The Patch Clamp Goes Planar

The story goes like this: At a 1976 Biophysical Society meeting, Erwin Neher presented the technique that would win a Nobel Prize for him and Bert Sakmann.

By | May 23, 2005


The story goes like this: At a 1976 Biophysical Society meeting, Erwin Neher presented the technique that would win a Nobel Prize for him and Bert Sakmann. The room was packed with scientists anxious to hear his talk. But Neher, never a dynamic speaker, gave a flat recitation of how he and his colleague used a micropipette to create a tight seal on a tiny patch of membrane, which greatly reduced surface area and thus noise, allowing them to detect fainter signals than was previously possible.1 To anyone not a specialist, it sounded like nothing more than a generic scientific presentation.

Another scientist, with a better sense of occasion, used the accompanying question-and-answer session to give the moment the theatrical flourish it deserved. Recapping Neher's talk, he ended his summation with the dramatic question, "So what you're saying is, you've created a way to record single-ion channels?"

Thirty years later, the story has become part of the scientific apocrypha. But the technique's impact has been undeniable: The method transformed both neuroscience and biophysics and helped usher in a new branch of drug-development research focused on ion channels. "Patch clamping is a marvelous technique. It allows you to have access to currents across a cell in two ways: One is to look at very few channels, very few molecules, or to look at the whole cell with high resolution," says Francisco Bezanilla of the University of California, Los Angeles.

Yet it is a truism of science that yesterday's dramatic breakthrough is today's restrictive routine. Although Neher and Sakmann refined their original technique1 into the familiar "suck-on-a-cell" method,2 patch clamping is still a meticulous, time-consuming procedure requiring a skilled technician to manipulate pipettes and cells under a microscope. "It's a blend of art and science," says Henry Lester of the California Institute of Technology. "It requires manual dexterity, great attention to instrumentation, and continual feedback between the experiment and the experimenter. It's almost like landing an airplane on instruments."

"There are different patch configurations," explains Niels Fertig, CEO of Munich-based Nanion Technologies. "Excised patches are used for single-ion channel work mostly performed by academics, but for pharmacology the dominant configuration is whole cell." With the whole-cell technique, an experimenter first forms a tight seal against the cell membrane with the pipette or chip, then ruptures the membrane to get whole-cell access. "You get a measure of the response of all of the ion channels in the cell to a compound," says Fred Sigworth, a professor of cellular and molecular physiology at the Yale University School of Medicine and Neher and Sakmann's occasional collaborator.

Both techniques as they now exist work well in academia, where solo researchers perform individual experiments. In industry, however, where streamlining and throughput are important, the technique is a time-consuming anachronism. But scientists and equipment manufacturers are developing workarounds. Several companies have released or are planning to release commercial implementations of a new high-throughput variation on the technique, called planar patch clamping. In development since the late 1990s by several scientists, including Fred Sigworth and Kathryn Klemic, both of the department of cellular and molecular physiology at Yale University School of Medicine, this new approach is beginning to bridge the divide between academia and industry.


Planar patch clamping, as the name implies, is a two-dimensional (or planar) variation on traditional patch clamping. The microelectrode pipette, whose aperture provides the suction that couples the instrument to the cell, is replaced by a planar electrode substrate into which a hole has been bored. A technician, or even a robotic arm, can then pour a cell suspension over the aperture for recording, forming a seal with individual cells by applying suction from below. The technique is easier and faster than its predecessor and allows for recordings from multiple wells.

"It's really not that hard to put a hole in a substrate," says Klemic. "The real challenge is to form a gigaseal," the tight seal at the heart of the patch clamp technique. For the direct-voltage control necessary for ion channel measurement, the resistance between the electrode and the cell must be greater than one gigaohm (1012 ohms), indicating virtually no current flow.

Such high resistance is not necessary for all applications, however. Molecular Devices in Sunnyvale, Calif., which recently purchased competitor Axon Instruments, has an industrial-class instrument called IonWorks HT, whose resistance of approximately 100 megaohms is more than adequate to measure large ionic currents.

The IonWorks HT also is high-throughput, allowing planar patch clamp testing in 384 wells at once. Though this level of throughput is de rigueur in the pharmaceutical industry, the original intent behind the planar patch clamp was neither automation nor speed, but noise reduction. Imagine taping a conversation, only to have the recorder make so much noise that it drowns out the words. That's what the current instrumentation does.

Noise is inherent in the use of glass pipettes, says Sigworth. "Glass is not entirely a solid. At the molecular level there are ions in the glass that can move around. The thermal motion of the semitrapped ions in the glass couple to little random currents into the interior of the glass pipette."

The alternative is to use quartz, which is a much better dielectric. But its high melting point, says Bezanilla, is problematic, requiring the use of a CO2 laser. (Klemic says this is true only for pipettes. Planar electrodes made of quartz are not technically more difficult to fabricate than glass, just more expensive.)

Adding to the problem with glass is the cost and noise of amplifiers required to conduct electrophysiology work. "The technology's been pretty stable since the 80's," says Sigworth, "although there has been an improvement in noise level." But the box is still expensive, and can cost as much as $15,000.

An amplifier itself is very simple, says Bezanilla; it essentially is just a few transistors. The important part is the head stage, which is the transducer between the pipette and the amplifier. No matter how a system is ultimately laid out, the head stage must come in contact with every cell in order to record data. For a planar system, packing the electronics together can be challenging.

Klemic and Sigworth are now working with colleagues at Yale to make an on-chip amplifier. "To do massively parallel patch clamping, it would be great to have a lot smaller amplifiers," says Sigworth, "And in principle we should be able to get the noise level down and much higher-resolution current recordings than we're doing now. Right now, we can watch a channel open and it lets about a hundred ions through, and that little pulse of current, 100 ions, we can detect. But it would be really fun if we count the ions as they go through one by one. The mechanisms of actions in things like the sodium-potassium pump would become clear."

The best commercially available amplifiers have a noise limit of 100 electronic charges, but Sigworth and Klemic hope to drive noise down to 5–10 charges to pull out charge movement within the protein itself. Klemic says that the system they're currently working on will likely go down to 12–15 charges.

For their substrate, Klemic and Sigworth first used silicon, then quartz. Currently the Yale researchers are working with polydimethylsiloxane, a silicone elastomer known to many laboratories as the slightly rubbery Sylgard.3 Nanion Technologies is successfully using glass, says Bezanilla.

Nanion's product is based on Fertig's research, which Bezanilla and other scientists praise highly.4 "The dream is, you collect your tissue culture cells, you spin them down in a micro-centrifuge to wash them, and then you pipette a few microliters of your cells into a little chamber in your patch clamp device and you close the lid, and you click the mouse on your computer to start recording. Nanion actually makes a little device; they it the Port-a-Patch. It's a little tiny box that can sit your benchtop and you literally can do that," says Sigworth. Fertig says the company also has a 16-chip robotic system in late development.


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Several companies offer or are planning to offer variations on planar patch clamp systems (see sidebar at right). In a March review in IEEE Transactions on Nanobioscience,5 which Bezanilla calls, "a very good introduction into why people are doing this," Sigworth and Klemic overview the state of the art and the commercial product landscape. Klemic says she wrote the article to inspire engineers to come up with next-generation ideas. It does a "very good analysis of all the noise sources," says Bezanilla, who is well-known at UCLA for a course in which he teaches students to build their own electrophysiology rigs.

The review discusses the pair's current research on planar patch clamps, which may offer a solution to some of the noise problems. "Their technique is really quite powerful, in principle," says Bezanilla, "[but] they're not there yet." Barbara Ehrlich, a Yale colleague and professor of pharmacology, calls their research "spectacular."

Some companies are working on variations on patch clamping other than the planar approach. One such firm is flyion of Tübingen, Germany, which has developed an inside-out variation on the traditional patch clamp technique. Rather than drawing cells up towards the system's glass pipette tip, the company's automated "Flip-the-Tip" system drops cells inside the pipette, where they fall down towards the tip to form a seal.

An acute need exists to screen for effects on ion channels not just for new drugs, but also for potentially lethal complications in those already on the market. Although solid numbers are not available, deaths have been attributed to drugs that block the hERG potassium channel and thus prolong the Q-T interval, which is the time interval between the beginning and end of a cardiac contraction. As a result, the US Food and Drug Administration has issued draft guidance on how to evaluate a drug's potential for prolonging the Q-T interval.


© 2004 Nature Publishing Group

Cells are deposited by a robotic arm into the top compartment, and the electrical resistance between the compartments is monitored continuously (A). Suction or electric fields are applied to direct a cell to the hole (B). When the cell blocks the hole, the resistance rises, signaling the control system to decrease the pressure until a high-resistance (>1 gigaohm) seal is formed (C). It is then possible to record the currents from one to several individual ion channels in this area. If the whole-cell current is sought, the membrane in the hole must be broken by applying negative pressure in the bottom compartment (D,E), or a large voltage pulse between the compartments. Whole-cell membrane current (Im) recording is done by applying different voltage waveforms (V). At any time, drugs can be applied to the top compartment, while the electrical recording continues (F). (From F. Bezanilla et al., Nat Rev Drug Discovery, 3:239–78, 2004.)

Though not yet a requirement, says Raymond Woosely, president of the Critical Path Institute, a nonprofit devoted to accelerating the safe development of drugs, he recommends to pharmaceutical manufacturers that "any compound have at least some consideration of its effects on hERG and Q-T before they go into humans, because the question of Q-T prolongation is very likely to be raised by the FDA."

Planar patch clamping could provide the throughput necessary for this type of work. Multi Channel Systems of Reutlingen, Germany, has developed a 96-well plate with electrodes on the bottom of each well to measure the field potentials that are directly correlated to the Q-T interval.

But the technology has other applications, too, the breadth of which Lester says scientists may not yet fully appreciate. One possible use is single-cell electroporation. "You could easily foresee using membranes and apertures as ways of doing more precise electroporation ... if you want to do it in a high-throughput fashion, then you'd like to combine single-cell electroporation with apertures in a dish."

As for the current and future state of the market, Lester says, "The major challenge going forward will be convincing the manufacturers that there is a research market involving small numbers of cells," that is, one targeted to academia rather than industry. He continues, "The instrument manufacturers are fixated at the moment on multiwell, highly parallel instruments that are required by drug companies and driven by the high cost of high-throughput screening. For the moment no one has yet figured out how to build an automated or semiautomated patch clamp that will complement the single postdoc in the research lab." Nanion's original Port-a-Patch notwithstanding, Lester's point is valid.

But if manufacturers do dedicate themselves to easy-to-use single devices, what will happen to the meticulous art of individual patch clamping? "It will go the way of analytic ultracentrifugation," predicts Lester. "Not a lot of people own [Beckman Instrument's] Model E's anymore."

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