We are grateful to our collaborators Knut Holthoff, Arthur Konnerth and Marco Canepari who participated in the initial development of this technique as well as to Leslie M. Loew for kindly providing dyes. Supported by NSF grant IOS-0817969, NIH grants NS068407 and M136043 and by Kavli Institute for Neuroscience at Yale University.

1. Equipment Setup
Step 1.1. Imaging setup
The key to recording voltage sensitive dye signals is appropriate setup design. We use an upright microscope (BX51WI Olympus or Zeiss AxioExaminer) equipped with three cameras. The setup is designed for illuminating individual neurons in brain slices by excitation light in epi-fluorescence, wide-field microscopy mode using either Nikon 60X/1.0 NA or Zeiss 63X/1.0 NA water dipping objectives. Our microscopes are bolted to a vibration isolation table and equipped with motorized movable stages. Each microscope is equipped with three camera ports. One camera port has a standard high spatial resolution CCD camera for infrared DIC video-microscopy (IR-1000, Dage MTI, USA). The second camera port has a fast data acquisition camera (up to 20 kHz frame rate) with relatively low spatial resolution (80 x 80 pixels) but outstanding dynamic range (14 bits) and exceptionally low read noise (NeuroCCD-SM, RedShirtImaging LLC, Decatur, GA). The third camera port has a CCD camera with high spatial resolution (PixelFly, 1392x1024 pixels; PCO AG, Germany) mounted on a spinning-disc confocal scanner (CSU-10, Yokogawa, Japan) used to collect z-stacks of confocal images for detailed morphological reconstruction of the stained cell. A frequency-doubled diode-pumped Nd:YVO4 continuous wave laser (400 mW) emitting at 532 nm (MLL-III/400 mW; CNI, Changchun, China) is the source of excitation light. The 2 mm diameter laser beam gated by a shutter (LS6; Vincent Associates) is directed to a light guide coupled to the microscope via a single-port epifluorescence condenser (TILL Photonics GmbH, Gr&auml;felfing, Germany) designed to overfill the back aperture of the objective and provide near uniform illumination of the object plane. The laser light is used in place of a conventional Xenon arc-lamp to maximize the sensitivity of Vm-imaging by: (1) using a monochromatic excitation light at the red wing of the absorption spectrum to maximize Vm sensitivity of the dye 9, 10 and (2) increasing the intensity of the excitation light beyond the level that can be achieved by an arc-lamp. The excitation light was reflected to the preparation by a dichroic mirror with a central wavelength of 560 nm and the recorded fluorescence light was passed through a 610 nm barrier filter (a Schott RG610). The combined effect of an increase in light intensity and the use of near optimal monochromatic excitation wavelengths was a dramatic improvement in the sensitivity of voltage imaging by a factor of about 50 compared to previous measurements 6.
Step 1.2. Adjust for uniform illumination
Use a fluorescence slide standard (green excitation/red emission). Insert appropriate neutral density filters in the laser beam path so that the CCD is not saturated. Focus the objective on the surface of the slide. Adjust the position of the receiving end of the quartz light guide in front of the laser launcher objective, and adjust the position of the output end of the light guide attached to the microscope using appropriate actuators on the epi-fluorescence condenser to achieve centered and uniform illumination of the field of view.
Step 1.3. Determine vibrational noise
Place a small black ink mark on the surface of the fluorescence slide. Record light intensity with NeuroCCD in the continuous recording mode. Focus the objective on a dark edge of the black ink mark. Record the light intensity for about 100 msec at 2 kHz frame rate. Display the spatial average of the fractional light intensity traces (&Delta;F/F) from ~ 20 pixels receiving light from the uniformly illuminated area and from ~20 pixels along the edge of the ink mark. The excess noise in recordings from pixels with high contrast edges reflects the vibrational noise in the system.
Step 1.4. Vibration isolation
It is mandatory to reduce vibrations by using a vibration-isolation table to a level below the shot noise at light intensities comparable to experimental recording conditions. Adjust the vibration isolation table until the vibration noise in the light intensity from pixels covering the sharp edge of the image is negligible. None of the equipment with moving parts (mechanical shutters, fans) can be mounted on the table. The cables from the equipment on the table attached to other components that are not isolated from vibration must be loose so that they do not transmit mechanical vibrations to the microscope.
2. Selection of an Appropriate Neuron for Vm-Imaging
Step 2.1. Selection of neurons
Make brain slices according to standard procedures. Use a mouse line expressing EGFP in individual nerve cells of interest. With a spinning disc confocal system, visualize EGFP labeled neurons in the slice. Select neurons for Vm-imaging with intact dendritic/axonal trees, and with processes running parallel and close to the surface of the slice. This cannot be accomplished in wild-type mice because axons and thin dendrites, for the most part, are not visible in DIC microscopy mode.
3. Loading Neurons with Voltage-sensitive Dye
Step 3.1. Filling of the patch pipette
Fill glass patch pipettes from the tip with dye-free intracellular solution by applying negative pressure for about 15 s up to 2/3 of the electrode taper. The dye free solution in the tip is necessary to prevent the spill of dye onto the slice which increases the background fluorescence and dramatically reduces signal to noise ratio. Back-fill the electrode with the solution containing the membrane impermeant voltage sensitive dye JPW3028 dissolved in intracellular solution (0.8 mM).
JPW3028, the most successful voltage probe for intracellular application, is a doubly positively charged analogue of the ANEP series of lipophilic styryl dyes that is still sufficiently water soluble to be used for microinjection. The di-ethyl analog of this dye has practically identical characteristics (including voltage-sensitivity) and is commercially available as JPW1114 (see Table 1). We prepare 20 mM stock solution in distilled H2O. 50 &mu;l aliquots of the stock solution are kept frozen at -20 &deg;C. For the final dye concentration of 0.8 mM, 2 &mu;l of the stock solution is dissolved in 50 &mu;l of intracellular solution on the day of the experiment. The stock dye solution is stable and can be kept at room temperature for several months. Thus, we keep one 50 &mu;l aliquot at room temperature until it is used up.
Step 3.2. Establish a giga-seal quickly
Patch a previously selected neuron and allow free diffusion of the dye from a somatic patch pipette to the soma in the whole-cell configuration for 20-45 min. It is essential to obtain a seal as fast as possible so that the amount of dye-free solution in the tip can be kept small. Start by practicing fast patching without the dye in the electrode. Next, practice patching and loading neurons with the dye with the goal of using a minimal amount of dye-free solution in the tip without contaminating the surrounding tissue. With practice, it is possible to establish a seal within 1-2 min.
Step 3.3. Monitor the level of staining
During dye diffusion in the whole-cell configuration, monitor the physiological state of the neuron by recording evoked action potentials in the current-clamp mode. Additionally, monitor the amount of staining by recording the resting light intensity (RLI) from the cell soma at a frame rate of 2 kHz and at a fraction of full light intensity adjusted with neutral density filters (we use 0.04% of the laser light intensity from a 400 mW laser). Continue the dye loading process until the action potential starts to broaden, usually after 20-40 minutes, depending on the electrode size and resistance.
The broadening of the spike is completely reversible and most likely due to capacitive load effect of the saturated concentration of the dye in the somatic membrane. The waveform of the spike is fully restored after the dye concentration is equilibrated throughout the neuron.
Step 3.4. Remove the dye electrode
At the end of staining period, carefully pull the patch pipette away from the soma in voltage clamp configuration ensuring that the transition from whole-cell to outside-out patch configuration is attained in the process.
Step 3.5. Wait for dye diffusion
Incubate the slice for an additional 1.5-2 hr at room temperature to allow for the voltage-sensitive dye to spread into neuronal processes. After a significant amount of dye diffuses away from the soma into dendritic and axonal processes, the waveform of the AP is completely restored.
4. Optical Recording
Step 4.1. Select a cellular compartment for imaging
Locate the soma of the stained neuron under low light level fluorescence and re-patch the neuron with a standard (no dye) patch electrode under DIC. Visualize neuronal processes under low light level at a frame rate of 10-40 Hz in the continuous recording mode of the CCD for voltage-imaging. Reduce the light level with neutral density filters to the minimum required to visualize the object of interest. We use 0.01% of the 400 mW laser intensity during positioning of the stained neuron. Using an X-Y stage, position the neuronal process of interest in the middle of the imaging area. Protect the soma from the excitation light using the partially closed field stop iris of the microscope. Shielding the soma from high intensity excitation light will significantly reduce the photodynamic damage during recording.
Step 4.2 Record optical signals related to membrane potential transients
Record optical signals related to backpropagated APs in individual dendritic branches. Record optical signals related to APs in the axon. Record optical signals related to backpropagating APs in dendritic spines. Use frame rates appropriate for accurate reconstruction of signal waveform and keep the recording periods and exposure to high intensity excitation light as short as possible to minimize dye bleaching and photodynamic damage. For example, examining the sequence of initiation and propagation of single action potentials in the axon requires recording periods of 5-10 msec. The excitation light intensity during recording is a compromise between the signal-to-noise ratio on the one hand and the degree of dye bleaching and photodynamic damage on the other hand. We use 100% of intensity of the 400 mW laser in recording from long sections of axons when an area of 300 &mu;m in diameter is illuminated. When the excitation light is focused to a 30 &mu;m diameter area for dendritic spine imaging, we use 10-25% of the laser light intensity. The required duration of recording is an important determinant of the optimal excitation light intensity; shorter recording periods allow higher light intensities. The optimal illumination intensity is best determined empirically for each preparation and measurement settings.
5. Data Analysis
Step 5.1. Correct the raw data for known errors
The analysis and display of data were carried out using the NeuroPlex program (RedShirtImaging) written in IDL (ITT Visual Information Solutions, Boulder, Colorado) and custom Visual Basic routines. Under the conditions of low light levels, the background fluorescence becomes a significant determinant of the &Delta;F/F signal size. The raw data were first corrected for this effect by subtracting the average background fluorescence intensity determined from an unstained area on the slice. Subsequently, the signal alignment software was used to correct for temporal jitter in AP initiation as well as for possible small movements of the preparation during averaging. In the temporal domain, AP signals were aligned by cross correlation of the electrically recorded APs in each trial to the reference signal acquired at the start of averaging (Figure 1B). In the spatial domain, images were aligned in two dimensions offline by image cross-correlation to compensate for possible small lateral movements of the preparation. The correct focus of the image in the z-dimension was verified before each individual trial; small adjustments were often necessary. The spatially and temporally aligned signals were averaged as shown in Figure 1B. Slow changes in light intensity due to bleaching of the dye were corrected by dividing the data by an appropriate dual exponential function derived from the recording trials with no stimulation (Figure 1B). The waveform of the AP signal was reconstructed from a set of data points using Cubic Spline Interpolation, a piecewise continuous curve passing through each data point. To confirm that the voltage-sensitive dye signal tracks membrane potential without significant distortion at the millisecond time scale, the electrical AP signal from the soma was compared to the optical AP signal from the adjacent axon hillock as shown in Figure 3B. The two signals superimpose very closely, allowing for the shot noise in optical recording.
<img alt="Figure 1" src="/files/ftp_upload/4261/4261fig1.jpg" /> 
 Figure 1. Data analysis (Animation). (A) Upper panel: high resolution confocal image of a stained neuron with axon in recording position. Recording electrode attached to soma and extracellular stimulating electrode next to basal dendrite shown schematically. Action potentials evoked by extracellular, synaptic stimulation. Lower panel: low spatial resolution fluorescence image of the axon obtained by CCD used for Vm imaging. (B) Electrode recordings from soma (black traces), optical recordings from AIS (red traces), and from node of Ranvier (green traces). Top traces: raw data from 9 trials showing temporal jitter in AP initiation. Second row of traces: temporally aligned signals. Third row of traces: averaged signal. Fourth row of traces: bleach correction. Bottom traces: cubic spline interpolation with one pass of temporal smoothing.

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Dejan Zecevic declares that he is a co-owner of RedShirtImaging LLC., a company specialized in high-speed, low noise CCD cameras used in voltage-sensitive dye recording. All other authors report no financial interests or potential conflicts of interest related to the current study.

This article describes a voltage-sensitive dye recording method for monitoring electrical activity of individual neurons with sub-micrometer and sub-millisecond spatiotemporal resolution. Laser excitation at near-optimal wavelength (regarding signal size) improved the sensitivity of recording by a factor of ~50 over previous approaches. The current sensitivity enables monitoring electrical signals from all parts of individual neurons, including dendrites, axons, axon collaterals and axon terminals as well as individual d...

<table border="1">
 <tbody>
 <tr>
 <td>Name of the component</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Setup components</td>
 </tr>
 <tr>
 <td>Upright Microscope</td>
 <td>Olympus Inc.</td>
 <td>BX51WI</td>
 <td>With three camera ports</td>
 </tr>
 <tr>
 <td>Motorized Movable Stage</td>
 <td>Siskiyou</td>
 <td>MXOPi.2</td>
 <td></td>
 </tr>
 <tr>
 <td>Epi-fluorescence Condenser for Olympus BX51</td>
 <td>TILL Photonics</td>
 <td>0000-560-11659</td>
 <td></td>
 </tr>
 <tr>
 <td>Upright Microscope</td>
 <td>Carl Zeiss, LLC </td>
 <td>AxioExaminer D1</td>
 <td>With three camera ports</td>
 </tr>
 <tr>
 <td>Motorized Top Plate</td>
 <td>Scientifica Limited</td>
 <td>MMBP</td>
 <td></td>
 </tr>
 <tr>
 <td>Epi-fluorescence Condenser for Zeiss AxioExaminer</td>
 <td>TILL Photonics</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Data Acquisition Camera</td>
 <td>RedShirtImaging LLC</td>
 <td>NeuroCCD-SM</td>
 <td>High speed, low read noise</td>
 </tr>
 <tr>
 <td>CCD for IR-DIC</td>
 <td>Dage-MTI</td>
 <td>IR-1000</td>
 <td></td>
 </tr>
 <tr>
 <td>Spinning-Disc Confocal Scanner</td>
 <td>Yokogawa</td>
 <td>CSU-10</td>
 <td></td>
 </tr>
 <tr>
 <td>High Spatial Resolution CCD on Confocal Scanner</td>
 <td>PCO AG</td>
 <td>PixelFly</td>
 <td>1392x1024 pixels </td>
 </tr>
 <tr>
 <td>DPSS CW Laser (532 Nm)</td>
 <td>CNI Optoelectronics Tech. Co., Ltd</td>
 <td>MLL-III-532 400mW</td>
 <td>Excitation light source</td>
 </tr>
 <tr>
 <td>Multi-Mode Fiber Launcher</td>
 <td>Siskiyou</td>
 <td>SM-CFT</td>
 <td></td>
 </tr>
 <tr>
 <td>Light Guide </td>
 <td>TILL Photonics</td>
 <td>0000-515-11524</td>
 <td></td>
 </tr>
 <tr>
 <td>Shutter</td>
 <td>Vincent Associates</td>
 <td>LS6</td>
 <td></td>
 </tr>
 <tr>
 <td>Vibration Isolation Table</td>
 <td>Minus k Technology</td>
 <td>MK26</td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Specific reagents</td>
 </tr>
 <tr>
 <td>Di-2-ANEPEQ (JPW 1114)</td>
 <td>Life Technologies</td>
 <td>D-6923</td>
 <td>Voltage sensitive dye</td>
 </tr>
 <tr>
 <td>Crym-EGFP Mouse Line</td>
 <td>GENSAT (MMRRC)</td>
 <td>STOCK Tg(Crym-EGFP)GF82Gsat/Mmcd</td>
 <td>Sparsely expressing EGFP in Layer 5 cortical neurons</td>
 </tr>
 </tbody>
</table>

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.

Successful confocal microscopy should allow clear identification of intact neuronal processes which are close to the surface of the slice and located in one plane of focus. The selection of nerve cells which are appropriate for voltage imaging prior to voltage-sensitive dye loading is critical. An example of confocal images of L5 pyramidal neurons expressing EGFP in a cortical slice (Crym transgenic mouse line) is shown in Figure 2. Axons of individual neurons are clearly visible. Cells with long intact ...

Watch this Scientific Journal Video about Voltage-sensitive Dye Recording from Axons, Dendrites and Dendritic Spines of Individual Neurons in Brain Slices at JoVE.com

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

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 ...

voltage-sensitive-dye-recording-from-axons-dendrites-dendritic-spines

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Neuroscience

16.6K Views.  Yale University School of Medicine. 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.

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

DiOLISTIC Labeling of Neurons from Rodent and Non-human Primate Brain Slices

DiOLISTIC staining uses the gene gun to introduce fluorescent dyes, such as DiI, into neurons of brain slices (Gan et al., 2009; O'Brien and Lummis, 2007; Gan et al., 2000). Here we provide a detailed description of each step required together with exemplary images of good and bad outcomes that will help when setting up the technique. In our experience, a few steps proved critical for the successful application of DiOLISTICS. These considerations include the quality of the DiI-coated bullets, the extent of fixative exposure, and the concentration of detergent used in the incubation solutions. Tips and solutions for common problems are provided. This is a versatile labeling technique that can be applied to multiple animal species at a wide range of ages. Unlike other fluorescent labeling techniques that are limited to preparations from young animals or restricted to mice because they rely on the expression of a fluorescent transgene, DiOLISTIC labeling can be applied to animals of all ages, species and genotypes and it can be used in combination with immunostaining to identify a specific subpopulation of cells. Here we demonstrate the use of DiOLISTICS to label neurons in brain slices from adult mice and adult non-human primates with the purpose of quantifying dendrite branching and dendritic spine morphology.

We demonstrate the use of the gene gun to introduce fluorescent dyes, such as DiI, into neurons in brain slices from rodents and non-human primates of different ages. In this particular case, we use adult mice (3-6 months old) and adult cynomologus monkeys (9-15 years old). This technique, originally described by the laboratory of Dr. Lichtman (Gan et al., 2000), is well suited for the study of dendritic branching and dendritic spine morphology and can be combined with traditional immunostaining, if detergents are kept at a low concentration.

DiOLISTIC staining uses the gene gun to introduce fluorescent dyes, such as DiI, into neurons of brain slices (Gan et al., 2009; O'Brien and Lummis, 2007; ...

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.

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 ...

Fluorescence Recovery After Photobleaching (FRAP) of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached in the region of interest. The fluorescence of the region recovers when the unbleached GFP-tagged protein from outside of the region diffuses into the region of interest. The mobility of the protein is then analyzed by measuring the fluorescence recovery rate. This technique could be used to characterize protein mobility and turnover rate.
In this study, we express the (enhanced green fluorescent protein) EGFP vector in cultured hippocampal neurons. Using the Zeiss 710 confocal microscope, we photobleach the fluorescence signal of the GFP protein in a single spine, and then take time lapse images to record the fluorescence recovery after photobleaching. Finally, we estimate the percentage of mobile and immobile fractions of the GFP in spines, by analyzing the imaging data using ImageJ and Graphpad softwares.
This FRAP protocol shows how to perform a basic FRAP experiment as well as how to analyze the data.

Fluorescence Recovery After Photobleaching FRAP of Fluorescence Tagged Proteins in Dendritic Spines of Cultured Hippocampal Neurons

FRAP has been used to quantify the mobility of Green Fluorescence Protein (GFP)-tagged proteins in cultured cells. We examined the mobile/immobile fractions of the GFP by analyzing the fluorescence recovery percentage after photobleaching. In this study, FRAP was performed at spines of hippocampal neurons.

FRAP has been used to quantify the mobility of GFP-tagged proteins. Using a strong excitation laser, the fluorescence of a GFP-tagged protein is bleached ...

Vibrodissociation of Neurons from Rodent Brain Slices to Study Synaptic Transmission and Image Presynaptic Terminals

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of interest. This allows for examination of synaptic transmission under conditions where the extracellular and postsynaptic intracellular environments can be well controlled. A vibration-based technique without the use of proteases, known as vibrodissociation, is the most popular technique for mechanical isolation. A micropipette, with the tip fire-polished to the shape of a small ball, is placed into a brain slice made from a P1-P21 rodent. The micropipette is vibrated parallel to the slice surface and lowered through the slice thickness resulting in the liberation of isolated neurons. The isolated neurons are ready for study within a few minutes of vibrodissociation. This technique has advantages over the use of primary neuronal cultures, brain slices and enzymatically isolated neurons including: rapid production of viable, relatively mature neurons suitable for electrophysiological and imaging studies; superior control of the extracellular environment free from the influence of neighboring cells; suitability for well-controlled pharmacological experiments using rapid drug application and total cell superfusion; and improved space-clamp in whole-cell recordings relative to neurons in slice or cell culture preparations. This preparation can be used to examine synaptic physiology, pharmacology, modulation and plasticity. Real-time imaging of both pre- and postsynaptic elements in the living cells and boutons is also possible using vibrodissociated neurons. Characterization of the molecular constituents of pre- and postsynaptic elements can also be achieved with immunological and imaging-based approaches.

This report demonstrates a technique for mechanical isolation of individual viable neurons retaining attached presynaptic boutons. Vibrodissociated neurons have the advantages of rapid production, excellent pharmacological control and improved space-clamp without influence from neighboring cells. This method can be used for imaging of synaptic elements and patch-clamp recording.

Mechanical dissociation of neurons from the central nervous system has the advantage that presynaptic boutons remain attached to the isolated neuron of ...

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic development and axonal wiring. Recently the dendritic arborization (DA) sensory neurons of the Drosophila larval peripheral nervous system (PNS) have become powerful genetic models in which to elucidate both general and class-specific mechanisms of neuron differentiation.
There are four main DA neuron classes (I-IV)1. They are named in order of increasing dendrite arbor complexity, and have class-specific differences in the genetic control of their differentiation2-10. The DA sensory system is a practical model to investigate the molecular mechanisms behind the control of dendritic morphology11-13 because: 1) it can take advantage of the powerful genetic tools available in the fruit fly, 2) the DA neuron dendrite arbor spreads out in only 2 dimensions beneath an optically clear larval cuticle making it easy to visualize with high resolution in vivo, 3) the class-specific diversity in dendritic morphology facilitates a comparative analysis to find key elements controlling the formation of simple vs. highly branched dendritic trees, and 4) dendritic arbor stereotypical shapes of different DA neurons facilitate morphometric statistical analyses.
DA neuron activity modifies the output of a larval locomotion central pattern generator14-16. The different DA neuron classes have distinct sensory modalities, and their activation elicits different behavioral responses14,16-20. Furthermore different classes send axonal projections stereotypically into the Drosophila larval central nervous system in the ventral nerve cord (VNC)21. These projections terminate with topographic representations of both DA neuron sensory modality and the position in the body wall of the dendritic field7,22,23. Hence examination of DA axonal projections can be used to elucidate mechanisms underlying topographic mapping7,22,23, as well as the wiring of a simple circuit modulating larval locomotion14-17.
We present here a practical guide to generate and analyze genetic mosaics24 marking DA neurons via MARCM (Mosaic Analysis with a Repressible Cell Marker)1,10,25 and Flp-out22,26,27 techniques (summarized in Fig. 1).

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

The dendritic arborization sensory neurons of the Drosophila larval peripheral nervous system are useful models to elucidate both general and neuron class-specific mechanisms of neuron differentiation. We present a practical guide to generate and analyze dendritic arborization neuron genetic mosaics.

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic ...

Recording Electrical Activity from Identified Neurons in the Intact Brain of Transgenic Fish

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be performed in an intact brain preparation where the neural circuitry of the CNS remains intact. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with green fluorescent protein for identification in the intact brain. Fish have multiple populations of GnRH neurons, and their functions are dependent on their location in the brain and the GnRH gene that they express1 . We have focused our demonstration on GnRH3 neurons located in the terminal nerves (TN) associated with the olfactory bulbs using the intact brain of transgenic medaka fish (Figure 1B and C). Studies suggest that medaka TN-GnRH3 neurons are neuromodulatory, acting as a transmitter of information from the external environment to the central nervous system; they do not play a direct role in regulating pituitary-gonadal functions, as do the well-known hypothalamic GnRH1 neurons2, 3 .The tonic pattern of spontaneous action potential firing of TN-GnRH3 neurons is an intrinsic property4-6, the frequency of which is modulated by visual cues from conspecifics2 and the neuropeptide kisspeptin 15. In this video, we use a stable line of transgenic medaka in which TN-GnRH3 neurons express a transgene containing the promoter region of Gnrh3 linked to enhanced green fluorescent protein7 to show you how to identify neurons and monitor their electrical activity in the whole brain preparation6.

In this video, we will demonstrate how to record electrical activity from identified single neurons in a whole brain preparation, which preserves complex neural circuits. We use transgenic fish in which gonadotropin-releasing hormone (GnRH) neurons are genetically tagged with a fluorescent protein for identification in the intact brain preparation.

Understanding the cell physiology of neural circuits that regulate complex behaviors is greatly enhanced by using model systems in which this work can be ...

Imaging Dendritic Spines of Rat Primary Hippocampal Neurons using Structured Illumination Microscopy

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. Technological advances have identified these structures as key elements in neuron connectivity and synaptic plasticity. The quantitative analysis of spine morphology using light microscopy remains an essential problem due to technical limitations associated with light's intrinsic refraction limit. Dendritic spines can be readily identified by confocal laser-scanning fluorescence microscopy. However, measuring subtle changes in the shape and size of spines is difficult because spine dimensions other than length are usually smaller than conventional optical resolution fixed by light microscopy's theoretical resolution limit of 200 nm.
Several recently developed super resolution techniques have been used to image cellular structures smaller than the 200 nm, including dendritic spines. These techniques are based on classical far-field operations and therefore allow the use of existing sample preparation methods and to image beyond the surface of a specimen. Described here is a working protocol to apply super resolution structured illumination microscopy (SIM) to the imaging of dendritic spines in primary hippocampal neuron cultures. Possible applications of SIM overlap with those of confocal microscopy. However, the two techniques present different applicability. SIM offers higher effective lateral resolution, while confocal microscopy, due to the usage of a physical pinhole, achieves resolution improvement at the expense of removal of out of focus light. In this protocol, primary neurons are cultured on glass coverslips using a standard protocol, transfected with DNA plasmids encoding fluorescent proteins and imaged using SIM. The whole protocol described herein takes approximately 2 weeks, because dendritic spines are imaged after 16-17 days in vitro, when dendritic development is optimal. After completion of the protocol, dendritic spines can be reconstructed in 3D from series of SIM image stacks using specialized software.

This article describes a working protocol to image dendritic spines from hippocampal neurons in vitro using Structured Illumination Microscopy (SIM). Super-resolution microscopy using SIM provides image resolution significantly beyond the light diffraction limit in all three spatial dimensions, allowing the imaging of individual dendritic spines with improved detail.

Dendritic spines are protrusions emerging from the dendrite of a neuron and represent the primary postsynaptic targets of excitatory inputs in the brain. ...

Acute Dissociation of Lamprey Reticulospinal Axons to Enable Recording from the Release Face Membrane of Individual Functional Presynaptic Terminals

Synaptic transmission is an extremely rapid process. Action potential driven influx of Ca2+&#160;into the presynaptic terminal, through voltage-gated calcium channels (VGCCs) located in the release face membrane, is the trigger for vesicle fusion and neurotransmitter release. Crucial to the rapidity of synaptic transmission is the spatial and temporal synchrony between the arrival of the action potential, VGCCs and the neurotransmitter release machinery. The ability to directly record Ca2+&#160;currents from the release face membrane of individual presynaptic terminals is imperative for a precise understanding of the relationship between presynaptic Ca2+&#160;and neurotransmitter release. Access to the presynaptic release face membrane for electrophysiological recording is not available in most preparations and presynaptic Ca2+&#160;entry has been characterized using imaging techniques and macroscopic current measurements &#8211; techniques that do not have sufficient temporal resolution to visualize Ca2+ entry. The characterization of VGCCs directly at single presynaptic terminals has not been possible in central synapses and has thus far been successfully achieved only in the calyx-type synapse of the chick ciliary ganglion and in rat calyces. We have successfully addressed this problem in the giant reticulospinal synapse of the lamprey spinal cord by developing an acutely dissociated preparation of the spinal cord that yields isolated reticulospinal axons with functional presynaptic terminals devoid of postsynaptic structures. We can fluorescently label and identify individual presynaptic terminals and target them for recording. Using this preparation, we have characterized VGCCs directly at the release face of individual presynaptic terminals using immunohistochemistry and electrophysiology approaches. Ca2+ currents have been recorded directly at the release face membrane of individual presynaptic terminals, the first such recording to be carried out at central synapses.

Recording Ca2+ currents at the presynaptic release face membrane is key to a precise understanding of Ca2+ entry and neurotransmitter release. We present an acute dissociation of the lamprey spinal cord that yields functional isolated reticulospinal axons, permitting recording directly from the release face membrane of individual presynaptic terminals.

Synaptic transmission is an extremely rapid process. Action potential driven influx of Ca2+&#160;into the presynaptic terminal, through voltage-gated ...

Recording Synaptic Plasticity in Acute Hippocampal Slices Maintained in a Small-volume Recycling-, Perfusion-, and Submersion-type Chamber System

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful analysis of synaptic transmission modulation when performing field potential or intracellular recordings. This manuscript describes methodological aspects that might be helpful in improving experimental conditions for the maintenance of acute brain slices and for recording field excitatory postsynaptic potentials in a commercially available submersion chamber with an outflow-carbogenation unit. The outflow-carbogenation helps to stabilize the oxygen level in experiments that rely on the recycling of a small buffer reservoir to enhance the cost-efficiency of drug experiments. In addition, the manuscript presents representative experiments that examine the effects of different carbogenation modes and stimulation paradigms on the activity-dependent synaptic plasticity of synaptic transmission.

This protocol describes the stabilization of the oxygen level in a small volume of recycled buffer and methodological aspects of recording activity-dependent synaptic plasticity in submerged acute hippocampal slices.

Even though experiments on brain slices have been in use since 1951, problems remain that reduce the probability of achieving a stable and successful ...

Two-photon Calcium Imaging in Neuronal Dendrites in Brain Slices

Calcium (Ca2+) imaging is a powerful tool to investigate the spatiotemporal dynamics of intracellular Ca2+ signals in neuronal dendrites. Ca2+ fluctuations can occur through a variety of membrane and intracellular mechanisms and play a crucial role in the induction of synaptic plasticity and regulation of dendritic excitability. Hence, the ability to record different types of Ca2+ signals in dendritic branches is valuable for groups studying how dendrites integrate information. The advent of two-photon microscopy has made such studies significantly easier by solving the problems inherent to imaging in live tissue, such as light scattering and photodamage. Moreover, through combination of conventional electrophysiological techniques with two-photon Ca2+ imaging, it is possible to investigate local Ca2+ fluctuations in neuronal dendrites in parallel with recordings of synaptic activity in soma. Here, we describe how to use this method to study the dynamics of local Ca2+ transients (CaTs) in dendrites of GABAergic inhibitory interneurons. The method can be also applied to studying dendritic Ca2+ signaling in different neuronal types in acute brain slices.

We present a method combining whole-cell patch-clamp recordings and two-photon imaging to record Ca2+ transients in neuronal dendrites in acute brain slices.