The microfluidic channels are fabricated in a cyclic polyolefin (Zeonex&reg; 690R, Zeon Chemical Inc., Louisville, KY). Zeonex has a glass-transition temperature (Tg) of 136&deg;C and is UV transparent, which is essential for the light directed chemistry used in the porous monolith formation. The microchannels are fabricated by micro-hot-embossing with a NiCo electroformed master-mold, which has the negative (inverse) features of the microfluidic platform. The hot-embossing is done as follows:
<ol>
<li>The mold and the polymer substrate are placed between two parallel metal platens.</li>
<li>The platens are heated up to the embossing temperature (166&deg;C), which is 30&ordm;C above the Tg of Zeonex.</li>
<li>The platens are then pressed together for approximately 5 min and the pressure is maintained at 250 psi.</li>
<li>The mold and the substrate are then removed from the heated platens, allowed to cool for 1-2 min, and then manually separated from each other.</li>
<li>Wells of 1.5-mm diameter are drilled at the ends of the channels for sample introduction and collection.</li>
<li>The embossed plastic substrate is then thermally bonded with another piece of Zeonex to form enclosed microfluidic channels. The embossed substrate and the cover piece can be UV-ozone or plasma treated prior to thermal bonding to enhance the sealing efficiency and ensure the fidelity of the enclosed channel structure (Bhattacharyya and Klapperich, 2007).</li>
</ol>
Fabrication of solid-phase extraction (SPE) column within the plastic microfluidic channels
Fabrication of the solid-phase is a two-step process. At first, the fabricated microchannels are surface modified in order to improve the adhesion of the SPE column to the plastic device. Grafting via surface photopolymerization is used to functionalize the walls of the polymeric microchannels.
The grafting procedure is as follows:
<ol>
<li>The microchannels are filled with a 1:1 mixture of ethylene diacrylate (EDA) and methyl methacrylate (MMA) with 3% benzophenone, which is a hydrogen abstracting photoinitiator. The EDA, MMA and benzophenone are purchased from Sigma-Aldrich, St. Louis, MO.</li>
<li>The chip is then UV-irradiated for 10 min at 254 nm UV wavelength and 200 mJ/cm2 energy in an ultraviolet exposure instrument (CL-1000 UV Crosslinker, UPV Inc., Upland, CA). The grafting occurs via H-abstraction polymerization and results in surface-tethered polymer chains, which get incorporated within the polymerization mixture used for the preparation of the monolith in the second step.</li>
<li>The excess monomer is removed from the channels by rinsing with 15 &mu;L of methanol using vacuum aspiration.</li>
</ol>
After the photografting process, a microporous polymer monolith is formed within the microchannels that entrap the silica particles for DNA extraction. The porous monolith is prepared in situ as follows:
<ol>
<li>The surface modified channels are filled with a monolith precursor mixture consisting of BuMA (15% wt), EDMA (10% wt), 1-dodecanol (52.5% wt), cyclohexanol (22.5% wt), DMPAP (1% wt with respect to monomers) and 0.7 &mu;m silica particles (15% with respect to the total volume of the pre-polymer mixture). The silica microspheres were purchased from Polysciences, Inc. (Warrington, PA) and the monomers and porogenic solvents are purchased from Sigma-Aldrich, St. Louis, MO.</li>
<li>The chip is irradiated with UV in the UV crosslinker at 200 mJ/cm2 for 1.1 min to initiate polymerization, and the excess monomer is removed from the channels by rinsing with 15 &mu;L of methanol using vacuum aspiration.</li>
<li>The fluidic connections (NanoPortsTM connections, Upchurch Scientific, Oak Harbor, WA) are then attached on the introduction and collection wells of the microfluidic channels.</li>
<li>The porous monolith is then washed again with methanol for at least 30 min using the syringe pump at a flow rate of 100mL/hour.</li>
</ol>
Fabrication of CNT lysis monolith within the plastic microfluidic channels
Fabrication of the CNT (carbon nanotube) lysis monolith is a two-step process. First, the fabricated microchannels are surface modified (grafted) in the same manner as for the SPE columns in the plastic device.
After the photografting process, a microporous polymer monolith is formed within the microchannel that is impregnated with multi-walled carbon nanotubes. The porous monolith is prepared in situ as follows:
<ol>
<li>Multi-walled nanotubes are resuspended in cyclohexanol at a concentration of 2.27M. This resuspension is ultrasonicated for 30 minutes at 50% amplitude and 50% duty cycle with a sonifier cell disruptor (Sonifier S-250D, manufactured by Branson Ultrasonic, Danbury, CT). The sonication provided an effective and stable 0.5M suspension of CNTs. The CNTs were purchased from Nanolabs (Brighton, MA).</li>
<li>The surface modified channels are filled with a similar monolith precursor mixture except that now the solution of cyclohexanol + CNTs is used. The monolith mixture consists of BuMA (15% wt), EDMA (10% wt), 1-dodecanol (52.5% wt), and cyclohexanol+CNTs (22.5% wt). To this solution the photoinitiator DMPAP is added (1.13% wt with respect to monomers). The monomers, porogenic solvents and DMPAPA are purchased from Sigma-Aldrich, St. Louis, MO.</li>
<li>The chip is irradiated with UV in the UV crosslinker at 120 mJ/cm2 for 0.9 min to initiate polymerization. The device is flipped 180 degrees in the crossliker and irradiated for another 0.9 min at 120 mJ/cm2. The excess monomer is removed from the channels by rinsing with 50 &mu;L of methanol using vacuum aspiration.</li>
<li>The fluidic connections (NanoPortsTM connections, Upchurch Scientific, Oak Harbor, WA) are then attached on the introduction and collection wells of the microfluidic channels.</li>
</ol>
Bacterial Sample Preparation for use with CNT lysis monolith
The sample preparation for a bacterial sample involves taking an optical density measurement at 600nm to ensure that the sample is not extremely concentrated so as to avoid clogging the channel.
<ol>
<li>Take an OD reading at 600nm with the bacteria in media, this is done with a biophotometer (Eppendorf BioPhotometer, Eppendorf Scientific, Inc., Westbury, NY). Dilute the sample until the OD readings fall within the range 0.18-0.25 A.</li>
<li>Centrifuge the sample at 6,500 rpm for 5 minutes and decant the supernatant. </li>
</ol>
Lysis
<ol>
<li>Resuspend bacteria in GuSCN* + .01%SDS + 4% Proteinase K (0.8mg/ml). Vortex briefly (1ml of bacteria in media should be resuspended in 1ml GuSCN). Proteinase K was added to the solution to aid with denaturing proteins that are attached to the bacterial DNA and to inhibit DNAases and RNAases. An additional benefit of the proteinase K is that it also aids with breaking down the cellular wall proteins which is desirable since both gram-positive bacteria and gram-negative bacteria cell walls contain large amounts of protein (although gram-negative bacteria typically have more protein). The chaotropic buffer additionally aids the proteinase K with denaturing proteins while the detergent disrupts the cell wall. Proteinase K and guanidinium thiocyanate (GuSCN) were purchased from Qiagen Inc. (Valencia, CA). SDS (Sodium Dodecyl Sulfate) was purchased from Pierce (Rockford, IL).</li>
<li>Immediately load the sample in a syringe connected to PEEK tubing. Aspirate the syringe so that no air is in the system. Connect the tubing to the nanoport and channel. </li>
</ol>
A KDS100 syringe pump (manufactured by KD Scientific, Holliston, MA) was used for the experiments and the flow rate used is 450 &mu;L/hour for all the steps. Flow the sample lysis channel at 450ul/hr.
<ol>
<li>Collect the sample and proceed to the DNA extraction via SPE (see DNA extraction protocol). NOTE: This bacterial sample will not need to follow Load Step 2, since the suspension is GuSCN*.</li>
</ol>
DNA Extraction Protocol
The extraction procedure consists of 3 steps:
<ol>
<li>Load.</li>
<li>Wash.</li>
<li>Elute.</li>
</ol>
KDS100 syringe pump (manufactured by KD Scientific, Holliston, MA) was used for the experiments and the flow rate used was 300 &mu;L/hour for all the steps.
Load
<ol>
<li>Condition the channel with chaotropic buffer (GuSCN*, guanidinium thiocyanate) for 1-2 min before starting the experiment.</li>
<li>Mix the sample with chaotropic buffer in 1:1 ratio.</li>
<li>Flow sample mixture through the channel for 3 min.</li>
</ol>
Wash
<ol>
<li>Wash the channel with 70% ethanol for 2 min.</li>
<li>Wash the channel with 100% ethanol for 2 min.</li>
</ol>
Elute
<ol>
<li>Elute the extracted DNA in millipore water for 3 min. Collect 15 &mu;L of purified, PCR-ready DNA.</li>
</ol>
*The GuSCN from the Qiagen kit (Buffer RLT) was used.
&nbsp;

<ol>
	<li>Bhattacharyya, A., Klapperich, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanical+and+chemical+analysis+of+plasma+and+ultraviolet-ozone+surface+treatments+for+thermal+bonding+of+polymeric+microfluidic+devices.">Mechanical and chemical analysis of plasma and ultraviolet-ozone surface treatments for thermal bonding of polymeric microfluidic devices.</a> Lab Chip. 7, 876-882 (2007).</li></ol>

The authors have nothing to disclose.

fabrication of the thermoplastic microfluidic channels

In our lab, we have successfully isolated nucleic acids directly from microliter and submicroliter volumes of human blood, urine and stool using polymer/nanoparticle composite microscale lysis and solid phase extraction columns. The recovered samples are concentrated, small volume samples that are PCRable, without any additional cleanup. Here, we demonstrate how to fabricate thermoplastic microfluidic chips using hot embossing and heat sealing. Then, we demonstrate how to use in situ light directed surface grafting and polymerization through the sealed chip to form the composite solid phase columns. We demonstrate grafting and polymerization of a carbon nanotube/polymer composite column for bacterial cell lysis. We then show the lysis process followed by solid phase extraction of nucleic acids from the sample on chip using a silica/polymer composite column. The attached protocols contain detailed instructions on how to make both lysis and solid phase extraction columns.

Watch this Scientific Journal Video about Fabrication of the Thermoplastic Microfluidic Channels at JoVE.com

Fabrication of the Thermoplastic Microfluidic Channels

Here we demonstrate how to fabricate thermoplastic microfluidic chips using hot embossing and heat sealing. Then we demonstrate how to use in situ light directed surface grafting and polymerization through the sealed chip to form the composite solid phase columns.

In our lab, we have successfully isolated nucleic acids directly from microliter and submicroliter volumes of human blood, urine and stool using ...

fabrication-of-the-thermoplastic-microfluidic-channels

Research

JoVE Journal

Biology

13.3K vues.  Boston University. Ici, nous montrer comment fabriquer des puces microfluidiques thermoplastiques en utilisant gaufrage à chaud et l'étanchéité de la chaleur. Ensuite, nous montrent comment utiliser la lumière à la surface in situ réalisé la greffe et la polymérisation à travers la puce scellée pour former des colonnes composites en phase solide.

Vidéo: Fabrication of the Thermoplastic Microfluidic Channels

Fabrication of a Microfluidic Device for the Compartmentalization of Neuron Soma and Axons

In this video, we demonstrate the technique of soft lithography with polydimethyl siloxane (PDMS) which we use to fabricate a microfluidic device for culturing neurons. Previously, a silicon wafer was patterned with the design for the neuron microfluidic device using SU-8 and photolithography to create a master mold, or what we simply refer to as a "master". Next, we pour the silicon polymer PDMS on top of the master which is then cured by heating the PDMS to 80&deg;C for 1 hour. The PDMS forms a negative mold of the device. The PDMS is then carefully cut and lifted away from the master. Holes are punched where the reservoirs will be and the excess PDMS trimmed away from the device. Nitrogen is used to blow away any excess debris from the device. At this point the devices are now ready for use and can either bonded to corning No. 1 cover glass with a plasma sterilizer/cleaner or can be reversibly bound to the cover glass by simply placing the device on top of the cover glass. The reversible bonding of the device to glass is covered in a separate video and requires first that the device be sterilized either with 70% ethanol or by autoclaving. Plasma treating sterilizes the devices so no further treatment is necessary. It is, however, important, when plasma-treating the devices, to add liquid to the devices within 10 minutes of the plasma treatment while the surfaces are still hydrophilic. Waiting longer than 10 minutes to add liquid to the device makes it difficult for the liquid to enter the device. The neuron devices are typically plasma-bound to cover glass and 0.5 mg/ml poly-L-lysine (PLL) in pH 8.5 borate buffer is immediately added to the device. After a minimum of 3 hours incubating with PLL, the devices are washed with dH2O water a minimum of 3 times with at least 15 minutes between each wash. Next, the water is removed and fresh media is added to the device. At this point the device is ready for use. It is important to remember at this point to never remove all the media from the device. Always leave media in the main channel.

In this video we demonstrate the technique of soft lithography with polydimethyl siloxane (PDMS) which we use to farbricate a microfluidic device for culturing neurons.

In this video, we demonstrate the technique of soft lithography with polydimethyl siloxane (PDMS) which we use to fabricate a microfluidic device for ...

Non-plasma Bonding of PDMS for Inexpensive Fabrication of Microfluidic Devices

In this video, we demonstrate how to use the neuron microfluidic device without plasma bonding.  In some cases it may be desirable to reversibly bond devices to the Corning No. 1 cover glass.  This could be due, perhaps, to a plasma cleaner not being available.  In other instances, it may be desirable to remove the device from the glass after the culturing of neurons for certain types of microscopy or for immunostaining, though it is not necessary to remove the device for immunostaining since the neurons can be stained in the device.  Some researchers, however, still prefer to remove the device.  In this case, reversible bonding of the device to the cover glass makes that possible.  There are some disadvantages to non-plasma bonding of the devices in that not as tight of a seal is formed.  In some cases axons may grow under the grooves rather than through them.  Also, because the glass and PDMS are hydrophobic, liquids do not readily enter the device making it necessary at times to force media and other reagents into the device.  Liquids will enter the device via capillary action, but it takes significantly longer as compared to devices that have been plasma bonded.  The plasma cleaner creates temporary hydrophilic charges on the glass and device that facilitate the flow of liquids through the device after bonding within seconds.  For non-plasma bound devices, liquid flow through the devices takes several minutes.  It is also important to note that the devices to be used with non-plasma bonding need to be sterilized first, whereas plasma treated devices do not need to be sterilized prior to use because the plasma cleaner will sterilize them.

In this video we demonstrate how to use the neuron microfluidic device without plasma bonding.

In this video, we demonstrate how to use the neuron microfluidic device without plasma bonding.  In some cases it may be desirable to reversibly bond ...

Microfluidic Applications for Disposable Diagnostics

In this interview, Dr. Klapperich discusses the fabrication of thermoplastic microfluidic devices and their application for development of new diagnostics.

In this interview, Dr. Klapperich discusses the fabrication of thermoplastic microfluidic devices and their application for development of new diagnostics.

In this interview, Dr. Klapperich discusses the fabrication of thermoplastic microfluidic devices and their application for development of new ...

Patch Clamp Recording of Ion Channels Expressed in  Xenopus Oocytes

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological measurement of single or multiple ion channels in cells. This technique can be applied to ion channels in both their native environment and expressed in heterologous cells, such as oocytes harvested from the African clawed frog, Xenopus laevis. Here, we describe the well-established technique of patch clamp recording from Xenopus oocytes. This technique is used to measure the properties of expressed ion channels either in populations (macropatch) or individually (single-channel recording). We focus on techniques to maximize the quality of oocyte preparation and seal generation. With all factors optimized, this technique gives a probability of successful seal generation over 90 percent. The process may be optimized differently by every researcher based on the factors he or she finds most important, and we present the approach that have lead to the greatest success in our hands.

This is intended as an introduction to patch clamp recording from Xenopus laevis oocytes. It covers vitelline membrane removal, formation of a gigaohm seal (gigaseal), and the optional conversion of the patch to the outside-out topology.

Since its development by Sakmann and Neher 1, 2, the patch clamp has become established as an extremely useful technique for electrophysiological ...

Fabrication of Amperometric Electrodes

Carbon fiber electrodes are crucial for the detection of catecholamine release from vesicles in single cells for amperometry measurements.  Here, we describe the techniques needed to generate low noise (&lt;0.5 pA) electrodes.  The techniques have been modified from published descriptions by previous researchers (1,2).  Electrodes are made by preparing carbon fibers and threading them individually into each capillary tube by using a vacuum with a filter to aspirate the fiber.  Next, the capillary tube with fiber is pulled by an electrode puller, creating two halves, each with a fine-pointed tip.  The electrodes are dipped in hot, liquid epoxy mixed with hardener to create an epoxy-glass seal.  Lastly, the electrodes are placed in an oven to cure the epoxy.  Careful handling of the electrodes is critical to ensure that they are made consistently and without damage.  This protocol shows how to fabricate and cut amperometric electrodes for recording from single cells.

This protocol describes how to generate carbon fiber electrodes. The electrodes are subsequently used to detect catecholamine release from vesicles with carbon fiber amperometry.

Carbon fiber electrodes are crucial for the detection of catecholamine release from vesicles in single cells for amperometry measurements.  Here, we ...

Fabrication of Myogenic Engineered Tissue Constructs

Despite the fact that electronic pacemakers are life-saving medical devices, their long-term performance in pediatric patients can be problematic owing to the restrictions imposed by a child's small size and their inevitable growth. Consequently, there is a genuine need for innovative therapies designed specifically for pediatric patients with cardiac rhythm disorders. We propose that a conductive biological alternative consisting of a collagen-based matrix containing autologously-derived cells could better adapt to growth, reduce the need for recurrent surgeries, and greatly improve the quality of life for these patients. In the present study, we describe a procedure for incorporating primary skeletal myoblast cell cultures within a hydrogel matrix to fashion a surgically-implantable tissue construct that will serve as an electrical conduit between the upper and lower chambers of the heart. Ultimately, we anticipate using this type of engineered tissue to restore atrioventricular electrical conduction in children with complete heart block. In view of that, we isolate myoblasts from the skeletal muscles of neonatal Lewis rats and plate them onto laminin-coated tissue culture dishes using a modified version of established protocols[2, 3]. After one to two days, cultured cells are collected and mixed with antibiotics, type 1 collagen, Matrigel&trade;, and NaHCO3. The result is a viscous, uniform solution that can be cast into a mold of nearly any shape and size[1, 4, 5]. For our tissue constructs, we employ type 1 collagen isolated from fetal lamb skin using standard procedures[6]. Once the tissue has solidified at 37&deg;C, culture media is carefully added to the plate until the construct is submerged. The engineered tissue is then allowed to further condense through dehydration for 2 more days, at which point it is ready for in vitro assessment or surgical-implantation.

Here, we demonstrate fabrication of collagen-based, tissue constructs containing skeletal myoblasts. These 3-D engineered constructs may be used to replace or repair tissues in vivo. For our purposes, we have designed these as an atrioventricular electrical conduit for the repair of complete heart block[1].

Despite the fact that electronic pacemakers are life-saving medical devices, their long-term performance in pediatric patients can be problematic owing to ...

The Microfluidic Probe: Operation and Use for Localized Surface Processing

Microfluidic devices allow assays to be performed using minute amounts of sample and have recently been used to control the microenvironment of cells. Microfluidics is commonly associated with closed microchannels which limit their use to samples that can be introduced, and cultured in the case of cells, within a confined volume. On the other hand, micropipetting system have been used to locally perfuse cells and surfaces, notably using push-pull setups where one pipette acts as source and the other one as sink, but the confinement of the flow is difficult in three dimensions. Furthermore, pipettes are fragile and difficult to position and hence are used in static configuration only. 
The microfluidic probe (MFP) circumvents the constraints imposed by the construction of closed microfluidic channels and instead of enclosing the sample into the microfluidic system, the microfluidic flow can be directly delivered onto the sample, and scanned across the sample, using the MFP. . The injection and aspiration openings are located within a few tens of micrometers of one another so that a microjet injected into the gap is confined by the hydrodynamic forces of the surrounding liquid and entirely aspirated back into the other opening. The microjet can be flushed across the substrate surface and provides a precise tool for localized deposition/delivery of reagents which can be used over large areas by scanning the probe across the surface. 
In this video we present the microfluidic probe1 (MFP). We explain in detail how to assemble the MFP, mount it atop an inverted microscope, and align it relative to the substrate surface, and finally show how to use it to process a substrate surface immersed in a buffer.

In this video we present the microfluidic probe1 (MFP). We explain in detail how to assemble the MFP, mount it atop an inverted microscope, and align it relative to the substrate surface, and finally show how to use it to process a substrate surface immersed in a buffer solution.

Microfluidic devices allow assays to be performed using minute amounts of sample and have recently been used to control the microenvironment of cells. ...

Mutagenesis and Functional Analysis of Ion Channels Heterologously Expressed in Mammalian Cells

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying potassium (referred to as GIRK) channels, which are important for regulating the excitability of neurons. There are four different mammalian GIRK channel subunits (GIRK1-GIRK4) - we focus on GIRK2 because it forms a homotetramer. Stimulation of different types of G protein-coupled receptors (GPCRs), such as the muscarinic receptor (M2R), leads to activation of GIRK channels. Alcohol also directly activates GIRK channels. We will show how to mutate one amino acid by specifically changing one or more nucleotides in the cDNA for the GIRK channel. This mutated cDNA sequence will be amplified in bacteria, purified, and the presence of the point mutation will be confirmed by DNA sequencing. The cDNAs for the mutated and wild-type GIRK channels will be transfected into human embryonic kidney HEK293T cells cultured in vitro. Lastly, whole-cell patch-clamp electrophysiology will be used to study the macroscopic potassium currents through the ectopically expressed wild-type or mutated GIRK channels. In this experiment, we will examine the effect of a L257W mutation in GIRK2 channels on M2R-dependent and alcohol-dependent activation.

We will demonstrate how to study the effect of a single point mutation on the function of an ion channel.

We will demonstrate how to study the functional effects of introducing a point mutation in an ion channel. We study G protein-gated inwardly rectifying ...

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may also be used to study the structure-function relationship for other ion channels and neurotransmitter receptors1. BK channels are widely expressed in different tissues and have been implicated in many physiological functions, including regulation of smooth muscle contraction, frequency tuning of inner hair cells and regulation of neurotransmitter release2-6. BK channels are activated by membrane depolarization and by intracellular Ca2+ and Mg2+ 6-9. Therefore, the protocol is designed to control both the membrane voltage and the intracellular solution. In this protocol, messenger RNA of BK channels is injected into Xenopus laevis oocytes (stage V-VI) followed by 2-5 days of incubation at 18&deg;C10-13. Membrane patches that contain single or multiple BK channels are excised with the inside-out configuration using patch clamp techniques10-13. The intracellular side of the patch is perfused with desired solutions during recording so that the channel activation under different conditions can be examined. To summarize, the mRNA of BK channels is injected into Xenopus laevis oocytes to express channel proteins on the oocyte membrane; patch clamp techniques are used to record currents flowing through the channels under controlled voltage and intracellular solutions.

Patch Clamp and Perfusion Techniques for Studying Ion Channels Expressed in Xenopus oocytes

Ionic current of BK channels is recorded using patch clamp techniques. BK channels are expressed in Xenopus oocytes by injecting messenger RNA. The intracellular solution during patch clamp recordings is controlled by a perfusion system.

The protocol presented here is designed to study the activation of the large conductance, voltage- and Ca2+-activated K+ (BK) channels. The protocol may ...

A Fluorescent Screening Assay for Identifying Modulators of GIRK Channels

G protein-gated inward rectifier K+ (GIRK) channels function as cellular mediators of a wide range of hormones and neurotransmitters and are expressed in the brain, heart, skeletal muscle and endocrine tissue1,2. GIRK channels become activated following the binding of ligands (neurotransmitters, hormones, drugs, etc.) to their plasma membrane-bound, G protein-coupled receptors (GPCRs). This binding causes the stimulation of G proteins (Gi and Go) which subsequently bind to and activate the GIRK channel. Once opened the GIRK channel allows the movement of K+ out of the cell causing the resting membrane potential to become more negative. As a consequence, GIRK channel activation in neurons decreases spontaneous action potential formation and inhibits the release of excitatory neurotransmitters. In the heart, activation of the GIRK channel inhibits pacemaker activity thereby slowing the heart rate.
GIRK channels represent novel targets for the development of new therapeutic agents for the treatment neuropathic pain, drug addiction, cardiac arrhythmias and other disorders3. However, the pharmacology of these channels remains largely unexplored. Although a number of drugs including anti-arrhythmic agents, antipsychotic drugs and antidepressants block the GIRK channel, this inhibition is not selective and occurs at relatively high drug concentrations3.
Here, we describe a real-time screening assay for identifying new modulators of GIRK channels. In this assay, neuronal AtT20 cells, expressing GIRK channels, are loaded with membrane potential-sensitive fluorescent dyes such as bis-(1,3-dibutylbarbituric acid) trimethine oxonol [DiBAC4(3)] or HLB 021-152 (Figure 1). The dye molecules become strongly fluorescent following uptake into the cells (Figure 1). Treatment of the cells with GPCR ligands stimulates the GIRK channels to open. The resulting K+ efflux out of the cell causes the membrane potential to become more negative and the fluorescent signal to decrease (Figure 1). Thus, drugs that modulate K+ efflux through the GIRK channel can be assayed using a fluorescent plate reader. Unlike other ion channel screening assays, such atomic absorption spectrometry4 or radiotracer analysis5, the GIRK channel fluorescent assay provides a fast, real-time and inexpensive screening procedure.

A real-time screening procedure for identifying drugs that interact with G protein-gated inward rectifier K+ (GIRK) channels is described. The assay utilizes membrane potential-sensitive fluorescent dyes to measure GIRK channel activity. This technique is adaptable for use on a number of cell lines.

G protein-gated inward rectifier K+ (GIRK) channels function as cellular mediators of a wide range of hormones and neurotransmitters and are expressed in ...