Name: patch clamp recordings on intact dorsal root ganglia from adult rats
Uploaded: 2016-09-29T13:00:00
Description: Watch this Scientific Journal Video about Patch Clamp Recordings on Intact Dorsal Root Ganglia from Adult Rats at JoVE.com

The authors would like to acknowledge the Painless Research Foundation for support of the work. This work was also supported by the NIH grants R01 NS080921-01 and R21 NS079897-01A1.

Ethics Statement: All procedures for the maintenance and use of the experimental animals conformed to the regulations of UCSF Committees on Animal Research and were carried out in accordance with the guidelines of the NIH regulations on animal use and care (Publication 85 - 23, Revised 1996). The UCSF Institutional Animal Care and Use Committee approved the protocols used in this study.
1. Preparation of Instruments, Solutions and Dishes
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	<li>Prepare Artificial Cerebrospinal Fluid (aCSF...

<ol>
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We report a method to prepare whole DRGs for patch clamp studies. There are several key elements for preparing an ideal specimen. Firstly, it is important to dissect the DRGs with dorsal roots attached. After that, the epineurium need to be carefully removed while avoiding damage to the neurons. Finally, to expose the neurons and their surrounding satellite glial cells, it is necessary to digest the remaining connective tissue. Intact DRGs from adult rats prepared with the method described here will maintain good viabili...

Sensation is essential to an organism&#39;s survival and wellbeing. The transmission of stimuli is dependent on the sensory pathways starting at peripheral endings of axons from primary sensory neurons. Primary sensory neurons, with the exception of the mesencephalic nucleus of the trigeminal nerve, are located in the trigeminal ganglia and dorsal root ganglia (DRGs). They serve as gatekeepers of the sensory information 1. At the perikarial membrane, just as at the central and peripheral terminals, DRG neurons express receptors and ion channels, such as glutamate receptors, TNF alpha receptors, transient receptor potential cation channel subfamily V member 1 (TRPV1), sodium channels, etc. 2-7. Patch clamp recordings of the perikarial membrane allow understanding functional changes of many of these receptors and channels throughout the neuron.
The patch clamp recording technique is a powerful tool for studying the activities of channels or receptors and a great number of studies have been conducted by applying this technique on DRG neurons 8-10. In most studies the DRG is removed by cutting the dorsal rootlets and spinal nerve close to the ganglion. After mincing, the ganglion is then placed in digestive enzymes that result in dissociation of the DRG neurons, which may then be recorded immediately or cultured for several days prior to recording. Unfortunately, dissociation of DRG neurons involves a necessary axotomy close to the perikarya. Once dissociated and axotomized, DRG neurons undergo phenotypic changes as well as changes in membrane excitability 11,12. The loss of contact between the perikarya of individual neurons and the satellite glial cells that normally surround them is likely to contribute to these changes 13. The crosstalk between neurons and satellite glial cells is both essential in physiological conditions and in adaptation to pathological conditions such as those leading to intractable pain 14,15. It would be challenging to study the interaction between neurons and satellite glial cells using a dissociated DRG preparation.
Intact DRGs, on the other hand, provide closer to in vivo conditions. In the past several years, our laboratory, as well as some other groups, has been using intact DRGs from adult rats to investigate changes of primary sensory neurons in different conditions associated with chronic pain 3-5,11,15-17. Although the techniques used in these studies are somewhat established, a step-by-step description has not yet been published. In the present manuscript, we describe a convenient and fast way to prepare intact DRGs and their use for patch clamp recordings.

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			<td>BD</td>
			<td>309659</td>
			<td>1 ml, 5 ml.</td>
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			<td>BD</td>
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			<td>size: 15</td>
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			<td height="20" style="height:20px;">Mayo straight scissor</td>
			<td>Fine Science Tools</td>
			<td>14010-15</td>
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			<td height="20" style="height:20px;">Mayo curved scissor</td>
			<td>Fine Science Tools</td>
			<td>14011-15</td>
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			<td height="20" style="height:20px;">Rongeur</td>
			<td>Fine Science Tools</td>
			<td>16021-14</td>
			<td></td>
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			<td height="20" style="height:20px;">Adson toothed forceps</td>
			<td>Fine Science Tools</td>
			<td>11027-12</td>
			<td></td>
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			<td height="20" style="height:20px;">Iris Scissor</td>
			<td>Fine Science Tools</td>
			<td>14084-08</td>
			<td></td>
		</tr>
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			<td height="20" style="height:20px;">Noyes spring scissor</td>
			<td>Fine Science Tools</td>
			<td>15124-12</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Bone scissors</td>
			<td>Fine Science Tools</td>
			<td>16044-10</td>
			<td>Special for cutting the bones.&#160;</td>
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			<td height="20" style="height:20px;">Forceps: Dumont, Dumoxel Biologie #5</td>
			<td>Fine Science Tools</td>
			<td>11252-30</td>
			<td>These have the fine tips that do not need sharpening when first purchased.</td>
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			<td height="21" style="height:21px;">periosteal elevator</td>
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			<td>WILD</td>
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			<td></td>
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			<td height="20" style="height:20px;">Transfer pipette</td>
			<td>Fisher brand</td>
			<td>13-711-5AM</td>
			<td></td>
		</tr>
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			<td height="20" style="height:20px;">Petri dish (10 cm)</td>
			<td>Pyrex</td>
			<td></td>
			<td>Glass petri dish can avoid damaging the tips of fine forceps</td>
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			<td height="20" style="height:20px;">Collagenase (Liberase TM)</td>
			<td>Roche</td>
			<td>05-401-119-001</td>
			<td>dissolve at the concentration of 13 u/ml, aliquot into glass pipette. Avoid repeated freeze and thaw.</td>
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			<td height="20" style="height:20px;">filter</td>
			<td>Thermo scientific</td>
			<td>7232520</td>
			<td>Filter the internal solutions for patch clamp recording to avoid clog.</td>
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		<tr height="20">
			<td height="20" style="height:20px;">Glass pipette</td>
			<td>Sutter</td>
			<td>BF150-110-7.5</td>
			<td></td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;">Anchor</td>
			<td>Havard apparatus</td>
			<td>64-0250</td>
			<td>stabilize the DRG to avoid drift.</td>
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		<tr height="20">
			<td height="20" style="height:20px;">Peristaltic pump</td>
			<td>WPI</td>
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			<td></td>
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			<td height="20" style="height:20px;">Pipette puller</td>
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			<td>P97</td>
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			<td>Molecular devices</td>
			<td>Axopatch 200B</td>
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			<td height="20" style="height:20px;">Digitizer</td>
			<td>Molecular devices</td>
			<td>1440D</td>
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			<td>NIKON</td>
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			<td>Sutter</td>
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			<td>Local Medical Gas supplier</td>
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patch clamp recordings on intact dorsal root ganglia from adult rats

Patch clamp studies from dorsal root ganglia (DRGs) neurons have increased our understanding of the peripheral nervous system. Currently, the majority of recordings are conducted on dissociated DRG neurons, which is a standard preparation for most laboratories. Neuronal properties, however, can be altered by axonal injury resulting from enzyme digestion used in acquiring dissociated neurons. Further, dissociated neuron preparations cannot fully represent the microenvironment of the DRG since loss of contact with satellite glial cells that surround the primary sensory neurons is an unavoidable consequence of this method. To overcome the limitations in using conventional dissociated DRG neurons for patch clamp recordings, in this report we describe a method to prepare intact DRGs and conduct patch clamp recordings on individual primary sensory neurons ex vivo. This approach permits the fast and straightforward preparation of intact DRGs, mimicking in vivo conditions by keeping DRG neurons associated with their surrounding satellite glial cells and basement membrane. Furthermore, the method avoids axonal injury from manipulation and enzyme digestion such as when dissociating DRGs. This ex vivo preparation can additionally be used to study the interaction between primary sensory neurons and satellite glial cells.

Figure 1 shows the process of preparing intact DRG for patch recording. Figure 1A shows the exposure and location of the ganglia after laminectomy. Figure1B shows L3, L4 and L5 DRGs with the nerve roots attached after removing the spinal cord. Then L4 and 5 DRGs are carefully dissected and freed from the vertebrae. Next, the epineurium, a transparent membrane surrounding the DRG, is removed (yellow arrow, Figure 1D). The ...

Watch this Scientific Journal Video about Patch Clamp Recordings on Intact Dorsal Root Ganglia from Adult Rats at JoVE.com

Patch Clamp Recordings on Intact Dorsal Root Ganglia from Adult Rats

This manuscript describes how to prepare intact dorsal root ganglia for patch clamp recordings. This preparation maintains the microenvironment for neurons and satellite glial cells, thus avoiding the phenotypic and functional changes seen using dissociated DRG neurons.

Patch clamp studies from dorsal root ganglia (DRGs) neurons have increased our understanding of the peripheral nervous system. Currently, the majority of ...

The overall goal for this experiment is to introduce methods to prepare the intact dorsal root ganglia and to conduct patch clamp recording using this preparation. This method allows single cell recording from the intact dorsal root ganglia while stimulating the entering spinal nerve or the exiting dorsal root. This approach is especially useful when studying the contribution of primary sensory neurons to pain.The main advantage of this technique is that it maintains both neuron and adjacent satellite glial cells. Therefore, this preparation is closer to in vivo situation. In this procedure, after the rat has been anesthetized, shave its lumbar area.Then, incise the skin and the subcutaneous tissue along the midline. Afterward, detach the perispinal muscles from the spinous processes at the L1 to S2 level using a fine periosteal elevator. Next, cut the spinous processes from L1 to S2.Subsequently, make two transverse cuts in the exposed vertebral column at approximately L1 and S1 and remove this segment of the vertebral column en bloc.Then, quickly submerge it into cold aCSF for two to three minutes. After that, flush off the blood from the removed vertebral column and transfer the vertebral column to a petri dish with cold aCSF, then use a fine rongeur to remove the laminae. Cut the dura mater along the midline with microscissors to expose the spinal cord.Next, transfer the remaining vertebra with DRG to the second petri dish filled with cold aCSF. Identify the L4 and L5 DRGs based on their relative position to transverse processes and the sciatic nerve. Then, free the DRG from the surrounding connective tissues and keep the nerve root and spinal nerve attached to the DRG.After that, transfer the DRG into the third petri dish for further dissection. Cut an opening where the dorsal and ventral roots join the DRG and separate the ventral root from it. Then, use forceps with blunt tips to hold the epineurium and use another fine forceps to roll the DRG from the epineurium.Continue to remove as much epineurium as possible that is attached to the DRG. Now, transfer the DRG to the recording chamber. Make sure the side of the DRG where the nerve roots originate faces down.Next, stabilize the DRG with an anchor. Then, perfuse the DRG with oxygenated aCSF through the plastic tubes connected with a peristaltic pump and allow the cells to recover for 30 minutes. After 30 minutes, assess the DRG quality under IR-DIC optics at 40x magnification through a CCD camera.Individually connected two one milliliter syringes to the pipette holders through two pieces of small diameter rubber tubing. Then, place one glass pipette filled with collagenase into one of the pipette holders and an empty glass pipette into the other one. Subsequently, use the micromanipulator to position the pipettes just above the DRG.Collide the tips of two pipettes against each other gently to enlarge the openings of the pipette tips. After that, move the pipette filled with collagenase close to the DRG surface. Briefly apply positive pressure to the pipette containing collagenase through the tube connecting the holder and the syringe by displacing the plunger about 0.5 milliliters.After 10 to 15 minutes, when the debris of the remaining epineurium is observed, apply gentle negative pressure to the empty pipette to suck away the debris. In this way, the neurons and surrounding satellite glial cells will be clearly exposed and ready for patch recording. In this procedure, place the electrode solution on ice to prevent degradation.Next, fill the glass patch pipette with filtered intracellular solution. Then, place the pipette into the head stage pipette holder. Apply gentle positive pressure to the pipette through a five milliliter syringe by displacing the plunger about one milliliter before lowering the pipette into the bath solution.Subsequently, choose V-Clamp mode on the amplifier and open the membrane test interface in the software. Under the microscope, move the pipette close to the target neuron. Next, apply positive pressure from the pipette to traverse the satellite glial cell layer until a sudden enlargement of space between the neuron and the surrounding layer of satellite glial cells is observed.Then, keep moving the pipette towards the neuron until a dimple is observed on the neuron. In our preparation, the neurons are surrounded by satellite glial cells. Therefore, to penetrate the glial cell layers and to reach individual neurons, the recording pipette is maintained at a high positive pressure.After that, reduce the positive pressure. Achieve a giga-ohm seal with gentle suction. To obtain whole cell recording configuration, penetrate the neuron cell membrane via a short, but strong, suction.Alternatively, use the zap function on the amplifier while suction is applied. Once a whole cell mode is established, compensate the whole cell capacitance and series resistance by turning the capacitance and resistance compensation knobs on the amplifier. Then, close the membrane test window.Choose I-Clamp Normal Mode by turning the mode knob on the amplifier. Afterward, click Open Protocol in the software. Select and load the protocol for measuring rheobase and click record to start recording.To examine the neuronal excitability, measure the input resistance and rheobase by injecting a graded series of depolarizing currents in steps of 100 picoamps. Next, click Open Protocol again and select the protocol for measuring membrane threshold. Subsequently, inject a 500 millisecond depolarizing ramp current to the neuron.To record the ligand induced currents, fill a pipette with specific agonists. Check the pipette to make sure there are no air bubbles inside. Then, place the pipette in the pipette holder which is connected to a drug dispensing system.Use the manipulator to move the pipette to within 15 micrometers of the neuron. Set the drug dispensing system pressure to one PSI and the duration to one second. Then, switch the recording mode to voltage clamp at 70 millivolts.Briefly apply pressure via the drug dispensing system to record a drug induced current. In this figure, rheobase was measured by injecting a series of currents into the DRG neuron. The lowest current intensity which can induce an action potential is defined as rheobase, as indicated by the arrow.The rheobase for this neuron is 300 picoamps. The membrane threshold was measured by injecting a ramp current. The potential is defined as the membrane threshold when an action potential is evoked.The membrane threshold for this neuron is 11.9 millivolts. In this figure, currents were induced by puff application of glutamate or AITC for one second. Both agonists induced inward currents, suggesting the presence of glutamate receptors and TRPA-1 receptors on small DRG neurons.Once mastered, this technique can be finished within one hour if it is performed properly. While attempting this procedure, it is important to remember to remove the epineurium and maintain the high pressure for the recording pipette. Following this procedure, patch clamp recording on satellite glial cells can be performed in order to answer additional questions like receptors and channels in satellite glial cells.This technique has paved the way for researchers in the field of pain to study the contribution of dorsal root ganglia to nociception. After watching this video, you should have a good understanding on how to prepare a whole dorsal root ganglia and then how to patch clamp from single cells on these whole dorsal root ganglia.

patch-clamp-recordings-on-intact-dorsal-root-ganglia-from-adult-rats

Research

JoVE Journal

Biochemistry

Time-resolved ElectroSpray Ionization Hydrogen-deuterium Exchange Mass Spectrometry for Studying Protein Structure and Dynamics

Intrinsically disordered proteins (IDPs) have long been a challenge to structural biologists due to their lack of stable secondary structure elements. Hydrogen-Deuterium Exchange (HDX) measured at rapid time scales is uniquely suited to detect structures and hydrogen bonding networks that are briefly populated, allowing for the characterization of transient conformers in native ensembles. Coupling of HDX to mass spectrometry offers several key advantages, including high sensitivity, low sample consumption and no restriction on protein size. This technique has advanced greatly in the last several decades, including the ability to monitor HDX labeling times on the millisecond time scale. In addition, by incorporating the HDX workflow onto a microfluidic platform housing an acidic protease microreactor, we are able to localize dynamic properties at the peptide level. In this study, Time-Resolved ElectroSpray Ionization Mass Spectrometry (TRESI-MS) coupled to HDX was used to provide a detailed picture of residual structure in the tau protein, as well as the conformational shifts induced upon hyperphosphorylation.

Conformational flexibility plays a critical role in protein function. Herein, we describe the use of time-resolved electrospray ionization mass spectrometry coupled to hydrogen-deuterium exchange for probing the rapid structural changes that drive function in ordered and disordered proteins.

Intrinsically disordered proteins (IDPs) have long been a challenge to structural biologists due to their lack of stable secondary structure elements. ...

Voltage-clamp Fluorometry in Xenopus Oocytes Using Fluorescent Unnatural Amino Acids

Voltage-Clamp Fluorometry (VCF) has been the technique of choice to investigate the structure and function of electrogenic membrane proteins where real-time measurements of fluorescence and currents simultaneously report on local rearrangements and global function, respectively1. While high-resolution structural techniques such as cryo-electron microscopy or X-ray crystallography provide static images of the proteins of interest, VCF provides dynamic structural data that allows us to link the structural rearrangements (fluorescence) to dynamic functional data (electrophysiology). Until recently, the thiol-reactive chemistry used for site-directed fluorescent labeling of the proteins restricted the scope of the approach because all accessible cysteines, including endogenous ones, will be labeled. It was thus required to construct proteins free of endogenous cysteines. Labeling was also restricted to sites accessible from the extracellular side. This changed with the use of Fluorescent Unnatural Amino Acids (fUAA) to specifically incorporate a small fluorescent probe in response to stop codon suppression using an orthogonal tRNA and tRNA synthetase pair2. The VCF improvement only requires a two-step injection procedure of DNA injection (tRNA/synthetase pair) followed by RNA/fUAA co-injection. Now, labelling both intracellular and buried sites is possible, and the use of VCF has expanded significantly. The VCF technique thereby becomes attractive for studying a wide range of proteins and &#8211; more importantly &#8211; allows investigating numerous cytosolic regulatory mechanisms.

Voltage-clamp Fluorometry in Xenopus Oocytes Using Fluorescent Unnatural Amino Acids

This article describes an enhancement of conventional Voltage-Clamp Fluorometry (VCF) where Fluorescent Unnatural Amino Acids (fUAA) are used instead of maleimide dyes, to probe structural rearrangements in ion channels. The procedure includes Xenopus oocyte DNA injection, RNA/fUAA coinjection, and simultaneous current and fluorescence measurements.

Voltage-Clamp Fluorometry (VCF) has been the technique of choice to investigate the structure and function of electrogenic membrane proteins where ...

Glycoproteomics of the Extracellular Matrix: A Method for Intact Glycopeptide Analysis Using Mass Spectrometry

Fibrosis is a hallmark of many cardiovascular diseases and is associated with the exacerbated secretion and deposition of the extracellular matrix (ECM). Using proteomics, we have previously identified more than 150 ECM and ECM-associated proteins in cardiovascular tissues. Notably, many ECM proteins are glycosylated. This post-translational modification affects protein folding, solubility, binding, and degradation. We have developed a sequential extraction and enrichment method for ECM proteins that is compatible with the subsequent liquid chromatography tandem mass spectrometry (LC-MS/MS) analysis of intact glycopeptides. The strategy is based on sequential incubations with NaCl, SDS for tissue decellularization, and guanidine hydrochloride for the solubilization of ECM proteins. Recent advances in LC-MS/MS include fragmentation methods, such as combinations of higher-energy collision dissociation (HCD) and electron transfer dissociation (ETD), which allow for the direct compositional analysis of glycopeptides of ECM proteins. In the present paper, we describe a method to prepare the ECM from tissue samples. The method not only allows for protein profiling but also the assessment and characterization of glycosylation by MS analysis.

This paper describes a methodology to prepare cardiovascular tissue samples for MS analysis that allows for (1) the analysis of ECM protein composition, (2) the identification of glycosylation sites, and (3) the compositional characterization of glycan forms. This methodology can be applied, with minor modifications, to the study of the ECM in other tissues.

Fibrosis is a hallmark of many cardiovascular diseases and is associated with the exacerbated secretion and deposition of the extracellular matrix (ECM). ...

A Simple Method for High Throughput Chemical Screening in Caenorhabditis Elegans

Caenorhabditis elegans is a useful organism for testing chemical effects on physiology. Whole organism small molecule screens offer significant advantages for identifying biologically active chemical structures that can modify complex phenotypes such as lifespan. Described here is a simple protocol for producing hundreds of 96-well culture plates with fairly consistent numbers of C. elegans in each well. Next, we specified how to use these cultures to screen thousands of chemicals for effects on the lifespan of the nematode C. elegans. This protocol makes use of temperature sensitive sterile strains, agar plate conditions, and simple animal handling to facilitate the rapid and high throughput production of synchronized animal cultures for screening.

A Simple Method for High Throughput Chemical Screening in Caenorhabditis Elegans

Here we describe a simple protocol for rapidly producing hundreds of nematode growth media agar, 96-well culture plates with consistent numbers of Caenorhhabditis elegans per well. These cultures are useful for the phenotypic screening of whole organisms. We focus here on using these cultures to screen chemicals for pro-longevity effects.

Caenorhabditis elegans is a useful organism for testing chemical effects on physiology. Whole organism small molecule screens offer significant advantages ...

Electrochemical Detection of Deuterium Kinetic Isotope Effect on Extracellular Electron Transport in Shewanella oneidensis MR-1

Direct electrochemical detection of c-type cytochrome complexes embedded in the bacterial outer membrane (outer membrane c-type cytochrome complexes; OM c-Cyts) has recently emerged as a novel whole-cell analytical method to characterize the bacterial electron transport from the respiratory chain to the cell exterior, referred to as the extracellular electron transport (EET). While the pathway and kinetics of the electron flow during the EET reaction have been investigated, a whole-cell electrochemical method to examine the impact of cation transport associated with EET has not yet been established. In the present study, an example of a biochemical technique to examine the deuterium kinetic isotope effect (KIE) on EET through OM c-Cyts using a model microbe, Shewanella oneidensis MR-1, is described. The KIE on the EET process can be obtained if the EET through OM c-Cyts acts as the rate-limiting step in the microbial current production. To that end, before the addition of D2O, the supernatant solution was replaced with fresh media containing a sufficient amount of the electron donor to support the rate of upstream metabolic reactions, and to remove the planktonic cells from a uniform monolayer biofilm on the working electrode. Alternative methods to confirm the rate-limiting step in microbial current production as EET through OM c-Cyts are also described. Our technique of a whole-cell electrochemical assay for investigating proton transport kinetics can be applied to other electroactive microbial strains.

Electrochemical Detection of Deuterium Kinetic Isotope Effect on Extracellular Electron Transport in Shewanella oneidensis MR-1

Here we present a protocol of whole-cell electrochemical experiments to study the contribution of proton transport to the rate of extracellular electron transport via the outer-membrane cytochromes complex in Shewanella oneidensis MR-1.

Direct electrochemical detection of c-type cytochrome complexes embedded in the bacterial outer membrane (outer membrane c-type cytochrome complexes; OM ...

Analyzing Dynamic Protein Complexes Assembled On and Released From Biolayer Interferometry Biosensor Using Mass Spectrometry and Electron Microscopy

In vivo, proteins are often part of large macromolecular complexes where binding specificity and dynamics ultimately dictate functional outputs. In this work, the pre-endosomal anthrax toxin is assembled and transitioned into the endosomal complex. First, the N-terminal domain of a cysteine mutant lethal factor (LFN) is attached to a biolayer interferometry (BLI) biosensor through disulfide coupling in an optimal orientation, allowing protective antigen (PA) prepore to bind (Kd 1 nM). The optimally oriented LFN-PAprepore complex then binds to soluble capillary morphogenic gene-2 (CMG2) cell surface receptor (Kd 170 pM), resulting in a representative anthrax pre-endosomal complex, stable at pH 7.5. This assembled complex is then subjected to acidification (pH 5.0) representative of the late endosome environment to transition the PAprepore into the membrane inserted pore state. This PApore state results in a weakened binding between the CMG2 receptor and the LFN-PApore and a substantial dissociation of CMG2 from the transition pore. The thio-attachment of LFN to the biosensor surface is easily reversed by dithiothreitol. Reduction on the BLI biosensor surface releases the LFN-PAprepore-CMG2 ternary complex or the acid transitioned LFN-PApore complexes into microliter volumes. Released complexes are then visualized and identified using electron microscopy and mass spectrometry. These experiments demonstrate how to monitor the kinetic assembly/disassembly of specific protein complexes using label-free BLI methodologies and evaluate the structure and identity of these BLI assembled complexes by electron microscopy and mass spectrometry, respectively, using easy-to-replicate sequential procedures.

Here we present a protocol to monitor the assembly and disassembly of the anthrax toxin using biolayer interferometry (BLI). Following assembly/disassembly on the biosensor surface, the large protein complexes are released from the surface for visualization and identification of components of the complexes using electron microscopy and mass spectrometry, respectively.

In vivo, proteins are often part of large macromolecular complexes where binding specificity and dynamics ultimately dictate functional outputs. In this ...

Measurement of Ion Concentration in the Unstirred Boundary Layer with Open Patch-Clamp Pipette: Implications in Control of Ion Channels by Fluid Flow

Fluid flow is an important environmental stimulus that controls many physiological and pathological processes, such as fluid flow-induced vasodilation. Although the molecular mechanisms for the biological responses to fluid flow/shear force are not fully understood, fluid flow-mediated regulation of ion channel gating may contribute critically. Therefore, fluid flow/shear force sensitivity of ion channels has been studied using the patch-clamp technique. However, depending on the experimental protocol, the outcomes and interpretation of data can be erroneous. Here, we present experimental and theoretical evidence for fluid flow-related errors and provide methods for estimating, preventing, and correcting these errors. Changes in junction potential between the Ag/AgCl reference electrode and bathing fluid were measured with an open pipette filled with 3 M KCl. Fluid flow could then shift the liquid/metal junction potential to approximately 7 mV. Conversely, by measuring the voltage shift induced by fluid flow, we estimated the ion concentration in the unstirred boundary layer. In the static condition, the real ion concentrations adjacent to the Ag/AgCl reference electrode or ion channel inlet at the cell-membrane surface can reach as low as approximately 30% of that in the flow condition. Placing an agarose 3 M KCl bridge between the bathing fluid and reference electrode may have prevented this problem of junction potential shifting. However, the unstirred layer effect adjacent to the cell membrane surface could not be fixed in this way. Here, we provide a method for measuring real ion concentrations in the unstirred boundary layer with an open patch-clamp pipette, emphasizing the importance of using an agarose salt-bridge while studying fluid flow-induced regulation of ion currents. Therefore, this novel approach, which takes into consideration the real concentrations of ions in the unstirred boundary layer, may provide useful insight on the experimental design and data interpretation related to fluid shear stress regulation of ion channels.

Mechanosensitive ion channels are often studied in terms of fluid flow/shear force sensitivity with patch-clamp recording. However, depending on the experimental protocol, the outcome on fluid flow-regulations of ion channels can be erroneous. Here, we provide methods for preventing and correcting such errors with a theoretical basis.

Fluid flow is an important environmental stimulus that controls many physiological and pathological processes, such as fluid flow-induced vasodilation. ...

Visualizing Surface T-Cell Receptor Dynamics Four-Dimensionally Using Lattice Light-Sheet Microscopy

The signaling and function of a cell are dictated by the dynamic structures and interactions of its surface receptors. To truly understand the structure-function relationship of these receptors in situ, we need to visualize and track them on the live cell surface with enough spatiotemporal resolution. Here we show how to use recently developed Lattice Light-Sheet Microscopy (LLSM) to image T-cell receptors (TCRs) four-dimensionally (4D, space and time) at the live cell membrane. T cells are one of the main effector cells of the adaptive immune system, and here we used T cells as an example to show that the signaling and function of these cells are driven by the dynamics and interactions of the TCRs. LLSM allows for 4D imaging with unprecedented spatiotemporal resolution. This microscopy technique therefore can be generally applied to a wide array of surface or intracellular molecules of different cells in biology.

The goal of this protocol is to show how to use Lattice Light-Sheet Microscopy to four-dimensionally visualize surface receptor dynamics in live cells. Here T cell receptors on CD4+ primary T cells are shown.

The signaling and function of a cell are dictated by the dynamic structures and interactions of its surface receptors. To truly understand the ...

Measuring Nucleotide Binding to Intact, Functional Membrane Proteins in Real Time

We have developed a method to measure binding of adenine nucleotides to intact, functional transmembrane receptors in a cellular or membrane environment. This method combines expression of proteins tagged with the fluorescent non-canonical amino acid ANAP, and FRET between ANAP and fluorescent (trinitrophenyl) nucleotide derivatives. We present examples of nucleotide binding to ANAP-tagged KATP ion channels measured in unroofed plasma membranes and excised, inside-out membrane patches under voltage clamp. The latter allows for simultaneous measurements of ligand binding and channel current, a direct readout of protein function. Data treatment and analysis are discussed extensively, along with potential pitfalls and artefacts. This method provides rich mechanistic insights into the ligand-dependent gating of KATP channels and can readily be adapted to the study of other nucleotide-regulated proteins or any receptor for which a suitable fluorescent ligand can be identified.

This protocol presents a method for measuring adenine nucleotide binding to receptors in real time in a cellular environment. Binding is measured as F&#246;rster resonance energy transfer (FRET) between trinitrophenyl nucleotide derivatives and protein labeled with a non-canonical, fluorescent amino acid.

We have developed a method to measure binding of adenine nucleotides to intact, functional transmembrane receptors in a cellular or membrane environment. ...

Transplantation of Neonatal Mouse Cardiac Macrophages into Adult Mice

In an injured neonatal myocardium, macrophages facilitate cardiomyocyte proliferation and angiogenesis and promote heart regeneration. The present study reveals that transplantation of neonatal cardiac macrophages recruited by injury promotes adult heart regeneration after myocardial infarction with improvement of cardiac function and cardiomyocyte proliferation. The results indicate that neonatal cardiac macrophage transplantation could be a promising strategy for cardiac injury treatment. Here, we provide the technical details, including the isolation of neonatal cardiac macrophages from apical resection-injured neonatal mouse hearts, the transplantation of macrophages into myocardial-infarcted adult mice, and the estimation of heart regeneration after a macrophage graft.

We provide a protocol for neonatal cardiac macrophage separation and transplantation into an adult mouse heart, which could be a promising way to promote cardiac repair.

In an injured neonatal myocardium, macrophages facilitate cardiomyocyte proliferation and angiogenesis and promote heart regeneration. The present study ...