: You might not need special equipment for an assay like
this, but you will need to think carefully about the experimental parameters.
For example, Mayer had to tweak the buffer solution so that the production of
acid could be measured over a reasonable amount of time. "Something less than a
day but more than a couple of minutes, because it takes time to set the assay
up," Mayer says. "You don't want the first embryos to be finished before the
last embryos are in [the wells]," or you won't be able to monitor progression of
the assay color change (i.e., CO2 production).
Infection of Candida albicans in the C. elegans intestine progresses as the worm’s cells are replaced by
metabolically active yeast cells (red).
Eleftherios Mylonakis and PLoS Pathogens
User: Eleftherios Mylonakis, assistant professor of medicine, Harvard
Medical School, Boston
Project: Infecting C. elegans with fungal species and
screening for potential antifungal agents
Problem: Many antifungal compounds can be toxic to mammalian cells. In
vitro assays, however, measure potential efficacy of the antifungal agents,
leaving toxicity for later stage trials. Mylonakis’s group needed to
develop a whole-animal screen that would allow them to easily pinpoint safe
antifungal agents.
Solution: The group can infect large numbers of worms, contained in
384-well plates, with a fungus and track the infections using microscope images.
“The rest is easy from there,“ Mylonakis says.
“Just add screening compounds and monitor the survival of the worm and
the expression of fungal virulence.”
Sounds simple, but coming up with the technique was a real challenge.
When Mylonakis first started developing the technique a decade ago, he had
minimal knowledge of how worms respond to fungal pathogens. He had to first
demonstrate that the worms would ingest and become infected by the same species
of fungus that serve as pathogens in humans. “When we started our work
it was not even known if fungi can be a food source for C.
elegans,” he recalls.
They were also able to make sure that the innate immune system molecules
that detected such fungal pathogens were similar in humans and worms, in the
case of a common, yeast-like fungus Cryptococcus neoformans (J Exp Med, 206:637–53, 2009).
Considerations: The most common mistake for beginners, says Mylonakis, is
not adjusting the assay to fit the specific endpoints, whether it’s
the effects of the antifungal compound or the worm’s immune response
to the fungus. If it’s the latter, for example, then the assay should
be adjusted to a fit a 4- to 6-hour window—the peak of a
worm’s immune response. The former might take longer to see.
Genes involved in appetite, which researchers tagged with GPF, are
activated in hungry worms during metabolism studies.
Eyleen O’Rourke
User: Eyleen O’Rourke, research fellow in the lab of Gary
Ruvkun, Harvard Medical School, Boston
Project: Developing automated screens of C. elegans to
identify genes controlling appetite
Problem: O’Rourke and her colleagues designed a worm that would
glow green by activating certain hunger genes linked to a green fluorescent
protein (GFP) when the animals were out of food. The worms could potentially
provide great high-throughput data, but the researchers needed to develop a
quantitative way to score brightness. Without that, says O’Rourke,
“there was no statistical analysis to be done.”
Solution: The first challenge involved acquiring the images.
Collaborating with two other labs, O’Rourke began by taking
high-resolution photos of 20 worms at a time for analysis. At lower
magnification, they could see more worms in the field, but they lost
sensitivity. To help overcome that, they outfitted their microscopes with a more
powerful ultraviolet light source, and scaled down GFP expression to reduce
background noise in their samples.
The brighter UV had an unwanted effect, though: When they switched it on,
the worms would instantly flee to the periphery of the wells. “We
didn’t expect to lose all of them within a few
milliseconds,” she says. So they redesigned the system to maintain the
worms under normal bulb light, programming the microscope to find the correct
focal plane before switching quickly to UV and snapping an image.
Considerations: For quantifying brightness, software advances have not
yet caught up with the hardware advances made by O’Rourke and her
colleagues. Currently, the group uses a freely available cell-imaging software,
called CellProfiler, to analyze fluorescence in the worms.
“It’s a very powerful and great tool, but worms have many
specific issues, like they can overlap on top of each other” in the
wells, she says. They are working with CellProfiler’s creator to
develop software tailored specifically to worms.
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