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Orange light bathing brain
Neurophysiology
charges ahead
At first glance, animal nervous systems seem to defy understanding. Even a “simple” animal such as a fly has over
100,000 neurons that can fire in billions of possible combinations, driving precise, nuanced responses to complex environmental stimuli. Nonetheless, evolution has given brains a
few features that make them tantalizingly easy to study—at
least in theory. By Alan Dove
N
eurons primarily operate electrically, a fact neuroscientists have exploited for decades. Using tiny
wire electrodes or capillary tubes, researchers can
measure current and voltage changes in isolated
cells or living animals. Simple electronic hardware can then
amplify and record the signals. The brain is also relatively
isolated from the body and composed of cell types found
nowhere else. That means investigators can genetically engineer animals’ nervous systems with exquisite precision.
Though both the electrophysiological and genetic approaches to neuroscience have been popular for years,
recent advances in both fields are now spawning a new generation of techniques. The developments range from gradual
but persistent progress in electrode design to automated
surgery systems and novel genetic engineering strategies.
Taken together, the new methods may soon let many more
researchers incorporate advanced neurophysiology analyses
into their work.
Getting wired
The most common way to measure electrical changes in
neurons is to stick metal electrodes into brain slices in a petri
dish, or directly into the brains of living animals, methods
investigators have used for over 50 years. For nearly as long,
researchers like David Farb, professor of pharmacology
at Boston University School of Medicine in Boston,
Massachusetts, have steadily expanded the capabilities of
these methods. “When I first started in electrophysiology we
used vacuum tube amplifiers,” says Farb. At that time, he
explains that “an investigator might spend a day and get a
recording from one cell, and maybe the next day nothing
worked, so it was ... very slow.”
After a long series of refinements, Farb and his colleagues
can now measure activity in entire brain regions while
an animal engages in natural behaviors. The work is still
tedious, though.
For each experiment, Farb’s lab builds a multilayered
apparatus. First, they bundle four electrodes into a device
they call a “tetrode,” which is about the diameter of a human
hair. Companies such as Thomas Recording in Giessen,
Germany and Tucker-Davis Technologies in Alachua,
Florida sell prebuilt tetrodes and related equipment, but
researchers without the resources to purchase this type of
equipment often build their own. Farb explains that he has
a small shop full of undergraduate students building the
single-use tetrodes continuously, “because we can’t afford
to buy them.”
Whether they buy or build the tetrodes, though, using
them requires patience and sophisticated surgical skill.
Two to three dozen tetrodes attach to a top section called a
“head stage,” which holds a set of microdrives, mounts that
hold and position the tetrodes. The microdrives contain tiny
screws that the researcher advances a short distance each
day, to avoid damaging the brain tissue. The process slowly
moves the tetrodes into the desired brain region of a rat or
other animal.
For those who can work with them, though, the
multiplexed electrodes can yield troves of data. Farb’s lab
has traced networks of “place cells” to show how animals
navigate in different environments, findings that have helped
illuminate everything from the fundamental mechanisms of
learning to the pathogenesis of Alzheimer’s disease.
Unfortunately, electrode analyses in conscious animals
have been traditionally restricted to nonhuman primates and
occasionally rats, as mice were too small to accommodate
the field’s standard equipment. Joshua Dudman, a group
leader at the Howard Hughes Medical Institute’s (HHMI)
Janelia Research Campus in Ashburn, Virginia, wanted
to change that. “I actually went and talked to my late
grandfather, who was a mechanical engineer who designed
carburetors, [and] we kind of bounced some ideas off each
other” says Dudman.
Armed with a cocktail napkin drawing and some
additional advice from Robert Wurtz, a distinguished
scientist at the National Institutes of Health’s (NIH) National Eye Institute in Bethesda, Maryland who developed
similar systems for monkeys, Dudman began building a
standardized system for probing mouse brains. His team
used a 3D printer to create mounting brackets that hold
the electrodes and restrain the animal’s head. Working
with another group at Janelia Research continued>
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NEUROTECHNIQUES
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NEUROTECHNIQUES
Patch clamping uses a tiny microcapillary tube
to track fluctuations in a single neuron’s ion
channels.
Campus, they then refined an older design for silicon chipbased electrode arrays. The result was the rodent in vitro/
vivo electrophysiology targeting system (RIVETS). As the
name implies, RIVETS allows a single, standardized set of
components to be used for studies in live animals or brain
slices. It also permits simultaneous imaging with two-photon
microscopy.
Besides the 3D printed parts, RIVETS also uses off-theshelf equipment that most neurobiology labs should already
have. “One of the big issues is how do you integrate it with
all these other pieces of equipment. If we want to position
something precisely, we don’t really want to remake micromanipulators, since there are many excellent companies
making really high-end stuff,” says Dudman. A micromanipulator is a precisely machined device usually built to attach
to a microscope stage, with joysticks or handles that allow
a researcher to move tiny components such as electrodes
into specific positions.
Scientists can download the 3D printing files and other
information from Dudman’s website, and use micromanipulators from companies such as Scientifica in East
Sussex, United Kingdom, or Sutter Instrument in Novato,
California. Dudman adds that Scientifica recently bought a
nonexclusive license to sell packaged RIVETS systems for
researchers who prefer a commercially supported product.
Dudman isn’t the only one trying to build a better mouse
cap. “The main innovation that we made in my lab was to
make ultralight but still ultrastable versions of chronic electrophysiology implants,” says Christopher Moore, associate
professor of neuroscience at Brown University
in Providence, Rhode Island.
The light weight was crucial in allowing the team to study
mouse brains. “The revolution in genetic engineering and
in vivo systems in mammals has been a huge boon, but
it’s all come in mice, and mice are little guys,” says Moore.
He explains that with the new arrays, “now you can do the
physiology you used to only be able to do in rats and monkeys but in a prep that’s so much better tolerated by
the mice.”
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Whether used in vitro or in vivo, wire electrodes generally measure electrical changes among groups of neurons.
A complementary technique, patch clamping, uses a tiny
microcapillary tube to track fluctuations in a single neuron’s
ion channels.
For those new to the patch clamp technique, the first step
should be a review of basic electronics. “Doing patch clamp
work, ultimately you’re treating a neuron like an electrical
device, and understanding how electrical devices work and
how those principles apply within a biological context is, I
think, of the utmost importance,” says Michael Markham,
assistant professor of biology at the University of Oklahoma
in Norman, Oklahoma. Markham maintains a free software
package called “Electrophysiology of the Neuron” to aid
that process.
Among other electrical challenges, Markham points out that
the further a capillary goes into an animal’s brain, the higher
its inherent resistance and capacitance. That means deeper
probes have less bandwidth than shallower ones.
Inserting capillaries into a live animal’s brain for patch
clamp experiments also entails a tricky operation, requiring
exceptional care and surgical skill. Edward Boyden, leader
of the synthetic neurobiology group at the Massachusetts
Institute of Technology (MIT) Media Lab in Cambridge,
Massachusetts, decided to turn the job over to a robot. “We
discovered an algorithm that ... allows you to patch clamp
neurons, and also allows for automation in a computer. It
doesn’t require human intuition,” says Boyden.
Boyden’s group uses the algorithm to drive a robot, which
slowly inserts a patch clamp capillary into the desired region
of an animal’s brain until it detects an increase in electrical
resistance, indicating that the capillary has encountered a
neuron. The robot can then attach the capillary, and not only
measure ion channel activation in the neuron but also extract
cytoplasmic material for biochemical analysis.
After publishing the technique, the researchers also created
a company, Neuromatic Devices, which now sells automated patch clamp equipment to make it easier for others
to duplicate the technique. Boyden anticipates automating
other aspects of neurophysiology in live animals, too. “What
we think we’ve stumbled across is an area that one might call
‘in vivo robotics,’ where we could deploy a wide variety of
technologies to automate and make turnkey these kinds of
processes,” he says.
Meanwhile, Moore’s team is trying to make wire electrode
physiology more accessible. Jakob Voigts, a graduate student in Moore’s lab, built a collection of standard electrophysiology equipment available under an open source license. The
project, called Open Ephys, allows researchers to assemble
a sophisticated set of neurophysiology gear from a few thousand dollars’ worth of parts. An online support community
helps debug any problems that arise.
Bright ideas
Besides making it easier to measure neuronal activity,
Boyden also helped pioneer new ways to activate neurons
experimentally. The innovation grew out of his frustration
with traditional pharmacological and electrical stimulation
methods, neither of which is especially precise. “You couldn’t
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Featured Participants
Boston University
School of Medicine
www.bumc.bu.edu
Brown University
www.brown.edu
Central Michigan
University
www.cmich.edu
HHMI Janelia Research
Campus
www.janelia.org
Massachusetts Institute
of Technology
www.mit.edu
NIH National Eye
Institute
www.nei.nih.gov
Neuromatic Devices
www.neuromaticdevices.com
Open Ephys
www.open-ephys.org
Scientifica
www.scientifica.uk.com
Sutter Instrument
www.sutter.com
Thomas Recording
www.thomasrecording.com
Tucker Davis Technologies
www.tdt.com
University of Oklahoma
www.ou.edu
just activate a specific set of neurons,” says Boyden.
While searching the literature, he and his colleagues
found a tantalizing lead. “We were brainstorming in the
Spring of 2000, and we stumbled across some papers on
so-called microbial opsins,” Boyden explains. The team
was especially intrigued by photosensitive opsins, which
open ion channels in the microbial cell membrane in response to light. When the investigators genetically modified animal neurons to express one of these proteins, the
neurons started to fire in response to pulses of light. Since
then, Boyden’s group and others have steadily improved
the technology, called “optogenetics,” which is now a standard tool for probing neurophysiology in live animals as well
as brain slices.
The latest developments in optogenetics include channel
proteins sensitive to red light, which can penetrate deeper
into brain tissue, and a photosensitive chloride channel that
can inhibit neurons in response to light instead of stimulating them. Clever mouse genetic techniques also allow
scientists to restrict the photosensitive channels’ expression
to very specific brain regions. Combining the techniques,
researchers can now shine light through an intact mouse
skull and either activate or inhibit target neuronal populations with exquisite precision. “The tools have really started
to get very routine in usage,” says Boyden.
Optical tricks can extend the optogenetic possibilities
even further. For example, researchers can now project
multiphoton holograms into animal brains to activate single
neurons. “You can really try to dial in the complex threedimensional configuration of neural coding, and that would
then allow for very specific hypotheses of neural codes to
be tested,” Boyden explains.
Optogenetics also combines well with electrophysiology.
One major challenge in traditional wire-electrode placement
has been figuring out where the electrode is in the animal’s
brain. With optogenetics, investigators can now be certain.
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“You can tell pretty well when you flash the light whether the
neuron you’ve isolated with your metal electrode is [the right
type],” says Brown’s Moore.
Moore’s lab has even figured out how to combine
optogenetics with another favorite tool of neuroscientists: functional MRI(fMRI). By revealing changes in blood
flow throughout the brain, fMRI gives an overview of the
response to a stimulus. Optogenetics can now provide very
precise stimuli. “It allows you to stimulate a specific cell
type in a specific spot, let’s say in the hand representation
of the neocortex; then you can use fMRI to say, ‘Where else
in the brain is activated when I stimulate just that cell type in
just that spot?’” says Moore, adding, “That’s an amazingly
powerful thing to know.”
Despite its advantages, optogenetics still has some
drawbacks. For the most accurate activation of cells,
researchers still have to insert fiber optics into a mouse’s
brain through the skull. Ute Hochgeschwender, associate
professor of neuroscience at Central Michigan University
in Mount Pleasant, Michigan, is working on a different strategy. “A way to get around that would be if instead of using a
physical fiber, if you can use basically a light source which
is also biological,” says Hochgeschwender.
Hochgeschwender and her colleagues fused luciferase
proteins from fireflies with light-sensitive opsins and fluorescent protein domains. The resulting luminopsins open their
ion channels in response to light, emit light when provided
with the appropriate luciferase substrate, and glow when
examined under a fluorescent microscope. Researchers
can activate the targeted neurons either by shining light
on them or giving the animal an injection of the luciferase
substrate, while the fluorescent tag reveals which cells are
expressing the protein. “Any opsin, any optogenetic element
developed, we can give the additional dimension of being
chemically accessible,” says Hochgeschwender.
In the latest iteration of the technology, Moore and Hochgeschwender are collaborating to build a system using
luciferase enzymes that are also calcium sensitive, requiring
both the substrate and a calcium ion to illuminate. Firing
causes an influx of calcium into the neuronal cytoplasm,
so “you can think about a system where actually neuronal
activity turns on the light in the presence of substrate,”
Hochgeschwender explains.
Researchers can then identify the luminescent cells, even
through an intact skull. Hochgeschwender envisions a strategy where a scientist could determine which mice were expressing the protein in the right cells before performing an
experiment, eliminating those with inappropriate expression
patterns and substantially reducing the number of animals
required for an experiment.
Whether they use wire electrodes or bioluminescent
enzymes, though, scientists working with the new generation of techniques are palpably excited about the deluge
of recent advances. As Farb says, “Now I feel like this kid,
where we’re getting results and I’m just like ‘Wow, I can’t
believe this, I never thought of this, I never imagined it.’”
Alan Dove is a science writer and editor based in Massachusetts.
DOI: 10.1126/science.opms.p1500098
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