We acknowledge support by the School of Computing Science and Faculty of Science, Agriculture and Engineering of Newcastle University, by the Ulm University, and by the SciMedia Ltd (Tokyo, Japan).

1. Preparation of the Crab Stomatogastric Nervous System
Adult Cancer pagurus L. were obtained from local sources (Newcastle University, Dove Marine Laboratories) and kept in filtered, aerated seawater (10 - 12&deg;C). Animals were anesthetized by packing them in ice for 20 - 40 min before dissection. We used isolated stomatogastric nervous systems (STNS)1. The STNS was pinned down in a silicone elastomer-lined (ELASTOSIL RT-601, Wacker, Munich, Germany) Petri dish and continuously superfused (7 - 12 mL/min) with chilled saline (10-13&deg;C). Physiological saline consisted of (mMol*l-1): NaCl, 440; MgCl2, 26; CaCl2, 13; KCl, 11; trisma base, 10; maleic acid, 5. Saline was kept at a constant temperature of 11 - 13&deg;C and at pH 7.4 - 7.6.
The detailed steps of the STNS dissection and preparation, including the desheathing of the stomatoastric ganglion (STG), are presented in the JoVE article by Guttierez and Grashow1. All experiments were carried out in accordance with the European Communities Council Directive of 24th November 1986 (86/609/EEC). Figure 1A shows a schematic diagram of the STNS and Figure 1B shows a typical desheathed STG having its neurons arranged as a flat semicircle in the posterior part of the ganglion.
The Petri dish with the STNS is fixed to the operating platform of the imaging microscope (BW51 WI; Olympus, Tokyo, Japan) with modelling clay. Both, the microscope stage and the microscope are mounted on a vibration isolated table (Scientifica, Uckfield, UK) to prevent movement artefacts during the optical recording. The motor patterns generated in the stomatogastric ganglion (STG) are monitored using extracellular recordings2-4. This is done through the following steps:
<ol>
<li>A petroleum jelly-based cylindrical compartment is built around a section of the main motor nerve (lvn) to electrically isolate the nerve from the bath (this is done on the dissection stage before the Petri dish is placed on the operating platform of the imaging microscope).</li>
<li>One of two stainless steel electrode wires is placed in this compartment, the other one in the bath as a reference electrode.</li>
<li>The differential signal is recorded, filtered and amplified with an AC differential amplifier (Univ. Kaiserslautern, Germany).</li>
<li>The motor activity of the ganglion is monitored using an oscilloscope (DL708E; Yokogawa, Tokyo, Japan) and is additionally recorded using a data acquisition board (CED Power, 1401, Cambridge Electronic Design, Cambridge, UK) and the Spike 2 v6.10 software (Cambridge Electronic Design, Cambridge, UK). Files are analyzed using Spike 2 v6.10 (Cambridge Electronic Design, Cambridge, UK).</li>
</ol>
2. Preparation of the Dye Solution
We used the fluorescent voltage-sensitive dye Di-8-ANEPPQ (Cambridge Bioscience)5, which has a double positive charge that facilitates the loading of the dye through microelectrodes by using positive current pulses. The dye solution should be protected from light as much as possible. The dye solution is prepared as follows:
<ol>
<li>5 mg of dye is mixed with 1 mL of F-127 Pluronic acid in DMSO (Invitrogen) solution.</li>
<li>The plastic vial containing the dissolved dye is centrifuged for 20 minutes at 12,000 rotation/minute in a Sigma Microcentrifuge 1-14 (Sigma, Osterode, Germany) in order to separate larger dye particles that may clog the microelectrode.</li>
<li>The supernatant (top 200 &mu;L) of the dye solution is separated using a pipettor and stored in a separate plastic vial. Both vials are wrapped in aluminium foil to prevent exposure to light.</li>
<li>The dye solution is frozen at -20 &deg;C.</li>
<li>Before use the dye solution is melted by placing the closed and wrapped vial in warm (25 &deg;C) water for 10-15 minutes.</li>
</ol>
3. Dye Loading by Intracellular Injection using Sharp Microelectrodes
We used sharp microelectrodes to load the neurons with the dye. To check the dye loading we use the MiCAM02 imaging system (SciMedia, Tokyo, Japan)6. The imaging system has two CCD cameras: the HR camera has a larger sensor chip (6.4 mm x 4.8 mm) providing better spatial resolution with the best temporal resolution being 1.4 ms, the HS camera has a smaller (2.9 mm x 2.1 mm) but faster sensor chip allowing faster imaging having 0.7 ms as its best temporal resolution. The steps of the dye loading are as follows:
<ol>
<li>Microelectrodes are pulled with a P-97 Flaming - Brown (Sutter Instruments, Novato, CA, USA) electrode puller using 10 cm long, 1 mm outer diameter glass tubes with filament (GB100TF 8P, Science Products, Hofheim, Germany). The electrodes are pulled to have a resistance in the range of 25 - 40 MOhm in 3M KCl (for appropriate pulling parameter settings follow the pipette cookbook provided by the maker of the electrode puller).</li>
<li>The frozen dye solution is molten and a very small amount of it is sucked up into a microfil needle (MicroFil - 34G, World Precision Instruments, Sarasota, FL, USA) - the sucked up dye fills around 2/3rd of the microfil needle.</li>
<li>The dye solution in the microfil needle is injected into the tip of a microelectrode. The microelectrode is placed in a dark box for 15 minutes.</li>
<li>The microelectrode is backfilled with 3M KCl solution. The electrode is then placed back into a dark electrode storage box for another 10-20 minutes to allow the dye solution to reach the fine tip of the electrode by the suction of the capillary force acting in the microelectrode.</li>
<li>The ready microelectrode is placed into an electrode holder and fixed onto the headstage of the intracellular amplifier (IA-251A, Warner Instruments Corporation, Hamden, CT, USA).</li>
<li>The intracellular amplifier is connected to the data acquisition board that provides a pulse signal to drive current pulses into the microelectrode.</li>
<li>The microelectrode is placed gently into the membrane of an STG neuron using a microelectrode manipulator (PatchStar, Scientifica Ltd, Uckfield, UK). Since the working distance of the 10x objective (UMPLFL10XW, NA 0.30, WD 3.3mm; Olympus Corporation, Tokyo, Japan) of the imaging microscope is very small, the microelectrode should be positioned in an oblique position (around 30&deg;) and the movement of the microelectrode into the right position requires several steps of re-focusing of the microscope objective and moving the microelectrode. To start the microelectrode is moved in a position such that its tip is above the STG. The microscope objective is lowered until the electrode tip appears in focus. Then it is lowered further but not as much to touch the microelectrode. The microelectrode is lowered until it re-appears in the focus of the objective. These steps are repeated until the neurons appear clearly in the focus plane of the objective. Then the microelectrode is lowered until it touches and gently pierces the membrane of the selected STG neuron. This is monitored using the oscilloscope.</li>
<li>The intracellular electrode is used to record the activity of the selected STG neuron in order to identify the neuron given its activity profile and the relationship between its activity and the STG activity recorded from the extracellular electrode2,3,7.</li>
<li>The intracellular amplifier is switched into current injection mode and is used for 30 minutes to inject 10 nA positive current steps into the microelectrode that last for 1 second and are separated by 1 second gaps with no current injected into the microelectrode. The injected current drives the positively charged dye molecules into the recorded neuron. If equipment is available multiple neurons may be injected simultaneously.</li>
<li>Ten minutes into the injection period and after the end of the injection the dye spread within the neuron is checked. To do this we use the MiCAM02 imaging system (SciMedia Ltd, Tokyo, Japan) with the 10x objective and illumination by the ultra-low ripple halogen light source (HL-151; Moritex, Tokyo, Japan) through the filter cube MSWG2 (with 480-550 nm excitation filter; Olympus Corporation, Tokyo, Japan). The imaging system is set to use the HR camera with 192 x 128 pixel resolution, 40 ms imaging time, the 'hbin' mode, and 10 frames per imaging session. If the dye filling is successful the imaging should show a dyed patch in the neuron around the location of the microelectrode, and this dyed patch should grow between the images taken after 10 minutes and after 30 minutes. If the spread of the dye is insufficient after 30 min, more injection time is granted. Alternatively, a new dye electrode may be used.</li>
<li>The electrode is pulled out of the neuron and the neuron is left to relax for at least 20-30 minutes. Within this time, the next neuron may be injected with dye.</li>
</ol>
4. Imaging of Dyed Neurons
The activity of the single neurons can be imaged after the dye filling procedure. Alternatively two or more neurons may be filled to image the simultaneous activity of several STG neurons. In principle many and possibly all STG neurons can be filled with dye. The imaging of the neurons is done with the MiCAM 02 imaging system using the HR camera. The procedure of the imaging is as follows:
<ol>
<li>The microscope objective is changed and a high light transmission efficiency 20x objective (XLUMPLFL20XW/0.95, NA 0.95, WD 2.0mm; Olympus Corporation, Tokyo, Japan) is used for the imaging data collection. This objective collects much more light that the 10x objective and also gives higher magnification providing more detailed image of the neurons. Being more light efficient means that small amount of dye loading is already visible with this objective and also that we can use lower light intensity for the imaging reducing the potential photo damage that may be caused to the neurons by the dye.</li>
<li>The microscope's operating stage is positioned such that the neurons that are meant to be imaged are in the field of view of the objective and the objective is focused onto them.</li>
<li>The extracellular recording data is also fed into the MiCAM 02 imaging system through its analogue input channel. This allows the relatively easy relating of optical signals and corresponding motor patterns or extracellularly recorded spikes in the analysis phase.</li>
<li>The MiCAM 02 system is set to use 96 x 64 pixel resolution with 2.2 ms imaging time or 48 x 32 pixel resolution with 1.5 ms imaging time. In both cases the 'hbin' setting may be used if the dyeing of the neurons is not very strong.</li>
<li>The light level is set as low as possible (i.e. the dye in the neurons should generate still sufficient fluorescence for the imaging) to avoid photo modulation and photo damage of the recorded neurons.</li>
<li>The room lighting and all remaining light sources are turned off. In addition, a black curtain may be closed around the recording rig to prevent light exposure from external sources.</li>
<li>The imaging sessions are set to be between 4 - 16 seconds long.</li>
<li>The spontaneous activity of the dyed neurons is recorded over several imaging sessions.</li>
</ol>
5. Analysis of the Optical Imaging Data
To analyse the imaging data we used the BVAna software (version 10.08; SciMedia Ltd, Tokyo, Japan) associated with the MiCAM 02 imaging system. The software supports the visualisation of the imaging data and provides a range of analysis tools as well. The steps of the data analysis and interpretation are as follows:
<ol>
<li>An appropriate circular region of interest is selected on each recorded neurons. Typically we use 2 - 3 pixel radius regions of interest for investigating neurons at the 48 x 32 pixel resolution, and 6 - 8 pixel radius regions of interest for investigating neurons at 96 x 64 resolution.</li>
<li>The temporal smoothing of the data may be necessary, especially if the dye generates relatively low amount of fluorescence (low background value in the data indicates this; it may be due to the low light level that is used or low level dye loading, for example in a neurite of the recorded neuron). Typically in such cases we used temporal smoothing with 3 - 5 time units.</li>
<li>In addition to the optical recordings we also display the extracellular recording by selecting to display the analogue input of the imaging system.</li>
<li>The parameters for the visualisation of the optical data and the analogue input should be set appropriately in order to see all recordings on the computer screen.</li>
<li>Considering the optical recordings of the neurons, the extracellular recording from the lvn, and the classification of the neuron based on the earlier intracellular recording, it is possible to determine the identity of the neuron and to relate its activity to the neural activities recorded from the nerve.</li>
<li>It is expected that the optical recordings show clearly, and without the need of averaging over multiple similar events, the neural activities corresponding to extracellularly recorded spikes and also subthreshold events such as inhibitory post-synaptic potentials caused by the activity of other neurons.</li>
<li>The data export features of the BVAna software can be used to export the optical recordings calculated for the selected regions of interest and also the analogue input that contains the recording from the lvn. The exported data can be processed and analyzed further using other data analysis software as well (e.g. in the simplest case Microsoft Excel (Microsoft, Redmond, WA, US) may be used for additional visualisation and data processing).</li>
</ol>
6. Representative Results
Figure 2 shows the simultaneous recording of two neurons (LP, lateral pyloric neuron; PD, pyloric dilator neuron) in the pyloric central pattern generator of a crab STG together with the recording from the lvn. The optical recordings of the LP and PD neurons identified on the basis of their intracellular recordings show that indeed these match well with the extracellular recordings corresponding to LP and PD neurons. There is a consistent 12 ms delay between the optical and lvn recordings of spike peaks (see inset of Figure 2) due to the axonal transmission delay between the ganglion and the extracellular recording site. The presented data also shows that the neural activity in the soma that corresponds to spikes of the recorded neurons is clearly detectable.
<img src="/files/ftp_upload/2567/2567fig1.jpg" alt="Figure 1" /> 
Figure 1. A) Schematic diagram of the stomatogastric nervous system (STNS). CoG, commissural ganglion; OG, esophageal ganglion; STG, stomatogastric ganglion; stn, stomatogastric nerve; lvn, lateral ventricular nerve. B) The crab stomatogastric ganglion (STG) - the image shows the semicircular posterior arrangement of STG neurons.
<img src="/files/ftp_upload/2567/2567fig2.jpg" alt="Figure 2" /> 
Figure 2. Simultaneous single-sweep recording of a PD and an LP neuron together with the recording of the lvn. The data shows that that the optical and nerve recordings match very well and that we can identify the neural activities corresponding to spikes recorded form the nerve. The inset shows an overlay of several sweeps of LP action potentials - both optically and electrically recorded - and their average demonstrating the match between optically and electrically recorded activities. Due to the fact that our optical recording method does not filter out the low frequency components of the data we are also able to record slow membrane potential changes characteristic for PD and LP neurons14.

<ol>
	<li>Gutierrez, G. J., Grashow, R. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cancer+borealis+stomatogastric+nervous+system+dissection.">Cancer borealis stomatogastric nervous system dissection.</a> J Vis Exp. , (2009).</li><li>Bartos, M., Nusbaum, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intercircuit+control+of+motor+pattern+modulation+by+presynaptic+inhibition.">Intercircuit control of motor pattern modulation by presynaptic inhibition.</a> J. Neurosci. 17, 2247-2256 (1997).</li><li>Blitz, D. M., Nusbaum, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Motor+pattern+selection+via+inhibition+of+parallel+pathways.">Motor pattern selection via inhibition of parallel pathways.</a> J Neurosci. 17, 4965-4975 (1997).</li><li>Stein, W., Smarandache, C. R., Nickmann, M., Hedrich, U. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Functional+consequences+of+activity-dependent+synaptic+enhancement+at+a+crustacean+neuromuscular+junction.">Functional consequences of activity-dependent synaptic enhancement at a crustacean neuromuscular junction.</a> J Exp Biol. 209, 1285-12300 (2006).</li><li>Loew, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Potentiometric+dyes:+Imaging+electrical+activity+of+cell+membranes.">Potentiometric dyes: Imaging electrical activity of cell membranes.</a> Pure &#38; Appl Chem. 68, 1405-1409 (1996).</li><li>Tominaga, T., Tominaga, Y., Yamada, H., Matsumoto, G., Ichikawa, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantification+of+optical+signals+with+electrophysiological+signals+in+neural+activities+of+Di-4-ANEPPS+stained+rat+hippocampal+slices.">Quantification of optical signals with electrophysiological signals in neural activities of Di-4-ANEPPS stained rat hippocampal slices.</a> J Neurosci Meth. 102, 11-23 (2000).</li><li>Heinzel, H. 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No conflicts of interest declared.

The voltage sensitive dye imaging8,9 of STG neurons in combination with traditional electrophysiology methods (intra- and extracellular electrode recordings) allows an improved understanding of how this small, well-known and still complex neural system works. The STG is a prototype of central pattern generator (CPG) neuronal networks (it includes the pyloric and gastric mill rhythm networks)10,,11, thus a better understanding of the emergent functional properties of the STG will also help to understand how in general CPG networks generate their network level functionality. CPG networks play a critical role in motor control12 and also in higher cognitive functions13, consequently the better understanding of their emergent properties may have a major impact in many areas of neuroscience.

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>MiCAM02</td>
<td>&nbsp;</td>
<td>SciMedia Ltd, Tokyo, Japan</td>
<td>&nbsp;</td>
<td>Imaging system</td>
</tr>
<tr>
<td>BVAna v.10.08</td>
<td>&nbsp;</td>
<td>SciMedia, Tokyo, Japab</td>
<td>&nbsp;</td>
<td>Data analysis software for the MiCAM 02 imaging system</td>
</tr>
<tr>
<td>HL-151</td>
<td>&nbsp;</td>
<td>Moritex, Tokyo, Japan</td>
<td>&nbsp;</td>
<td>Ultra-low ripple halogen light source</td>
</tr>
<tr>
<td>BX51 WI</td>
<td>&nbsp;</td>
<td>Olympus, Tokyo, Japan</td>
<td>&nbsp;</td>
<td>Upright microscope for epifluorescence imaging</td>
</tr>
<tr>
<td>20x imaging objective </td>
<td>&nbsp;</td>
<td>Olympus Corporation, Tokyo, Japan</td>
<td>XLUMPLFL20XW/0.95</td>
<td>NA 0.95, WD 2.0mm</td>
</tr>
<tr>
<td>10x imaging objective </td>
<td>&nbsp;</td>
<td>Olympus Corporation, Tokyo, Japan</td>
<td>UMPLFL10XW</td>
<td>NA 0.30, WD 3.3mm</td>
</tr>
<tr>
<td>Filter cube with 480-550 nm excitation filter and 590nm emission filter </td>
<td>&nbsp;</td>
<td>Olympus Corporation, Tokyo, Japan</td>
<td>MSWG2</td>
<td>&nbsp;</td>
<tr>
<td>Antivibration table</td>
<td>&nbsp;</td>
<td>Scientifica Ltd, Uckfield, UK</td>
<td>63-534</td>
<td>&nbsp;</td>
<tr>
<td>PatchStar electrode manipulator</td>
<td>&nbsp;</td>
<td>Scientifica Ltd, Uckfield, UK</td>
<td>PS-7000C</td>
<td>Micro-electrode manipulator</td>
</tr>
<tr>
<td>CED Power 1401</td>
<td>&nbsp;</td>
<td>Cambridge Electronic Design, Cambridge, UK</td>
<td>&nbsp;</td>
<td>Data acquisition board</td>
</tr>
<tr>
<td>DL708E Oscilloscope</td>
<td>&nbsp;</td>
<td>Yokogawa, Tokyo, Japan</td>
<td>&nbsp;</td>
<td>Intracellular amplifier</td>
</tr>
<tr>
<td>AC differential amplifier</td>
<td>&nbsp;</td>
<td>University of Kaiserslautern, Germany</td>
<td>&nbsp;</td>
<td>Extracellular amplifier</td>
</tr>
<tr>
<td>P-97 Flaming &#x2013; Brown type electrode puller</td>
<td>&nbsp;</td>
<td>Sutter Instruments, Novato, CA, USA</td>
<td>&nbsp;</td>
<td>Electrode puller</td>
</tr>
<tr>
<td>Sigma Microcentrifuge 1-14</td>
<td>&nbsp;</td>
<td>Sigma, Osterode, Germany</td>
<td>&nbsp;</td>
<td>Centrifuge</td>
</tr>
<tr>
<td>Genex Pipettors Mline</td>
<td>&nbsp;</td>
<td>Scientific Laboratory Supplies, UK</td>
<td>PIP7774</td>
<td>PIP7774</td>
</tr>
<tr>
<td>Glass micropipette with filament</td>
<td>&nbsp;</td>
<td>Science Products, Hofheim, Germany</td>
<td>GB100TF 8P</td>
<td>Glass for microelectrodes</td>
</tr>
<tr>
<td>MicroFil &#x2013; 34G</td>
<td>&nbsp;</td>
<td>World Precision Instruments, Sarasota, FL, USA</td>
<td>&nbsp;</td>
<td>Microfil needle</td>
</tr>
<tr>
<td>Spike 2 v6.10</td>
<td>&nbsp;</td>
<td>Cambridge Electronic Design, Cambridge, UK</td>
<td>&nbsp;</td>
<td>Electrophysiology software</td>
</tr>
<tr>
<td>Di-8-ANEPPQ</td>
<td>&nbsp;</td>
<td>Cambridge Bioscience Ltd, Cambridge, UK</td>
<td>BT61014</td>
<td>Fluorescent voltage sensitive dye</td>
</tr>
<tr>
<td>F-127 Pluronic Acid, DMSO</td>
<td>&nbsp;</td>
<td>Invitrogen, Paisley, UK</td>
<td>P-3000MP</td>
<td>Solvent</td>
</tr>
<tr>
<td>NaCl</td>
<td>&nbsp;</td>
<td>Sigma &#x2013; Aldrich, Poole, UK</td>
<td>S9888</td>
<td>Crab saline component</td>
</tr>
<tr>
<td>CaCl2 . 2H2O</td>
<td>&nbsp;</td>
<td>Sigma &#x2013; Aldrich, Poole, UK</td>
<td>C3881</td>
<td>Crab saline component</td>
</tr>
<tr>
<td>KCl</td>
<td>&nbsp;</td>
<td>Sigma &#x2013; Aldrich, Poole, UK</td>
<td>P3911</td>
<td>Crab saline component, electrolyte solution</td>
</tr>
<tr>
<td>MgCl2 . 6H2O</td>
<td>&nbsp;</td>
<td>Sigma &#x2013; Aldrich, Poole, UK</td>
<td>M9272</td>
<td>Crab saline component</td>
</tr>
<tr>
<td>Trizma base</td>
<td>&nbsp;</td>
<td>Sigma &#x2013; Aldrich, Poole, UK</td>
<td>T1503</td>
<td>Crab saline component</td>
</tr>
<tr>
<td>Maleic acid</td>
<td>&nbsp;</td>
<td>Sigma &#x2013; Aldrich, Poole, UK</td>
<td>M0375</td>
<td>Crab saline component</td>
</tr>
<tr>
<td>Elastosil RT-601</td>
<td>&nbsp;</td>
<td>Wacker, Munich, Germany</td>
<td>&nbsp;</td>
<td>Lining of Petri dishes</td>
</tr>		</tbody>
		</table>
		</div>

optical imaging of neurons in the crab stomatogastric ganglion with voltage-sensitive dyes

Voltage-sensitive dye imaging of neurons is a key methodology for the understanding of how neuronal networks are organised and how the simultaneous activity of participating neurons leads to the emergence of the integral functionality of the network. Here we present the methodology of application of this technique to identified pattern generating neurons in the crab stomatogastric ganglion. We demonstrate the loading of these neurons with the fluorescent voltage-sensitive dye Di-8-ANEPPQ and we show how to image the activity of dye loaded neurons using the MiCAM02 high speed and high resolution CCD camera imaging system. We demonstrate the analysis of the recorded imaging data using the BVAna imaging software associated with the MiCAM02 imaging system. The simultaneous voltage-sensitive dye imaging of the detailed activity of multiple neurons in the crab stomatogastric ganglion applied together with traditional electrophysiology techniques (intracellular and extracellular recordings) opens radically new opportunities for the understanding of how central pattern generator neural networks work.

Watch this Scientific Journal Video about Optical Imaging of Neurons in the Crab Stomatogastric Ganglion with Voltage-sensitive Dyes at JoVE.com

Optical Imaging of Neurons in the Crab Stomatogastric Ganglion with Voltage-sensitive Dyes

Here we present the methodology for fast and high resolution fluorescent voltage-sensitive dye imaging of detailed activity of neurons in the crab stomatogastric ganglion.

Voltage-sensitive dye imaging of neurons is a key methodology for the understanding of how neuronal networks are organised and how the simultaneous ...

optical-imaging-neurons-crab-stomatogastric-ganglion-with-voltage

Nanyang Technological University

Research

JoVE Journal

Neuroscience

12.9K Views.  Ulm University. Here we present the methodology for fast and high resolution fluorescent voltage-sensitive dye imaging of detailed activity of neurons in the crab stomatogastric ganglion.

Video: Optical Imaging of Neurons in the Crab Stomatogastric Ganglion with Voltage-sensitive Dyes

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured neurons for extended durations1-3. This is now possible through the use of vital dyes and fluorescent proteins with which cytoskeletal components, organelles, and other structures in living cells can be labeled and then visualized via dynamic fluorescence microscopy. For example, in embryonic chicken sympathetic neurons, mitochondrial movement was characterized using the vital dye rhodamine 1234. In another study, mitochondria were visualized in rat forebrain neurons by transfection of mitochondrially targeted eYFP5. However, imaging of primary neurons over minutes, hours, or even days presents a number of issues. Foremost among these are: 1) maintenance of culture conditions such as temperature, humidity, and pH during long imaging sessions; 2) a strong, stable fluorescent signal to assure both the quality of acquired images and accurate measurement of signal intensity during image analysis; and 3) limiting exposure times during image acquisition to minimize photobleaching and avoid phototoxicity.
Here, we describe a protocol that permits the observation, visualization, and analysis of mitochondrial movement in cultured hippocampal neurons with high temporal resolution and under optimal life support conditions. We have constructed an affordable stage-top incubator that provides good temperature regulation and atmospheric gas flow, and also limits the degree of media evaporation, assuring stable pH and osmolarity. This incubator is connected, via inlet and outlet hoses, to a standard tissue culture incubator, which provides constant humidity levels and an atmosphere of 5-10% CO2/air. This design offers a cost-effective alternative to significantly more expensive microscope incubators that don't necessarily assure the viability of cells over many hours or even days. To visualize mitochondria, we infect cells with a lentivirus encoding a red fluorescent protein that is targeted to the mitochondrion. This assures a strong and persistent signal, which, in conjunction with the use of a stable xenon light source, allows us to limit exposure times during image acquisition and all but precludes photobleaching and phototoxicity. Two injection ports on the top of the stage-top incubator allow the acute administration of neurotransmitters and other reagents intended to modulate mitochondrial movement. In sum, lentivirus-mediated expression of an organelle-targeted red fluorescent protein and the combination of our stage-top incubator, a conventional inverted fluorescence microscope, CCD camera, and xenon light source allow us to acquire time-lapse images of mitochondrial transport in living neurons over longer durations than those possible in studies deploying conventional vital dyes and off-the-shelf life support systems.

We describe a protocol that allows imaging of mitochondria in living neurons via fluorescence microscopy over long durations. Imaging over extended periods is accomplished through lentivirus-mediated expression of a mitochondrially targeted fluorescent protein and use of an inexpensive stage-top incubator that was designed and built in our laboratory.

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured ...

Genetic Study of Axon Regeneration with Cultured Adult Dorsal Root Ganglion Neurons

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability to support axon growth and a hostile environment in the mature CNS1,2. In contrast, mature neurons in the peripheral nervous system (PNS) regenerate readily after injuries3. Adult dorsal root ganglion (DRG) neurons are well known to regenerate robustly after peripheral nerve injuries. Each DRG neuron grows one axon from the cell soma, which branches into two axonal branches: a peripheral branch innervating peripheral targets and a central branch extending into the spinal cord. Injury of the DRG peripheral axons results in substantial axon regeneration, whereas central axons in the spinal cord regenerate poorly after the injury. However, if the peripheral axonal injury occurs prior to the spinal cord injury (a process called the conditioning lesion), regeneration of central axons is greatly improved4. Moreover, the central axons of DRG neurons share the same hostile environment as descending corticospinal axons in the spinal cord. Together, it is hypothesized that the molecular mechanisms controlling axon regeneration of adult DRG neurons can be harnessed to enhance CNS axon regeneration. As a result, adult DRG neurons are now widely used as a model system to study regenerative axon growth5-7.
Here we describe a method of adult DRG neuron culture that can be used for genetic study of axon regeneration in vitro. In this model adult DRG neurons are genetically manipulated via electroporation-mediated gene transfection6,8. By transfecting neurons with DNA plasmid or si/shRNA, this approach enables both gain- and loss-of-function experiments to investigate the role of any gene-of-interest in axon growth from adult DRG neurons. When neurons are transfected with si/shRNA, the targeted endogenous protein is usually depleted after 3-4 days in culture, during which time robust axon growth has already occurred, making the loss-of-function studies less effective. To solve this problem, the method described here includes a re-suspension and re-plating step after transfection, which allows axons to re-grow from neurons in the absence of the targeted protein. Finally, we provide an example of using this in vitro model to study the role of an axon regeneration-associated gene, c-Jun, in mediating axon growth from adult DRG neurons9.

An in vitro model for genetic study of axon regeneration using cultured adult mouse dorsal root ganglion neurons is described. The method includes a re-suspension/re-plating step to allow axon re-growth from neurons undergoing genetic manipulation. This approach is especially useful for loss-of-function studies of axon regeneration using RNAi-based protein knockdown.

It is well known that mature neurons in the central nervous system (CNS) cannot regenerate their axons after injuries due to diminished intrinsic ability ...

Voltage-sensitive Dye Recording from Axons, Dendrites and Dendritic Spines of Individual Neurons in Brain Slices

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding how the brain works. The primary function of any nerve cell is to process electrical signals, usually from multiple sources. Electrical properties of neuronal processes are extraordinarily complex, dynamic, and, in the general case, impossible to predict in the absence of detailed measurements. To obtain such a measurement one would, ideally, like to be able to monitor, at multiple sites, subthreshold events as they travel from the sites of origin on neuronal processes and summate at particular locations to influence action potential initiation. This goal has not been achieved in any neuron due to technical limitations of measurements that employ electrodes. To overcome this drawback, it is highly desirable to complement the patch-electrode approach with imaging techniques that permit extensive parallel recordings from all parts of a neuron. Here, we describe such a technique - optical recording of membrane potential transients with organic voltage-sensitive dyes (Vm-imaging) - characterized by sub-millisecond and sub-micrometer resolution. Our method is based on pioneering work on voltage-sensitive molecular probes 2. Many aspects of the initial technology have been continuously improved over several decades 3, 5, 11. Additionally, previous work documented two essential characteristics of Vm-imaging. Firstly, fluorescence signals are linearly proportional to membrane potential over the entire physiological range (-100 mV to +100 mV; 10, 14, 16). Secondly, loading neurons with the voltage-sensitive dye used here (JPW 3028) does not have detectable pharmacological effects. The recorded broadening of the spike during dye loading is completely reversible 4, 7. Additionally, experimental evidence shows that it is possible to obtain a significant number (up to hundreds) of recordings prior to any detectable phototoxic effects 4, 6, 12, 13. At present, we take advantage of the superb brightness and stability of a laser light source at near-optimal wavelength to maximize the sensitivity of the Vm-imaging technique. The current sensitivity permits multiple site optical recordings of Vm transients from all parts of a neuron, including axons and axon collaterals, terminal dendritic branches, and individual dendritic spines. The acquired information on signal interactions can be analyzed quantitatively as well as directly visualized in the form of a movie.

An imaging technique for monitoring of membrane potential changes with sub-micrometer spatial and sub-millisecond temporal resolution is described. The technique, based on laser excitation of voltage-sensitive dyes, allows measurements of signals in axons and axon collaterals, terminal dendritic branches, and individual dendritic spines.

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding ...

Implementing Dynamic Clamp with Synaptic and Artificial Conductances in Mouse Retinal Ganglion Cells

Ganglion cells are the output neurons of the retina and their activity reflects the integration of multiple synaptic inputs arising from specific neural circuits. Patch clamp techniques, in voltage clamp and current clamp configurations, are commonly used to study the physiological properties of neurons and to characterize their synaptic inputs. Although the application of these techniques is highly informative, they pose various limitations. For example, it is difficult to quantify how the precise interactions of excitatory and inhibitory inputs determine response output. To address this issue, we used a modified current clamp technique, dynamic clamp, also called conductance clamp 1, 2, 3 and examined the impact of excitatory and inhibitory synaptic inputs on neuronal excitability. This technique requires the injection of current into the cell and is dependent on the real-time feedback of its membrane potential at that time. The injected current is calculated from predetermined excitatory and inhibitory synaptic conductances, their reversal potentials and the cell's instantaneous membrane potential. Details on the experimental procedures, patch clamping cells to achieve a whole-cell configuration and employment of the dynamic clamp technique are illustrated in this video article. Here, we show the responses of mouse retinal ganglion cells to various conductance waveforms obtained from physiological experiments in control conditions or in the presence of drugs. Furthermore, we show the use of artificial excitatory and inhibitory conductances generated using alpha functions to investigate the responses of the cells.

This video article illustrates the set-up, the procedures to patch cell bodies and how to implement dynamic clamp recordings from ganglion cells in whole-mount mouse retinae. This technique allows the investigation of the precise contribution of excitatory and inhibitory synaptic inputs, and their relative magnitude and timing to neuronal spiking.

Ganglion cells are the output neurons of the  retina and their activity reflects the integration of multiple synaptic inputs  arising from specific neural ...

Retrograde Labeling of Retinal Ganglion Cells in Adult Zebrafish with Fluorescent Dyes

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs survival and regeneration experiments. Many studies have been performed in mammalian animals to research RGCs survival after optic nerve injury. However, retrograde labeling of RGCs in adult zebrafish has not yet been reported, though some alternative methods can count cell numbers in retinal ganglion cell layers (RGCL). Considering the small size of the adult zebrafish skull and the high risk of death after drilling on the skull, we open the skull with the help of acid-etching and seal the hole with a light curing bond, which could significantly improve the survival rate. After absorbing the dyes for 5 days, almost all the RGCs are labeled. As this method does not need to transect the optic nerve, it is irreplaceable in the research of RGCs survival after optic nerve crush in adult zebrafish. Here, we introduce this method step by step and provide representative results.

We introduce an efficient method to retrograde label retinal ganglion cells (RGCs) in adult zebrafish.

As retrograde labeling retinal ganglion cells (RGCs) can isolate RGCs somata from dying sites, it has become the gold standard for counting RGCs in RGCs ...

Optical Control of a Neuronal Protein Using a Genetically Encoded Unnatural Amino Acid in Neurons

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to photo-regulate endogenous proteins expressed in their native environment. Here, we present an approach to optically control the function of a neuronal protein directly in neurons using a genetically encoded unnatural amino acid (Uaa). By using an orthogonal tRNA/aminoacyl-tRNA synthetase pair to suppress the amber codon, a photo-reactive Uaa 4,5-dimethoxy-2-nitrobenzyl-cysteine (Cmn) is site-specifically incorporated in the pore of a neuronal protein Kir2.1, an inwardly rectifying potassium channel. The bulky Cmn physically blocks the channel pore, rendering Kir2.1 non-conducting. Light illumination instantaneously converts Cmn into a smaller natural amino acid Cys, activating Kir2.1 channel function. We express these photo-inducible inwardly rectifying potassium (PIRK) channels in rat hippocampal primary neurons, and demonstrate that light-activation of PIRK ceases the neuronal firing due to the outflux of K+ current through the activated Kir2.1 channels. Using in utero electroporation, we also express PIRK in the embryonic mouse neocortex in vivo, showing the light-activation of PIRK in neocortical neurons. Genetically encoding Uaa imposes no restrictions on target protein type or cellular location, and a family of photoreactive Uaas is available for modulating different natural amino acid residues. This technique thus has the potential to be generally applied to many neuronal proteins to achieve optical regulation of different processes in brains. The current protocol presents an accessible procedure for intricate Uaa incorporation in neurons in vitro and in vivo to achieve photo control of neuronal protein activity on the molecular level.

Here, a procedure to selectively activate a neuronal protein with a short pulse of light by genetically encoding a photo-reactive unnatural amino acid into a target neuronal protein expressed in neurons in culture or in vivo is presented.

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to ...

Optical Coherence Tomography: Imaging Mouse Retinal Ganglion Cells In Vivo

Structural changes in the retina are common manifestations of ophthalmic diseases. Optical coherence tomography (OCT) enables their identification in vivo&#8212;rapidly, repetitively, and at a high resolution. This protocol describes OCT imaging in the mouse retina as a powerful tool to study optic neuropathies (OPN). The OCT system is an interferometry-based, non-invasive alternative to common post mortem histological assays. It provides a fast and accurate assessment of retinal thickness, allowing the possibility to track changes, such as retinal thinning or thickening. We present the imaging process and analysis with the example of the Opa1delTTAG mouse line. Three types of scans are proposed, with two quantification methods: standard and homemade calipers. The latter is best for use on the peripapillary retina during radial scans; being more precise, is preferable for analyzing thinner structures. All approaches described here are designed for retinal ganglion cells (RGC) but are easily adaptable to other cell populations. In conclusion, OCT is efficient in mouse model phenotyping and has the potential to be used for the reliable evaluation of therapeutic interventions.

Optical Coherence Tomography: Imaging Mouse Retinal Ganglion Cells In Vivo

This manuscript describes a protocol for the in vivo imaging of the mouse retina with high-resolution spectral domain optical coherence tomography (SD-OCT). It focuses on retinal ganglion cells (RGC) in the peripapillary region, with several scanning and quantifying approaches described.

Structural changes in the retina are common manifestations of ophthalmic diseases. Optical coherence tomography (OCT) enables their identification in ...

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging methods exist for examining the brain tissue of live newborn mammals. Herein we summarize a protocol for imaging individual cortical neurons in living neonatal mice. This protocol includes the following two methodologies: (1) the Supernova system for sparse and bright labeling of cortical neurons in the developing brain, and (2) a surgical procedure for the fragile neonatal skull. This protocol allows the observation of temporal changes of individual cortical neurites during neonatal stages with a high signal-to-noise ratio. Labeled cell-specific gene silencing and knockout can also be achieved by combining the Supernova with RNA interference and CRISPR/Cas9 gene editing systems. This protocol can, thus, be used for analyzing the developmental dynamics of cortical neurons, molecular mechanisms that control the neuronal dynamics, and changes in neuronal dynamics in disease models.

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

We present an in vivo two-photon imaging protocol for imaging the cerebral cortex of neonatal mice. This method is suitable for analyzing the developmental dynamics of cortical neurons, the molecular mechanisms that control the neuronal dynamics, and the changes in neuronal dynamics in disease models.

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging ...

Wide-field Single-photon Optical Recording in Brain Slices Using Voltage-sensitive Dye

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural circuits. Due to the fractional change in the light signal, it has been difficult to use this method as a quantitative assay. This article describes special optics and slice handling systems, which render this technique stable and reliable. The present article demonstrates the slice handling, staining, and recording of the VSD-stained hippocampal slices in detail. The system maintains physiological conditions for a long time, with good staining, and prevents mechanical movements of the slice during the recordings. Moreover, it enables staining of slices with a small amount of the dye. The optics achieve high numerical aperture at low magnification, which allows recording of the VSD signal at the maximum frame rate of 10 kHz, with 100 pixel x 100-pixel spatial resolution. Due to the high frame rate and spatial resolution, this technique allows application of the post-recording filters that provide sufficient signal-to-noise ratio to assess the changes in neural circuits.

We introduce a reproducible and stable optical recording method for brain slices using voltage-sensitive dye. The article describes voltage-sensitive dye staining and recording of optical signals using conventional hippocampal slice preparations.

Wide-field single photon voltage-sensitive dye (VSD) imaging of brain slice preparations is a useful tool to assess the functional connectivity in neural ...

Advanced Diffusion Imaging in The Hippocampus of Rats with Mild Traumatic Brain Injury

Mild traumatic brain injury (mTBI) is the most common type of acquired brain injury. Since patients with traumatic brain injury show a tremendous variability and heterogeneity (age, gender, type of trauma, other possible pathologies, etc.), animal models play a key role in unraveling factors that are limitations in clinical research. They provide a standardized and controlled setting to investigate the biological mechanisms of injury and repair following TBI. However, not all animal models mimic the diffuse and subtle nature of mTBI effectively. For example, the commonly used controlled cortical impact (CCI) and lateral fluid percussion injury (LFPI) models make use of a craniotomy to expose the brain and induce widespread focal trauma, which are not commonly seen in mTBI. Therefore, these experimental models are not valid to mimic mTBI. Thus, an appropriate model should be used to investigate mTBI. The Marmarou weight drop model for rats induces similar microstructural alterations and cognitive impairments as seen in patients who sustain mild trauma; therefore, this model was selected for this protocol. Conventional computed tomography and magnetic resonance imaging (MRI) scans commonly show no damage following a mild injury, because mTBI induces often only subtle and diffuse injuries. With diffusion weighted MRI, it is possible to investigate microstructural properties of brain tissue, which can provide more insight into the microscopic alterations following mild trauma. Therefore, the goal of this study is to obtain quantitative information of a selected region-of-interest (i.e., hippocampus) to follow up disease progression after obtaining a mild and diffuse brain injury.

The overall goal of this procedure is to obtain quantitative microstructural information of the hippocampus in a rat with mild traumatic brain injury. This is done using an advanced diffusion-weighted magnetic resonance imaging protocol and region-of-interest based analysis of parametric diffusion maps.