This work was supported by DGAPA-UNAM IN224616-RN224616 grant. The authors want to thank Mrs. Josefina Bolado, Head of the Scientific Paper Translation Department, from Divisi&#243;n de Investigaci&#243;n at Facultad de Medicina, UNAM, for editing the English-language version of this manuscript.

NOTE: The experiments comply with the Laws of Animal Protection of Mexico.
1. Experimental Setup
<ol>
	<li>General connections
	<ol>
		<li>Connect the amplifier to a suitable computer through an analog-to-digital converter and use an oscilloscope to monitor the experiment (Figure 1).</li>
		<li>Connect the photostimulator to the A/D converted.</li>
	</ol>
	</li>
	<li>Recording chamber
	<ol>
		<li>Place the recording chamber on top of an anti-vibration table and locate it inside a Faraday cage. 
		NOTE: This prevents mechanical vibration and electrical noise that may affect the recording. Our Faraday cage was made of darkened wire mesh. A homemade, 2 mL, acrylic recording chamber is used (Figure 2).</li>
		<li>Prepare the bath solution to keep the preparation alive13: 205 mM NaCl; 5 mM KCl; 2 mM MgSO4; 13 mM CaCl2; 5 mM Hepes-NaOH, pH 7.3. 
		&#8203;NOTE: Solution does not include dextrose or any bubbling by 95% O2, 5% CO2 mixture because the experimental procedure is short enough to detect damage by the electrical recording (see step 4.2.3 for completeness).</li>
	</ol>
	</li>
	<li>Electrodes
	<ol>
		<li>Use a chloride-coated silver-wire electrode as the reference electrode 
		NOTE: Silver chloride coating is necessary to allow the electrode reaction: AgCl(s) + e&#8722;&#8596; Ag(s) + Cl-.</li>
		<li>Sand a silver wire (thickness ~ 1 mm, length ~ 10 cm) with a fine grade emery paper in order to clean its surface.</li>
		<li>Rinse the silver wire with distilled water.</li>
		<li>Immerse the clean silver-wire in a 4-6% sodium hypochlorite solution (NaClO) until it appears dark (approximately 20 min).</li>
		<li>Use thin glass pipettes filled with an electrolyte solution (2.7 M KCl) as the intracellular electrode.</li>
		<li>Pull a glass capillary tube (internal diameter &#8776; 1 mm) with a micro-pipette puller to obtain a thin tip with a small opening (0.01-0.1 &#181;m)14.</li>
		<li>Fill the pulled capillary glass with a 2.7 M KCl solution. First fill it by capillarity (immerse the pipette tip into the 2.7 M KCl solution) and then fill the half of the pipette with a fine injection needle. If necessary, tap the electrode pipette to eliminate air bubbles.</li>
		<li>Connect the electrode to its holder. Connect the holder to the amplifier headstage with the amplifier. Position the electrode-holder-headstage with a stable 3-D micromanipulator. Lower the electrode until bath solution covers its tip. 
		NOTE: The electrode must have a vertical orientation</li>
	</ol>
	</li>
	<li>Select the Bridge Mode of the amplifier (in the Mode section of the amplifier, press the Bridge button) and measure the electrode resistance. Make sure that the resistance is about 50 M&#937;11. 
	&#8203;NOTE: This electrode size allows a good-quality electrical recording with limited damage to the cell.</li>
	<li>Null the offset current and compensate capacitive transients using the Holding Position button in the section Voltage Clamp of the amplifier, and the Capacitance Neutralization button in Microelectrode 1 section of the amplifier.</li>
	<li>Superfusion system
	<ol>
		<li>Pour the bath solution (step 1.2.2) into a suitable receptacle (a 250 mL serum bottle) and connect it to an irrigation tubing set (3.2 mm of internal diameter), thus connecting the recording chamber. Use a gravity driven superfusion system. Regulate the flow rate to ~ 0.5 mL/s.</li>
		<li>Connect the chamber to a suction device. Regulate the solution suction system in such a way that the total volume of the recording chamber does not vary. Use a vacuum pump for suction. 
		NOTE: A constant volume in the recording chamber is important to keep stray capacitances constant throughout the experiment.</li>
	</ol>
	</li>
</ol>
2. Biological Material
Note: Use adult crayfishes P. clarkii (7-10 cm long) in the intermolt stage of indistinct sex.
<ol>
	<li>Circadian time
	<ol>
		<li>One month prior to the experiments, maintain 100 crayfishes under a 12-h light/12-h dark cycle15 (white light, 2.4 kW/m2). 
		NOTE: One month is sufficient time to synchronize the crayfish population.</li>
		<li>Determine the circadian time of the crayfish population by assessing the amplitude of electroretinograms of five randomly chosen animals15. 
		NOTE: 0 h circadian time (CT 0) indicates the beginning of a subjective day, i.e., time during which an organism is normally active16,17,18,19 (Figure 3). The crayfish is a nocturnal animal, so under a 12-h light/12-h dark cycle, it is active during the dark phase.</li>
	</ol>
	</li>
	<li>Eyestalk isolation procedure
	<ol>
		<li>At the desired circadian time, anesthetize a selected crayfish by immersion in tap water at 0-4 &#176;C, for 15 min.</li>
		<li>By means of a fine scissor, detach the eyestalk from the base.</li>
		<li>Access the retina using a razor blade to make an opening (~ 1-mm2) in the dorsal cornea.</li>
		<li>Place the eyestalk in the center of the recording chamber (the hole covered with silicone) with the access opening to the retina on the top of the chamber.</li>
	</ol>
	</li>
	<li>Keep the eyestalk under constant darkness for 20 min. 
	NOTE: 20 min is sufficient time for the photoreceptor cell to be fully dark adapted.</li>
</ol>
3. Photoreceptor Impaling
<ol>
	<li>Put the microelectrode and the eyestalk&#39;s longitudinal axis parallel in such a way that the microelectrode is centered for access to the retina. Use a stereoscopic microscope (10X) to place the devices in the correct configuration.</li>
	<li>Monitor the voltage difference between the reference and recording electrodes by selecting the Bridge Mode11 of the amplifier.</li>
	<li>Lower the electrode into the bath and thereafter place it right over the retina.</li>
	<li>Move away the microscope and position the photostimulator lamp parallel to the eyestalk&#39;s longitudinal axis. 
	NOTE: Distance between lamp and superfusion chamber is approximately 15 cm.</li>
	<li>Slowly lower the microelectrode until a sudden voltage drop is detected. 
	NOTE: A voltage drop of around &#8722;50 mV indicates the impalement of a healthy photoreceptor cell.</li>
	<li>Deliver a test light-flash (white light, 7.2 kW/m2, 10 &#181;s duration) to record the photoreceptor&#39;s receptor potential. 
	NOTE: The photoreceptor&#39;s receptor potential is a depolarizing potential of 10-15 mV amplitude and 300 ms duration (Figure 4).</li>
</ol>
4. Electrical Recording
<ol>
	<li>Ensure that the photoreceptor cells are totally adapted to dark conditions (see step 2.3).
	<ol>
		<li>Deliver a test-light flash every 2 min. Automate the time between flashes by choosing an adequate value (120 s) of &#34;time between episodes&#34; in the data acquisition software. 
		&#8203;NOTE: 2 min is the necessary time to allow the total recovery of the currents underlying the photoreceptor&#39;s electrical response.</li>
		<li>Monitor the resting membrane potential, as well as the amplitude and duration of the receptor potential. Assume that the photoreceptor is completely adapted once the membrane potential, amplitude, and duration of the receptor potential remain unchanged. Eyestalks previously kept under constant darkness for 20 min (step 2.3) complete their adaptation in approximately 5 min.</li>
		<li>Stop stimulating the cell 2 min before recording the light-elicited current.</li>
	</ol>
	</li>
	<li>Current recording
	<ol>
		<li>Clamp the voltage at the measured resting membrane potential value of the cell by selecting &#34;Holding amplitude&#34; in the data acquisition software (in Waveform on the Analog Output Channel section).</li>
		<li>Select the dSEVC mode of the amplifier. In the Mode section of the amplifier, select the SEVC button and switch the lever to the Discont SEVC position.</li>
		<li>Set the switching rate to 500-1,000 Hz (by using the Rate Adjust button of the amplifier), as determined by the speed of the electrode11.</li>
		<li>Deliver a light-flash, and observe the evoked ion influx. 
		NOTE: This is the light-elicited or transduction current (Figure 5).</li>
		<li>Return to the Bridge mode on the amplifier and record a receptor potential by sending a light flash (see section 3). Make sure that the photoreceptor cell is totally adapted to dark conditions. Measure the receptor potential characteristics (amplitude and duration) and compare with the first measurements. Assume that the impaled photoreceptor cell is a healthy cell if they have the same characteristics. 
		NOTE: After eyestalk ablation, biological preparation is viable during the following 2 h.</li>
	</ol>
	</li>
	<li>Two pulse protocol: Measure the recovery from desensitization with a two light-flash protocol. 
	&#8203;NOTE: The two light-flash protocol is similar to the standard two-pulse voltage protocol used to measure recovery from inactivation of voltage-gated channels20. The first light-flash causes a temporary change in the sensitivity of the photoreceptor cell and the second flash evaluates the state of the light-activated conductance.
	<ol>
		<li>Deliver a pair of light pulses. Apply the second flash after a desired time interval (from 300 ms to 2 min).</li>
		<li>Digitize the currents at 10 KHz sampling with the data acquisition software and save the data for off-line analysis11. 
		NOTE: A table with the configuration values of the software is included as supplementary material.</li>
	</ol>
	</li>
</ol>
5. Data Analysis
<ol>
	<li>Kinetics of the light-elicited current
	<ol>
		<li>Measure three current parameters: activation latency L, the time elapsed from the light-flash delivery until the current attains 10% of its maximal amplitude (Figure 5); peak or maximal current amplitude Ip; and desensitization time constant T. Measure the desensitization time constant (&#932;) by fitting the current decay phase to: 
		<img alt="Equation 1" src="/files/ftp_upload/56258/56258eq1.jpg" /> 
		where, A = -I(t=0) is a positive constant (Figure 5).</li>
	</ol>
	</li>
	<li>Desensitization and recovery 
	&#8203;Note: Recovery from desensitization was assessed as the ratio p2/p1, where p1 is the relevant parameter (either L, Ip, or T) of the control current, and p2 is the corresponding parameter of the second test-current.
	<ol>
		<li>Plot p2/p1 (either L, Ip, or T) as a function of time between pulses.</li>
		<li>Depending on the parameter, fit the points of each plot to: 
		<img alt="Equation 2" src="/files/ftp_upload/56258/56258eq2.jpg" /> 
		or to: 
		<img alt="Equation 3" src="/files/ftp_upload/56258/56258eq3.jpg" /></li>
		<li>Use an appropriate statistical test to determine the number of exponential terms required to fit the experimental data.</li>
	</ol>
	</li>
</ol>

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We have nothing to disclose.

The crayfish has proven to be an excellent model due to its ability to survive under non-natural conditions. There is easy access to in vivo and in vitro electrophysiological analyses. In addition, crustaceans are a favorable group for neurobiological research in the field of comparative chronobiology21.
In this paper, the study of desensitization and recovery of the light-activated transduction-current of crayfish photoreceptor cells is shown using th...

In order to be perceived as a visual stimulus, light reaching the eyes must be transduced into an electrical signal. Hence, in all visual organisms, light triggers a transduction ion-current, which in turn produces a change in the membrane potential of photoreceptor cells, the so-called receptor potential. Due to this, the light sensitivity of the eye primarily depends on the state of the light activated conductance, which can be either available to be activated or desensitized.
In crayfish photoreceptors, light triggers a slow, transient, ionic current1. Upon illumination, the transduction current arises after a lag or latency before reaching its maximum; thereafter it decays, as the transduction channels fall into a desensitized state in which they are unresponsive to further light stimulus2. That is, light, in addition to activating the transduction current responsible of vision, also induces a transient decrement of the sensitivity of photoreceptor cells. Desensitization may represent a general protective mechanism against overexposure to an adequate stimulus. The eye&#39;s sensitivity to light is recovered as the transduction conductance recovers from desensitization.
Intracellular recording is a useful technique for measuring electrical activity of excitable cells3,4,5,6,7,8. Although intracellular recording has become less frequent with the advent of the patch-clamp technique9, it is still a convenient approach when cells are either difficult to isolate, or present a geometry that makes the formation of the patch-clamping giga-seals difficult (i.e., seals or tight-contacts between the patch electrode and membranes with electrical resistance of the order of 109ohms). Examples of the latter are sperm cells10 and the photoreceptor cells herein studied. In our experience, Procambarus clarkii photoreceptors are difficult to isolate and keep in primary culture; additionally, they are thin rods that make giga-seal formation difficult to achieve. In intracellular recordings, a sharp electrode is advanced into a cell that is kept in place by the surrounding tissue. The electrode is chopped by the high-speed switching circuitry of the amplifier, so current is sampled between voltage pulses. This mode is known as discontinuous single-electrode voltage clamp (dSEVC mode)11. The high resistance (small opening) of the electrode hinders the diffusional exchange between the cell and the pipette solutions, yielding a minimal disturbance of the intracellular milieu3. A potential drawback of this technique is that electrode insertion may produce a non-selective leak current; therefore, care must be taken to avoid recording from cells where the size of the leak current may interfere with the intended measurements4,12.
Herein, we use isolated crayfish eyestalks to assess desensitization and recovery of the light-activated ion conductance by performing intracellular electrical recordings of photoreceptor cells under voltage clamp conditions.

<table><tbody><tr><td>Axoclamp2A&amp;nbsp;</td><td>Axon Instruments Inc</td><td></td><td>Amplifier</td></tr><tr><td>Digidata 1200 Interface</td><td>Axon Instruments Inc</td><td></td><td>Digitizer</td></tr><tr><td>Oscilloscope TDS430A</td><td>Tektronix</td><td></td><td>Analogic Oscilloscope</td></tr><tr><td>Photostimulator PS33 Plus</td><td>Grass</td><td></td><td>Lamp</td></tr><tr><td>Puller PC-100</td><td>Narishige</td><td></td><td>Micropipette Puller</td></tr><tr><td>Puller P-97</td><td>Sutter Instruments</td><td></td><td>Micropipette Puller</td></tr><tr><td>Glass Capillary Tube Kimax-51</td><td>Kimble Products</td><td>34502</td><td>0.8, 1.10, 100 mm</td></tr><tr><td>HS-2 Headstage</td><td>Axon Instruments Inc</td><td></td><td>Headstage</td></tr><tr><td>Micromanipulator MX-4</td><td>Narishige</td><td></td><td>Mechanical Micromanipulator</td></tr><tr><td>Stereoscopic Microscope</td><td>Zeiss</td><td></td><td>Microscope</td></tr><tr><td>pClamp</td><td>Axon Instruments Inc</td><td></td><td>Data acquisition software for digidata 1200 interface</td></tr><tr><td>Clampfit</td><td>Axon Instruments, Inc</td><td></td><td>Analysis software linked to pClamp</td></tr><tr><td>Origin</td><td>OriginLab Corp.</td><td></td><td>Data analysis and graphing software</td></tr><tr><td>Sodium Chloride</td><td>Sigma</td><td>S7653</td><td>&gt;99.5%</td></tr><tr><td>Potassium Chloride</td><td>Sigma</td><td>P-9333</td><td>Minimum 99%</td></tr><tr><td>Magnesium Sulfate</td><td>Sigma</td><td>M7506</td><td>Minimum 99.5%</td></tr><tr><td>Calcium Chloride</td><td>Sigma</td><td>C5080</td><td>Minimum 99.0%</td></tr><tr><td>Hepes</td><td>Sigma</td><td>H7523</td><td>&gt;99.5%</td></tr><tr><td>Sodium Hydroxide</td><td>Sigma</td><td>S8045</td><td>98.00%</td></tr><tr><td>Sodium hypochlorite solution</td><td>Sigma</td><td>425044</td><td>Available chlorine, 10-15%&amp;nbsp;</td></tr></tbody></table>

desensitization and recovery of crayfish photoreceptors upon delivery of a light stimulus

A method to study desensitization and recovery of crayfish photoreceptors is presented. We performed intracellular electrical recordings of photoreceptor cells in isolated eyestalks using the discontinuous single electrode-switched voltage-clamp configuration. First, with a razor blade we made an opening in the dorsal cornea to get access to the retina. Thereafter, we inserted a glass electrode through the opening, and penetrated a cell as reported by the recording of a negative potential. Membrane potential was clamped at the photoreceptor's resting potential and a light-pulse was applied to activate currents. Finally, the two light-flash protocol was employed to measure current desensitization and recovery. The first light-flash triggers, after a lag period, the transduction ionic current, which after reaching a peak amplitude decays towards a desensitized state; the second flash, applied at varying time intervals, assesses the state of the light-activated conductance. To characterize the light-elicited current, three parameters were measured: 1) latency (the time elapsed between light flash delivery and the moment in which current achieves 10% of its maximum value); 2) peak current; and 3) desensitization time constant (exponential time constant of the current decay phase). All parameters are affected by the first pulse.
To quantify recovery from desensitization, the ratio p2/p1 was employed versus time between pulses. p1 is the peak current evoked by the first light-pulse, and p2 is the peak current evoked by the second pulse. These data were fitted to a sum of exponential functions. Finally, these measurements were carried out as function of circadian time.

First, a representative receptor potential of crayfish photoreceptor cells is obtained (Figure 4). Afterwards, a test light-flash was applied to trigger the light transduction current (Figure 5). The cationic transduction current1 activates after a lag, reaching a maximal and thereafter slowly drops into an absorbing desensitized state from which it slowly recovers.
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Desensitization and Recovery of Crayfish Photoreceptors Upon Delivery of a Light Stimulus

A protocol for the study of desensitization and sensitivity recovery of crayfish photoreceptors as a function of circadian time is presented.

A method to study desensitization and recovery of crayfish photoreceptors is presented. We performed intracellular electrical recordings of photoreceptor ...

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6.6K Views.  Universidad Nacional Autónoma de México. A protocol for the study of desensitization and sensitivity recovery of crayfish photoreceptors as a function of circadian time is presented.

Video: Desensitization and Recovery of Crayfish Photoreceptors Upon Delivery of a Light Stimulus

Recording Behavioral Responses to Reflection in Crayfish

Social behavior depends on sensory input from the visual, mechanical and olfactory systems. One important issue concerns the relative roles of each sensory modality in guiding behavior. The role of visual inputs has been examined by isolating visual stimuli from mechanical and chemosensory stimuli. In some studies (Bruski &amp; Dunham, 1987: Delgado-Morales et al., 2004) visual inputs have been removed with blindfolds or low light intensity, and effects of remaining sensory modalities have been elucidated. An alternative approach is to study the effects of visual inputs in the absence of any appropriate mechanical and chemosensory cues. This approach aims to identify the exclusive role of visual inputs. 
We have used two methods to provide visual stimuli to crayfish without providing chemical and mechanical cues. In one method, crayfish are videotaped in an aquarium where half of the walls are covered in mirrors to provide a reflective environment, and the other half are covered in a non-reflective (matte finish) plastic. This gives the crayfish a choice between reflective and non-reflective environments. The reflective environment provides visual cues in the form of reflected images of the crayfish as it moves throughout half of the tank; these visual cues are missing from the non-reflective half of the tank. An alternative method is to videotape the behavior of crayfish in an aquarium separated by a smaller chamber at each end, with a crayfish in one small chamber providing visual cues and an inert object in the opposite small chamber providing visual input from a non-moving, non-crayfish source. 
Our published results indicate that responses of crayfish to the reflective environment depend on socialization and dominance rank. Socialized crayfish spent more time in the reflective environment and exhibited certain behaviors more frequently there than in the non-reflective environment; isolated crayfish showed no such differences. Crayfish that were housed in same-sex pairs developed a social rank of either dominant or subordinate. Responses to reflection differed between dominant and subordinate crayfish (May &amp; Mercier, 2006; May &amp; Mercier, 2007). Dominant crayfish spent more time on the reflective side, entered reflective corners more frequently and spent more time in reflective corners compared to the non-reflective side. Subordinate crayfish walked in reverse more often on the reflective side than on the non-reflective side. Preliminary data suggest similar effects from visual cues provided by a crayfish in a small adjoining chamber (May et al., 2008).

We have developed two methods for studying effects of visual cues on behavior in the absence of tactile and chemical cues. One method involves videotaping responses of crayfish to reflective walls in an aquarium; the other examines effects of visual inputs provided by a live crayfish behind a transparent partition.

Social behavior depends on sensory input from the visual, mechanical and olfactory systems.  One important issue concerns the relative roles of each ...

Membrane Potentials, Synaptic Responses, Neuronal Circuitry, Neuromodulation and Muscle Histology Using the Crayfish: Student Laboratory Exercises

The purpose of this report is to help develop an understanding of the effects caused by ion gradients across a biological membrane. Two aspects that influence a cell's membrane potential and which we address in these experiments are: (1) Ion concentration of K+ on the outside of the membrane, and (2) the permeability of the membrane to specific ions. The crayfish abdominal extensor muscles are in groupings with some being tonic (slow) and others phasic (fast) in their biochemical and physiological phenotypes, as well as in their structure; the motor neurons that innervate these muscles are correspondingly different in functional characteristics. We use these muscles as well as the superficial, tonic abdominal flexor muscle to demonstrate properties in synaptic transmission. In addition, we introduce a sensory-CNS-motor neuron-muscle circuit to demonstrate the effect of cuticular sensory stimulation as well as the influence of neuromodulators on certain aspects of the circuit. With the techniques obtained in this exercise, one can begin to answer many questions remaining in other experimental preparations as well as in physiological applications related to medicine and health. We have demonstrated the usefulness of model invertebrate preparations to address fundamental questions pertinent to all animals.

The experiments demonstrate an easy approach for students to gain experience in examining muscle structure, synaptic responses, the effects of ion gradients and permeability on membrane potentials. Also, a sensory-CNS-motor-muscle circuit is presented to show a means to test effects of compounds on a neuronal circuit.

The purpose of this report is to help develop an understanding of the effects caused by ion gradients across a biological membrane. Two aspects that ...

Light-mediated Reversible Modulation of the Mitogen-activated Protein Kinase Pathway during Cell Differentiation and Xenopus Embryonic Development

Kinase activity is crucial for a plethora of cellular functions, including cell proliferation, differentiation, migration, and apoptosis. During early embryonic development, kinase activity is highly dynamic and widespread across the embryo. Pharmacological and genetic approaches are commonly used to probe kinase activities. Unfortunately, it is challenging to achieve superior spatial and temporal resolution using these strategies. Furthermore, it is not feasible to control the kinase activity in a reversible fashion in live cells and multicellular organisms. Such a limitation remains a bottleneck for achieving a quantitative understanding of kinase activity during development and differentiation. This work presents an optogenetic strategy that takes advantage of a bicistronic system containing photoactivatable proteins Arabidopsis thaliana cryptochrome 2 (CRY2) and the N-terminal domain of cryptochrome-interacting basic-helix-loop-helix (CIBN). Reversible activation of the mitogen-activated protein kinase (MAPK) signaling pathway is achieved through light-mediated protein translocation in live cells. This approach can be applied to mammalian cell cultures and live vertebrate embryos. This bicistronic system can be generalized to control the activity of other kinases with similar activation mechanisms and can be applied to other model systems.

Light-mediated Reversible Modulation of the Mitogen-activated Protein Kinase Pathway during Cell Differentiation and Xenopus Embryonic Development

This protocol describes an optogenetic strategy to modulate mitogen-activated protein kinase (MAPK) activity during cell differentiation and Xenopus embryonic development. This method allows for the reversible activation of the MAPK signaling pathway in mammalian cell culture and in multicellular live organisms, like Xenopus embryos, with high spatial and temporal resolution.

Kinase activity is crucial for a plethora of cellular functions, including cell proliferation, differentiation, migration, and apoptosis. During early ...

Developmental Biology

Using the Horseshoe Crab, Limulus Polyphemus, in Vision Research

The American horseshoe crab, Limulus Polyphemus is one of the oldest creatures on earth, and the animal continues to play an indispensable role in biomedical research. Not only does their blood contain special cells that scientists use to detect bacteriotoxins in our medicines, but their eyes also contain a neural network that has provided much insight about physiological processes operating in our visual system, such as light adaptation and lateral inhibition. The horseshoe crab remains an attractive model for vision research because the animal is large and hardy for an invertebrate, its retinal neurons are big and easily accessible, its visual system is compact and extensively studied, and its visual behavior is well defined. Moreover, the structure and function of the eyes are modulated on a daily basis by a circadian clock in the animal s brain. In short, the visual system of horseshoe crabs is simple enough to be understood yet complex enough to be interesting.
In this video we present three electrophysiological paradigms for investigating the neural basis of vision that can be performed in vivo with Limulus. They are electroretinogram recording, optic nerve recording, and intraretinal recording. Electroretinogram (ERG) recordings measure with a surface electrode the summed electrical response of all cells in the eye to a flash of light. They can be used to monitor the overall sensitivity of the eye for prolong periods of time. Optic nerve recordings measure the spiking activity of single nerve fibers with an extracellular microsuction electrode. They can be used to study visual messages conveyed from the eye to the brain as well as circadian-clock messages fed back from the brain to the eye. Intraretinal recordings measure with an intracellular microelectrode the voltage fluctuations induced by light in individual cells of the eye. They can be used to elucidate cellular mechanisms of retinal processing.

Using the Horseshoe Crab, Limulus Polyphemus, in Vision Research

In this video we perform electroretinogram recording, optic nerve recording, and intraretinal recording with the American horseshoe crab, Limulus Polyphemus. These electrophysiological paradigms can be used for investigating the neural basis of vision in a research or teaching lab.

The American horseshoe crab, Limulus Polyphemus is one of the oldest creatures on earth, and the animal continues to play an indispensable role in ...

Electroretinogram Analysis of the Visual Response in Zebrafish Larvae

The electroretinogram (ERG) is a noninvasive electrophysiological method for determining retinal function. Through the placement of an electrode on the surface of the cornea, electrical activity generated in response to light can be measured and used to assess the activity of retinal cells in vivo. This manuscript describes the use of the ERG to measure visual function in zebrafish. Zebrafish have long been utilized as a model for vertebrate development due to the ease of gene suppression by morpholino oligonucleotides and pharmacological manipulation. At 5-10 dpf, only cones are functional in the larval retina. Therefore, the zebrafish, unlike other animals, is a powerful model system for the study of cone visual function in vivo. This protocol uses standard anesthesia, micromanipulation and stereomicroscopy protocols that are common in laboratories that perform zebrafish research. The outlined methods make use of standard electrophysiology equipment and a low light camera to guide the placement of the recording microelectrode onto the larval cornea. Finally, we demonstrate how a commercially available ERG stimulator/recorder originally designed for use with mice can easily be adapted for use with zebrafish. ERG of larval zebrafish provides an excellent method of assaying cone visual function in animals that have been modified by morpholino oligonucleotide injection as well as newer genome engineering techniques such as Zinc Finger Nucleases (ZFNs), Transcription Activator-Like Effector Nucleases (TALENs), and Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR)/Cas9, all of which have greatly increased the efficiency and efficacy of gene targeting in zebrafish. In addition, we take advantage of the ability of pharmacological agents to penetrate zebrafish larvae to evaluate the molecular components that contribute to the photoresponse. This protocol outlines a setup that can be modified and used by researchers with various experimental goals.

We present a method for the electroretinographic (ERG) analysis of zebrafish larvae utilizing micromanipulation and electroretinography techniques. This is a simple and straightforward method for assaying visual function of zebrafish larvae in vivo.

The electroretinogram (ERG) is a noninvasive electrophysiological method for determining retinal function. Through the placement of an electrode on the ...

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.

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.

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

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.

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

Electrophysiological Method for Whole-cell Voltage Clamp Recordings from Drosophila Photoreceptors

Whole-cell voltage clamp recordings from Drosophila melanogaster photoreceptors have revolutionized the field of invertebrate visual transduction, enabling the use of D. melanogaster molecular genetics to study inositol-lipid signaling and Transient Receptor Potential (TRP) channels at the single-molecule level. A handful of labs have mastered this powerful technique, which enables the analysis of the physiological responses to light under highly controlled conditions. This technique allows control over the intracellular and extracellular media; the membrane voltage; and the fast application of pharmacological compounds, such as a variety of ionic or pH indicators, to the intra- and extracellular media. With an exceptionally high signal-to-noise ratio, this method enables the measurement of dark spontaneous and light-induced unitary currents (i.e.&#160;spontaneous and quantum bumps) and macroscopic Light-induced Currents (LIC) from single D. melanogaster photoreceptors. This protocol outlines, in great detail, all the key steps necessary to perform this technique, which includes both electrophysiological and optical recordings. The fly retina dissection procedure for the attainment of intact and viable ex vivo isolated ommatidia in the bath chamber is described. The equipment needed to perform whole-cell and fluorescence imaging measurements are also detailed. Finally, the pitfalls in using this delicate preparation during extended experiments are explained.

Electrophysiological Method for Whole-cell Voltage Clamp Recordings from Drosophila Photoreceptors

Whole-cell recordings from Drosophila melanogaster photoreceptors enable the measurement of spontaneous dark bumps, quantum bumps, macroscopic responses to light, and current-voltage relationships under various conditions. In combination with D. melanogaster genetic manipulation tools, this method enables the study of the ubiquitous inositol-lipid signaling pathway and its target, the TRP channel.

Whole-cell voltage clamp recordings from Drosophila melanogaster photoreceptors have revolutionized the field of invertebrate visual transduction, ...

Long-range Channelrhodopsin-assisted Circuit Mapping of Inferior Colliculus Neurons with Blue and Red-shifted Channelrhodopsins

When investigating neural circuits, a standard limitation of the in vitro patch clamp approach is that axons from multiple sources are often intermixed, making it difficult to isolate inputs from individual sources with electrical stimulation. However, by using channelrhodopsin assisted circuit mapping (CRACM), this limitation can now be overcome. Here, we report a method to use CRACM to map ascending inputs from lower auditory brainstem nuclei and commissural inputs to an identified class of neurons in the inferior colliculus (IC), the midbrain nucleus of the auditory system. In the IC, local, commissural, ascending, and descending axons are heavily intertwined and therefore indistinguishable with electrical stimulation. By injecting a viral construct to drive expression of a channelrhodopsin in a presynaptic nucleus, followed by patch clamp recording to characterize the presence and physiology of channelrhodopsin-expressing synaptic inputs, projections from a specific source to a specific population of IC neurons can be mapped with cell type-specific accuracy. We show that this approach works with both Chronos, a blue light-activated channelrhodopsin, and ChrimsonR, a red-shifted channelrhodopsin. In contrast to previous reports from the forebrain, we find that ChrimsonR is robustly trafficked down the axons of dorsal cochlear nucleus principal neurons, indicating that ChrimsonR may be a useful tool for CRACM experiments in the brainstem. The protocol presented here includes detailed descriptions of the intracranial virus injection surgery, including stereotaxic coordinates for targeting injections to the dorsal cochlear nucleus and IC of mice, and how to combine whole cell patch clamp recording with channelrhodopsin activation to investigate long-range projections to IC neurons. Although this protocol is tailored to characterizing auditory inputs to the IC, it can be easily adapted to investigate other long-range projections in the auditory brainstem and beyond.

Channelrhodopsin-assisted circuit mapping (CRACM) is a precision technique for functional mapping of long-range neuronal projections between anatomically and/or genetically identified groups of neurons. Here, we describe how to utilize CRACM to map auditory brainstem connections, including the use of a red-shifted opsin, ChrimsonR.

When investigating neural circuits, a standard limitation of the in vitro patch clamp approach is that axons from multiple sources are often intermixed, ...

Electrophysiological Methods for Measuring Photopigment Levels in Drosophila Photoreceptors

The Drosophila G-protein-coupled photopigment rhodopsin (R) is composed of a protein (opsin) and a chromophore. The activation process of rhodopsin is initiated by photon absorption-inducing isomerization of the chromophore, promoting conformational changes of the opsin and resulting in a second dark-stable photopigment state (metarhodopsin, M). Investigation of this bi-stable photopigment using random mutagenesis requires simple and robust methods for screening mutant flies. Therefore, several methods for measuring reductions in functional photopigment levels have been designed. One such method exploits the charge displacements within the photopigment following photon absorption and the huge amounts of photopigment molecules expressed in the photoreceptors. This electrical signal, named the early receptor potential (or early receptor current), is measured by a variety of electrophysiological methods (e.g., electroretinogram and whole-cell recordings) and is linearly proportional to functional photopigment levels. The advantages of this method are the high signal-to-noise ratio, direct linear measurement of photopigment levels, and independence of phototransduction mechanisms downstream to rhodopsin or metarhodopsin activation. An additional electrophysiological method called prolonged depolarizing afterpotential (PDA) exploits the bi-stability of Drosophila photopigment and the absorption-spectral differences of fly R and M pigment states. The PDA is induced by intense blue light, converting saturating amounts of rhodopsin to metarhodopsin, resulting in the failure of light-response termination for an extended time in darkness, but it can be terminated by metarhodopsin to rhodopsin conversion using intense orange light. Since the PDA is a robust signal that requires massive photopigment conversion, even small defects in the biogenesis of the photopigment lead to readily detected abnormal PDA. Indeed, defective PDA mutants led to the identification of novel signaling proteins important for phototransduction.

Electrophysiological Methods for Measuring Photopigment Levels in Drosophila Photoreceptors

We present a protocol to electrophysiologically characterize bi-stable photopigments: (i) exploiting the charge displacements within the photopigment molecules following photon-absorption and their huge amount in the photoreceptors, and (ii) exploiting the absorption-spectra differences of rhodopsin and metarhodopsin photopigment states. These protocols are useful to screen for mutations affecting bi-stable photopigment systems.&#160;

The Drosophila G-protein-coupled photopigment rhodopsin (R) is composed of a protein (opsin) and a chromophore. The activation process of rhodopsin is ...

Desensitization and Recovery of Crayfish Photoreceptors Upon Delivery of a Light Stimulus

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Chapters in this video

Recording Behavioral Responses to Reflection in Crayfish

Membrane Potentials, Synaptic Responses, Neuronal Circuitry, Neuromodulation and Muscle Histology Using the Crayfish: Student Laboratory Exercises

Light-mediated Reversible Modulation of the Mitogen-activated Protein Kinase Pathway during Cell Differentiation and Xenopus Embryonic Development

Using the Horseshoe Crab, Limulus Polyphemus, in Vision Research

Electroretinogram Analysis of the Visual Response in Zebrafish Larvae

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

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

Electrophysiological Method for Whole-cell Voltage Clamp Recordings from Drosophila Photoreceptors

Long-range Channelrhodopsin-assisted Circuit Mapping of Inferior Colliculus Neurons with Blue and Red-shifted Channelrhodopsins

Electrophysiological Methods for Measuring Photopigment Levels in Drosophila Photoreceptors