We thank the late&#160;Rudy Limburg for fabricating the cardan arm perimeter, Kimberly Jamison, Matthew McHenry, and Raju Metherate for lending us equipment, and Almut Kelber and Kentaro Arikawa, for encouragement. This work was supported by a National Science Foundation (NSF) Graduate Research Fellowship to KJM and NSF grant IOS-1257627 to A.D.B.

The authors declare that they have no competing financial interests.

Intracellular recording can be a difficult technique to master due to the many technical steps involved. For successful experiments several important points must be considered. First, it is important to have a properly vibrationally-isolated table on which the experiment is performed. Many researchers use air tables, which completely separate the tabletop from the base, giving superior vibration isolation. Our setup involves a thick marble table with a sandbox on top, into which is placed the micromanipulator/electrode holder/specimen stage apparatus. This is an effective and more affordable alternative to an air table, especially if access to in-house gas or compressed air is a limitation. Additional vibration-absorbing measures may be taken such as passive air suspension, or the addition of cushioning elements to the table legs (e.g., opened tennis balls, bike tubes, thick gel pads). Furthermore, it is essential that the experimental setup be inside a Faraday cage with everything properly grounded. The Faraday cage should have a metal mesh screen in the front that can be removed when working inside the cage and replaced easily when recording. Even a small amount of ambient electrical noise (especially 50 Hz noise from the main AC power supply) can make an otherwise good recording unusable.
When preparing the specimen, hemolymph and pigment layers surrounding the ommatidia may prevent successful recordings. If the hole in the cornea is cut too large, normal pumping of hemolymph in the body causes the liquid surface to move up and down at the cut site, resulting in an unstable recording. Once the hole is cut, hemolymph and surface pigment layers will clot rapidly into an impenetrable scab even when sealed with petroleum jelly, so it is important to get the electrode into the eye as soon as possible. Ringer&#39;s solution may also be used instead of Vaseline. The ground electrode may be introduced into the mouthparts or into the stump of a cut antenna.
Additionally, it is often helpful to keep the animal as dark-adapted as possible. For this method, steps include keeping the recording room very dark, blocking stray light from the Xenon lamp from entering the Faraday cage, short duration stimulus flashes (30-50 msec), and a low enough frequency between flashes (0.5 or greater). When a visual pigment absorbs a photon, the chromophore in rhodopsin switches from 11-cis-3-hydroxyretinal to all-trans-retinal, inducing the conformational change of the opsin protein, and activating the entire complex as metarhodopsin, which initiates the G protein cascade. Photo-bleaching occurs when high intensity light causes the chromophore to physically separate from the opsin protein. Time is a limiting factor in this experiment because the electrode will only stably record responses from within the cell for a certain period of time before it falls out or the membrane is damaged. For this reason we do not break to allow the cell to recover, but we do use a flash duration and frequency that we have found does not degrade the cell&#39;s response over time. It is important to decrease both frequency and intensity of light if photo-bleaching occurs.
During recording, a large electrode tip or large movement by the electrode may damage the cell when penetrating the membrane. Only fine tips (at least ~100 M&#937;) and small movements should be used when approaching a cell for recording. If intracellular recording is applied to other applications, such as brain recordings, extremely fine, sturdy electrodes may be pulled using quartz glass, but a specialized puller must be used for these electrodes. When first making an electrode pulling program, we checked tip resistance by backfilling the electrode, securing it to the electrode holder in a mock setup, and placing the tip and ground electrode in saline solution. Next we applied a current to measure change in voltage on the oscilloscope. To move the electrode tip we use a manual micromanipulator that moves along two axes. Other manipulators exist including digital ones that allow movement along all three axes and these may be used for this or more complex applications. There are many ways to build a stage for recording, and there are many different types of hardware and software used in recording and analyzing the observed data. Our setup represents one simple, easy, and affordable setup of the recording rig.
In constructing the VlogI curve, functions developed by Naka and Rushton33 and others34,35 account for the non-linear portions of the plotted responses. Various methods are used to fit this curve to the data, and we plotted the results of one such method that does not require curve-fitting software, though other methods are also suitable36,37 (Figure 7). It may also be useful to compare spectral sensitivities to models of rhodopsin absorbance at a given peak wavelength. Several published models aim to reproduce rhodopsin absorbance spectra38,39. A more precise idea of the absorbance spectrum of the visual pigment expressed in an insect photoreceptor cell may be modeled by taking into account ommatidial properties such as filtering pigments, but this requires measurement of additional physiological and anatomical parameters11,40.
One limitation of the method is that if the study organism expresses more than one genetically similar opsin in the eye, it can be difficult to identify which opsin mRNA likely corresponds to which spectral class of photoreceptor cell. To overcome this problem, this method has been combined with dye-injections and in situ hybridization or immunohistochemistry to successfully identify the opsins expressed in recorded cells10.
Our method is simple and accessible for researchers unfamiliar with visual electrophysiology. This technique is common in neuroscience, but specific and clear methods are absent in the literature, making this method difficult to reproduce. Although many variations of this technique exist, we offer a straightforward way to measure spectral sensitivity in the compound eye. The physiological data is an important piece of evidence in stories of visual ecology and evolution41. Opsin sequence variation is linked closely with the sensitivity of a photoreceptor cell, making this method ideal for studies examining the genetic basis for phenotypic change. Measurement of photoreceptor cell sensitivities may also be paired with behavioral color discrimination assays, showing the physiological basis for important discrimination thresholds in color vision42-46. In genetic or therapeutic manipulations in Drosophila for example, this technique can be a good way to measure proper physiological function of the eye or brain as well47,48. Although ours is not the first or the most complex method of intracellular recording in the eye, our hope is that we can make this method more easily available for reproduction and integration in research programs outside of formal neuroscience.

The electrical properties of cells such as neurons are observed by measuring ion flow across cell membranes as a change in voltage or current. A variety of electrophysiological techniques have been developed to measure bioelectric events in cells. Neurons found in the eyes of animals are accessible and their circuitry is often less complex than in the brain, making these cells good candidates for electrophysiological study. Common applications of electrophysiology in the eye include electroretinography (ERG)1,2 and microelectrode intracellular recording. ERG involves placing an electrode in or on the eye of an animal, applying a light stimulus, and measuring the change in voltage as a sum of the responses of all nearby cells3-6. If one is specifically interested in characterizing spectral sensitivities of individual photoreceptor cells, often multiple cell types simultaneously respond at different strengths&#160;to a given stimulus; thus it can be difficult to determine the sensitivities of specific cell types from ERG data especially if there are several different kinds of spectrally-similar photoreceptor cells in the eye. One potential solution is to create transgenic Drosophila with the photoreceptor (opsin) gene of interest expressed in the majority R1-6 cells in the eye and then perform ERGs7. Potential drawbacks of this method include no to low-expression of the photoreceptor protein8, and the long time frame for the generation and screening of transgenic animals. For eyes with fewer kinds of spectrally distinct photoreceptors, adaptation of the eye with colored filters can help with lowering the contribution of some cell types to the ERG, thereby permitting estimation of spectral sensitivity maxima9.
Intracellular recording is another technique where a fine electrode impales a cell and a stimulus is applied. The electrode records only that individual cell&#39;s response so that recording from and analyzing multiple individual cells can yield specific sensitivities of physiologically different cell types10-14. Although our protocol focuses on analysis of spectral sensitivity, the basic principles of intracellular recording with sharp electrodes are modifiable for other applications. Using a different preparation of a specimen, for instance, and using sharp quartz electrodes, one may record from deeper in the optic lobe or other regions in the brain, depending on the question being asked. For example, response times of individual photoreceptor cells15, cell activity in the optic lobes16 (lamina, medulla or lobula17), brain18 or other ganglia19 can also be recorded with similar techniques, or color stimuli could be replaced with polarization20-22 or motion stimuli23,24.
Phototransduction, the process by which light energy is absorbed and converted into an electrochemical signal, is an ancient trait common to nearly all present day animal phyla25. The visual pigment found in photoreceptor cells and responsible for initiating visual phototransduction is rhodopsin. Rhodopsins in all animals are made up of an opsin protein, a member of the 7 transmembrane G protein-coupled receptor family, and an associated chromophore which is derived from retinal or a similar molecule26,27. Opsin amino acid sequence and chromophore structure affect the absorbance of rhodopsin to different wavelengths of light. When a photon is absorbed by the chromophore the rhodopsin becomes activated, initiating a G-protein cascade in the cell that ultimately leads to the opening of membrane-bound ion channels28. Unlike most neurons, photoreceptor cells undergo graded potential changes that can be measured as a relative change in response amplitude with changing light stimulus. Typically a given photoreceptor type expresses only one opsin gene (though exceptions exist8,10,29-31). Sophisticated color vision, of the kind found in many vertebrates and arthropods, is achieved with a complex eye of hundreds or thousands of photoreceptor cells each expressing one or occasionally more rhodopsin types. Visual information is captured by comparing responses over the photoreceptor mosaic via complex downstream neural signaling in the eye and brain, resulting in the perception of an image complete with color and motion.
After measuring the raw responses of a photoreceptor cell to different wavelengths of light via intracellular recording, it is possible to calculate its spectral sensitivity. This calculation is based on the Principle of Univariance, which states that a photoreceptor cell&#39;s response is dependent on the number of photons it absorbs, but not on the particular properties of the photons it absorbs32. Any photon that is absorbed by rhodopsin will induce the same kind of response. In practice, this means that a cell&#39;s raw response amplitude will increase due to either an increase in light intensity (more photons to absorb), or to a shift in wavelength toward its peak sensitivity (higher probability of rhodopsin absorbing that wavelength). We make use of this principle in relating cellular responses at known intensity and the same wavelength to responses at different wavelengths and the same intensity but unknown relative sensitivity. Cell types are often identified by the wavelength at which their sensitivity peaks.
Here we show one method for intracellular recording and analysis of spectral sensitivity of photoreceptors in the eye of a butterfly, with a focus on making this method more accessible to the wider research community. Although intracellular recording remains common in the literature, particularly with respect to color vision in insects, we have found that descriptions of materials and methods are usually too brief to allow for reproduction of the technique. We present this method in video format with the aim of permitting its easier replication. We also describe the technique using easily obtainable and affordable equipment. We address common caveats that often are not reported, which slow down research when optimizing a new and complex technique.

determination of photoreceptor cell spectral sensitivity in an insect model from in vivo intracellular recordings

Intracellular recording is a powerful technique used to determine how a single cell may respond to a given stimulus. In vision research, intracellular recording has historically been a common technique used to study sensitivities of individual photoreceptor cells to different light stimuli that is still being used today. However, there remains a dearth of detailed methodology in the literature for researchers wishing to replicate intracellular recording experiments in the eye. Here we present the insect as a model for examining eye physiology more generally. Insect photoreceptor cells are located near the surface of the eye and are therefore easy to reach, and many of the mechanisms involved in vision are conserved across animal phyla. We describe the basic procedure for in vivo intracellular recording of photoreceptor cells in the eye of a butterfly, with the goal of making this technique more accessible to researchers with little prior experience in electrophysiology. We introduce the basic equipment needed, how to prepare a live butterfly for recording, how to insert a glass microelectrode into a single cell, and finally the recording procedure itself. We also explain the basic analysis of raw response data for determining spectral sensitivity of individual cell types. Although our protocol focuses on determining spectral sensitivity, other stimuli (e.g., polarized light) and variations of the method are applicable to this setup.

For many elements of the recording setup, a written description does not provide enough detail. Figure 1 is a schematic of the components involved in the complete recording setup. In Figure 2, spectra are plotted for white light and each interference filter to give a sense of why a correction factor is needed and what is needed to calculate this correction. Figure 3 shows photos and a diagram of the Cardan arm that is used for these experiments. Figure 4 is a composite image showing the recording stage and micromanipulator. Figure 5 shows several steps in the preparation of a butterfly for the experiment.&#160;
Once a recording begins, a negative change in voltage in response to a light flash means that the electrode is outside of a cell, as in Figure 6a. The strength of the response depends on the proximity of the electrode tip to a photoreceptor cell, and the angle of incidence of the light flash. The response should be large (&#62;30 mV) before the tip is near enough to a cell to impale. Figure 6b shows a clear depolarizing response to a light stimulus, signifying entry into a photoreceptor cell. The resting potential should be stable and the response amplitude should be large (at least 40 mV), although the absolute amplitude may vary considerably. We measure relative response, so it is more important that the signal to noise ratio is high. If the resting potential changes greatly, the response waveform looks unusual, or the maximum response is too low, then comparing relative responses across all interference filters becomes impossible. Examples of unusable recordings are shown in Figure 6c, 6d.
After completing a successful recording, the ND responses must be plotted and the Naka-Rushton equation should be fitted to the data33, shown in Figure 7. This figure is plotted using the ND filter series without any interference filters. If the recording is stable, the data from the ND filter series should be similar before and after the experiment. Spectral sensitivity is determined by fitting the Naka-Rushton equation to the VlogI plot in Figure 7, then solving for (I) for each response (V) at a given wavelength, as explained in the calculations of Section 6 of the protocol.
A representative example of spectral sensitivity derived from a single recording is plotted in Figure 8a (please note this example shows real calculated data, but the peak has been shifted as this result is unpublished). Cell types may be classified by peak sensitivity at a similar wavelength and overall shape of the sensitivity spectrum. Similar cell types are then averaged and the mean sensitivity is plotted with standard error bars at each wavelength in Figure 8b. Spectral sensitivities of three typical cell types found in an insect are shown in Figure 8c (magnitude and error bars are calculated from real data, but the peaks are shifted).
<img alt="Figure 3" src="/files/ftp_upload/53829/53829fig3.jpg" style="opacity: 0.9; font-size: 13.888888359069824px; line-height: 28px;" />
Figure 1:&#160;Schematic of Recording Components. Components of the light path, specimen setup, and recording hardware are indicated. Optical track sits on the wooden bench outside the Faraday cage. Xe, xenon arc lamp, Cd, condenser assembly, Lx, convex lens, ND, neutral density filter wheel, Cf, interference filter, S, shutter, Lc, concave lens, Co, collimating beam probe, Fo, fiber optic cable. The Faraday cage sits on the wooden bench around the specimen. The wooden bench sits above but does not touch the vibrationally isolated marble table underneath. Recording (Rc) and reference (Rf) electrodes are attached to the headstage (Hs). Rc is introduced into the eye (E) and Rf is introduced into another part of the body. The headstage is part of the preamplifier (Pa) setup outside the Faraday cage. The signal is passed from the preamplifier through the Humbug noise reducer (Hb), and into the oscilloscope (Os). From the oscilloscope the signal is passed through the Powerlab hardware (Plb) and into the laptop computer (Cpu) where it is read by the Labchart software. The shutter is controlled by a function generator (Fg) which is passed through a second channel on the oscilloscope, and may also be passed through a second channel on the Powerlab hardware if this signal is going to be recorded as well.
<img alt="Figure 5" src="/files/ftp_upload/53829/53829fig5.jpg" style="opacity: 0.9; font-size: 13.888888359069824px; line-height: 28px;" />&#160;
Figure 2:&#160;White Light and Interference Filter Spectra Used to Calculate Correction Factors. The spectra measured in step 2.5 of the protocol are shown from 300 to 700 nm, for white light as well as each of the 41 interference filters. Each interference filter spectrum is measured with only that filter in the light path. T520 corresponds to the area under the spectrum for the filter with peak 520 nm, and s520 corresponds to the intensity of white light at peak wavelength of the filter, 520 nm. These values are used in calculating the correction factors for each filter (in this case 520 nm) as described in step 2.6 in the protocol.
<img alt="Figure 4" src="/files/ftp_upload/53829/53829fig4.jpg" style="opacity: 0.9; font-size: 13.888888359069824px; line-height: 28px;" />
Figure 3:&#160;Description of the Cardan Arm Perimeter Used in the Experiments.&#160;The Cardan arm holds the fiber optic cable and allows full spherical rotation and angular adjustment so that light may be delivered at the proper angle of incidence to the cell being recorded. (A) Top down view. (B) View from the side at an angle. Numbers correspond to the same part in all panels. (1) The bottom plate allows full circular rotation in the horizontal plane. (2) The vertical plate allows full circular rotation of the arm in a vertical plane. (3) This cylinder holds the metal arm with the fiber optic cable on the end, and it allows a second vertical plane of circular rotation perpendicular to (2). (4) The fiber optic cable is held in place at the end of the arm, and light is directed toward the location of the specimen in the experimental setup.
<img alt="Figure 1" src="/files/ftp_upload/53829/53829fig1.jpg" /> 
Figure 4: Components of the Recording Setup. (A) Plastic tube used to hold the specimen. (B) Overhead view of the platform on which the specimen and tube are mounted. (C) Side view of ball-and-joint platform with magnetic base (1). Reference electrode kept immobile with glue and wax on side of platform (2). Reference electrode wrapped around alligator clip and soldered in place, providing an attachment area where the headstage reference electrode can clip (3). (D) Electrode tip under 20X&#160;magnification. Scale bar, 25 &#181;m. (E) Stage, electrode holder, and micromanipulator setup. The headstage (1) is fixed above the apparatus with the silver recording wire (2), and the reference electrode with alligator clip (3) attached. The electrode holder (4) is fixed to a manual micromanipulator (5) with a post below that may be adjusted with a knob for vertical movement or may be pushed or pulled for horizontal movement of the electrode holder. The magnetic platform with specimen sits on the stage (6) just below the electrode holder. (F) The Faraday cage surrounds the recording setup with a screen that can be pulled up or down in the front. Aluminum foil is placed underneath all equipment with rubber pads on top. The fiber optic cable (1) leads into the cage from the optical track outside, and is directed by the Cardan arm (2) toward the stage (3). The recording stage and manipulator apparatus is placed in a sand box (4) resting on a marble table underneath the setup. All other equipment rests on the wooden bench top that does not touch the marble table. The sand box sits on top of the marble table in a hole cut out of the wooden table, so that the specimen is completely vibrationally isolated from the equipment on the wooden tabletop. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53829/53829fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53829/53829fig2.jpg" /> 
Figure 5:&#160;Butterfly Prep. (A) The butterfly is inserted into the tube and the head, wings, and antennae are immobilized with hot wax. The indifferent electrode is inserted into the mouthparts (1), a piece of dry wax is used to hold down the abdomen (2), and a wet tissue is placed behind the specimen. (B) Close-up of the head waxed down. The eye to be recorded from (1) is kept clear from wax or debris, and the reference electrode (2) is inserted into the mouthparts and hot wax is quickly melted over it to keep it in place. (C) A hole cut into the eye where the pink-white photoreceptor cell layer can be seen. Black pigment and yellow hemolymph are absent. (D) Lateral view of the specimen with a glass recording electrode (1) placed in the eye, and the indifferent electrode (2) attached to the head stage, which should complete a circuit as seen on the oscilloscope. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53829/53829fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
 
<img alt="Figure 6" src="/files/ftp_upload/53829/53829fig6.jpg" style="font-size: 14px;" />
Figure 6:&#160;Raw Responses from Sample Recordings. Each response corresponds to a single light flash of 50 msec duration (black bars). (A) An example of the large negative voltage change that should be seen just before entering a cell. (B) A clean recording should have little background noise and a large depolarizing response, typically of at least 40 mV. (C) An example of a poor recording due to the negative potential change after the main peak (arrowheads). (D) Another example of a bad recording. The resting potential is undergoing large fluctuations (red bar) and the large amount of background noise can obscure the amplitude of response (arrowhead).
<img alt="Figure 7" src="/files/ftp_upload/53829/53829fig7.jpg" /> 
Figure 7:&#160;Response-Intensity Log-Linear Function. Solid circles show the measured responses of a cell from 3.5 to 0 OD, for this experimental setup. Light intensity is on a logarithmic scale. At very high intensities the response is saturated, and at very low intensities a small response persists instead of dropping to zero along the line. The Naka-Rushton equation33 is fitted to this non-linear shape (dotted line).
<img alt="Figure 8" src="/files/ftp_upload/53829/53829fig8.jpg" /> 
Figure 8:&#160;Spectral Sensitivity Examples. (A) A single representative cell&#39;s responses were recorded and relative spectral sensitivity was calculated. The peak of this cell is at 440 nm, meaning it responds best to blue light. Single cell data may look noisy (peak at 380 nm). (B) Cells with the same relative peak and shape are averaged together and standard error bars are added. Here, seven blue cells from seven individuals were averaged providing strong evidence that a cell type exists in this species maximally sensitive to light at 440 nm. (C) This process can be repeated for all cell types found, and plotted together. Insect eyes vary widely in their spectral sensitivities but a typical insect may have peaks shown here, at 370 nm, 440 nm, and 510 nm. Note, these spectral sensitivities are all calculated using real data, but the peaks have been shifted because the data is not yet published for this species.

Watch this Scientific Journal Video about Determination of Photoreceptor Cell Spectral Sensitivity in an Insect Model from In Vivo Intracellular Recordings at JoVE.com

Determination of Photoreceptor Cell Spectral Sensitivity in an Insect Model from In Vivo Intracellular Recordings

The electrophysiological technique of intracellular recording is demonstrated and used to determine spectral sensitivities of single photoreceptor cells in the compound eye of a butterfly.

Determination of Photoreceptor Cell Spectral Sensitivity in an Insect Model from In Vivo Intracellular Recordings

Intracellular recording is a powerful technique used to determine how a single cell may respond to a given stimulus. In vision research, intracellular ...

determination-photoreceptor-cell-spectral-sensitivity-an-insect-model

Nanyang Technological University

Research

JoVE Journal

Neuroscience

11.3K Views.  University of California, Irvine. The electrophysiological technique of intracellular recording is demonstrated and used to determine spectral sensitivities of single photoreceptor cells in the compound eye of a butterfly.

Video: Determination of Photoreceptor Cell Spectral Sensitivity in an Insect Model from In Vivo Intracellular Recordings

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

Single-unit In vivo Recordings from the Optic Chiasm of Rat

Information about the visual world is transmitted to the brain in sequences of action potentials in retinal ganglion cell axons that make up the optic nerve. In vivo recordings of ganglion cell spike trains in several animal models have revealed much of what is known about how the early visual system processes and encodes visual information. However, such recordings have been rare in one of the most common animal models, the rat, possibly owing to difficulty in detecting spikes fired by small diameter axons. The many retinal disease models involving rats motivate a need for characterizing the functional properties of ganglion cells without disturbing the eye, as with intraocular or in vitro recordings. Here, we demonstrate a method for recording ganglion cell spike trains from the optic chiasm of the anesthetized rat. We first show how to fabricate tungsten-in-glass electrodes that can pick up electrical activity from single ganglion cell axons in rat. The electrodes outperform all commercial ones that we have tried. We then illustrate our custom-designed stereotaxic system for in vivo visual neurophysiology experiments and our procedures for animal preparation and reliable and stable electrode placement in the optic chiasm.

Single-unit In vivo Recordings from the Optic Chiasm of Rat

Retinal ganglion cells transmit visual information from the eye to the brain with sequences of action potentials. Here, we demonstrate how to record the action potentials of single ganglion cells in vivo from anesthetized rats.

Information about the visual world is transmitted to the brain in sequences of action potentials in retinal ganglion cell axons that make up the optic ...

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from individual neurons are critical for understanding how neural circuits function under natural conditions. Traditionally, these recordings have been performed 'blind', meaning the identity of the recorded cell is unknown at the start of the recording. Cellular identity can be subsequently determined via intracellular1, juxtacellular2 or loose-patch3 iontophoresis of dye, but these recordings cannot be pre-targeted to specific neurons in regions with functionally heterogeneous cell types. Fluorescent proteins can be expressed in a cell-type specific manner permitting visually-guided single-cell electrophysiology4-6. However, there are many model systems for which these genetic tools are not available. Even in genetically accessible model systems, the desired promoter may be unknown or genetically homogenous neurons may have varying projection patterns. Similarly, viral vectors have been used to label specific subgroups of projection neurons7, but use of this method is limited by toxicity and lack of trans-synaptic specificity. Thus, additional techniques that offer specific pre-visualization to record from identified single neurons in vivo are needed. Pre-visualization of the target neuron is particularly useful for challenging recording conditions, for which classical single-cell recordings are often prohibitively difficult8-11. The novel technique described in this paper uses retrograde transport of a fluorescent dye applied using tungsten needles to rapidly and selectively label a specific subset of cells within a particular brain region based on their unique axonal projections, thereby providing a visual cue to obtain targeted electrophysiological recordings from identified neurons in an intact circuit within a vertebrate CNS.
The most significant novel advancement of our method is the use of fluorescent labeling to target specific cell types in a non-genetically accessible model system. Weakly electric fish are an excellent model system for studying neural circuits in awake, behaving animals12. We utilized this technique to study sensory processing by "small cells" in the anterior exterolateral nucleus (ELa) of weakly electric mormyrid fish. "Small cells" are hypothesized to be time comparator neurons important for detecting submillisecond differences in the arrival times of presynaptic spikes13. However, anatomical features such as dense myelin, engulfing synapses, and small cell bodies have made it extremely difficult to record from these cells using traditional methods11, 14. Here we demonstrate that our novel method selectively labels these cells in 28% of preparations, allowing for reliable, robust recordings and characterization of responses to electrosensory stimulation.

Retrograde Fluorescent Labeling Allows for Targeted Extracellular Single-unit Recording from Identified Neurons In vivo

Retrograde transport of fluorescent dye labels a sub-population of neurons based on anatomical projection. Labeled axons can be visually targeted in vivo, permitting extracellular recording from identified axons. This technique facilitates recording when neurons cannot be labeled through genetic manipulation or are difficult to isolate using 'blind' in vivo approaches.

The overall goal of this method is to record single-unit responses from an identified population of neurons. In vivo electrophysiological recordings from ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

Design and Assembly of an Ultra-light Motorized Microdrive for Chronic Neural Recordings in Small Animals

The ability to chronically record from populations of neurons in freely behaving animals has proven an invaluable tool for dissecting the function of neural circuits underlying a variety of natural behaviors, including navigation1 , decision making 2,3, and the generation of complex motor sequences4,5,6. Advances in precision machining has allowed for the fabrication of light-weight devices suitable for chronic recordings in small animals, such as mice and songbirds. The ability to adjust the electrode position with small remotely controlled motors has further increased the recording yield in various behavioral contexts by reducing animal handling.6,7
 Here we describe a protocol to build an ultra-light motorized microdrive for long-term chronic recordings in small animals. Our design evolved from an earlier published version7, and has been adapted for ease-of use and cost-effectiveness to be more practical and accessible to a wide array of researchers. This proven design 8,9,10,11 allows for fine, remote positioning of electrodes over a range of ~ 5 mm and weighs less than 750 mg when fully assembled. We present the complete protocol for how to build and assemble these drives, including 3D CAD drawings for all custom microdrive components.

The design, fabrication and assembly of an ultra-light motorized microdrive is described. The device provides a cost-effective and easy-to-use solution for chronic recordings of single units in small behaving animals.

The ability to chronically record from populations of neurons in freely behaving animals has proven an invaluable tool for dissecting the function of ...

Dual Electrophysiological Recordings of Synaptically-evoked Astroglial and Neuronal Responses in Acute Hippocampal Slices

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs through activation of their ion channels and neurotransmitter receptors, and process information in part through activity-dependent release of gliotransmitters. Furthermore, astrocytes constitute the main uptake system for glutamate, contribute to potassium spatial buffering, as well as to GABA clearance. These cells therefore constantly monitor synaptic activity, and are thereby sensitive indicators for alterations in synaptically-released glutamate, GABA and extracellular potassium levels. Additionally, alterations in astroglial uptake activity or buffering capacity can have severe effects on neuronal functions, and might be overlooked when characterizing physiopathological situations or knockout mice. Dual recording of neuronal and astroglial activities is therefore an important method to study alterations in synaptic strength associated to concomitant changes in astroglial uptake and buffering capacities. Here we describe how to prepare hippocampal slices, how to identify stratum radiatum astrocytes, and how to record simultaneously neuronal and astroglial electrophysiological responses. Furthermore, we describe how to isolate pharmacologically the synaptically-evoked astroglial currents.

The preparation of acute brain slices from isolated hippocampi, as well as the simultaneous electrophysiological recordings of astrocytes and neurons in stratum radiatum during stimulation of schaffer collaterals is described. The pharmacological isolation of astroglial potassium and glutamate transporter currents is demonstrated.

Astrocytes form together with neurons tripartite synapses, where they integrate and modulate neuronal activity. Indeed, astrocytes sense neuronal inputs ...

Simultaneous Whole-cell Recordings from Photoreceptors and Second-order Neurons in an Amphibian Retinal Slice Preparation

One of the central tasks in retinal neuroscience is to understand the circuitry of retinal neurons and how those connections are responsible for shaping the signals transmitted to the brain. Photons are detected in the retina by rod and cone photoreceptors, which convert that energy into an electrical signal, transmitting it to other retinal neurons, where it is processed and communicated to central targets in the brain via the optic nerve. Important early insights into retinal circuitry and visual processing came from the histological studies of Cajal1,2 and, later, from electrophysiological recordings of the spiking activity of retinal ganglion cells - the output cells of the retina3,4.
A detailed understanding of visual processing in the retina requires an understanding of the signaling at each step in the pathway from photoreceptor to retinal ganglion cell. However, many retinal cell types are buried deep in the tissue and therefore relatively inaccessible for electrophysiological recording. This limitation can be overcome by working with vertical slices, in which cells residing within each of the retinal layers are clearly visible and accessible for electrophysiological recording.
Here, we describe a method for making vertical sections of retinas from larval tiger salamanders (Ambystoma tigrinum). While this preparation was originally developed for recordings with sharp microelectrodes5,6, we describe a method for dual whole-cell voltage clamp recordings from photoreceptors and second-order horizontal and bipolar cells in which we manipulate the photoreceptor's membrane potential while simultaneously recording post-synaptic responses in horizontal or bipolar cells. The photoreceptors of the tiger salamander are considerably larger than those of mammalian species, making this an ideal preparation in which to undertake this technically challenging experimental approach. These experiments are described with an eye toward probing the signaling properties of the synaptic ribbon - a specialized synaptic structure found in a only a handful of neurons, including rod and cone photoreceptors, that is well suited for maintaining a high rate of tonic neurotransmitter release7,8 - and how it contributes to the unique signaling properties of this first retinal synapse.

We describe the preparation of thin retinal slices from aquatic tiger salamanders (Ambystoma tigrinum) and explain how we use these slices to study synaptic processing in the retina by obtaining dual whole-cell voltage clamp recordings from photoreceptors and second-order horizontal and bipolar cells.

One of the central tasks in retinal neuroscience  is to understand the circuitry of retinal neurons and how those connections are  responsible for shaping ...

A Simple and Inexpensive Method for Determining Cold Sensitivity and Adaptation in Mice

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life. The cold plantar assay allows the objective and inexpensive assessment of cold sensitivity in mice, and can quantify both analgesia and hypersensitivity. Mice are acclimated on a glass plate, and a compressed dry ice pellet is held against the glass surface underneath the hindpaw. The latency to withdrawal from the cooling glass is used as a measure of cold sensitivity.
Cold sensation is also important for survival in regions with seasonal temperature shifts, and in order to maintain sensitivity animals must be able to adjust their thermal response thresholds to match the ambient temperature. The Cold Plantar Assay (CPA) also allows the study of adaptation to changes in ambient temperature by testing the cold sensitivity of mice at temperatures ranging from 30 &#176;C to 5 &#176;C. Mice are acclimated as described above, but the glass plate is cooled to the desired starting temperature using aluminum boxes (or aluminum foil packets) filled with hot water, wet ice, or dry ice. The temperature of the plate is measured at the center using a filament T-type thermocouple probe. Once the plate has reached the desired starting temperature, the animals are tested as described above.
This assay allows testing of mice at temperatures ranging from innocuous to noxious. The CPA yields unambiguous and consistent behavioral responses in uninjured mice and can be used to quantify both hypersensitivity and analgesia. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

The Cold Plantar Assay (CPA) measures cold responsiveness between 30 &#176;C and 5 &#176;C, and can also measure cold adaptation. This protocol describes how to use the CPA to measure cold hypersensitivity, analgesia, and adaptation in mice.

Cold hypersensitivity is a serious clinical problem, affecting a broad subset of patients and causing significant decreases in quality of life.  The cold ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.