Electroretinogram Analysis of the Visual Response in Zebrafish Larvae

Summary

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.

Abstract

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.

Introduction

The electroretinogram (ERG) is a noninvasive electrophysiological method that has been used extensively in the clinic for determining the function of the retina in humans. The electrical activity in response to a light stimulus is measured by placing recording electrodes on the outer surface of the cornea. The characteristics of the stimulus paradigm and the response waveform define the retinal neurons contributing to the response. This method has been adapted for use with a number of animal models including mice and zebrafish. The typical vertebrate ERG response has four principal components: the a-wave, which is a cornea-negative potential derived from photoreceptor cell activity; the b-wave, a cornea-positive potential derived from the ON bipolar cells; the d-wave, a cornea-positive potential interpreted as the activity of the OFF bipolar cells; and the c-wave, which occurs several seconds after the b-wave and reflects activity in Müller glia and the retinal pigment epithelium^1-4. Additional references for understanding the history and principles of ERG analysis in humans and model animals are the online textbook, Webvision, from the University of Utah and texts such as the Principles and Practice of Clinical Electrophysiology of Vision^4,5.

Daniorerio (zebrafish) has long been favored as a model for vertebrate development, due to its rapid maturation and transparency, which allows for noninvasive morphological analysis of organ systems, behavioral assays and both forward and reverse genetic screens (for review, see Fadool and Dowling⁶). Zebrafish larvae are highly amenable to genetic and pharmacological manipulation, which, when coupled with their high fecundity, make them an excellent animal model for high-throughput biological analyses. The higher ratio of cones to rods in larval zebrafish – roughly 1:1 compared to mice (~3% cones) – make them particularly useful for the study of cone function^7-9.

In the vertebrate retina, cones develop before rods¹⁰. Interestingly, zebrafish cones are operative as early as 4 dpf, allowing for selective electrophysiological analysis of cones at that stage^6,11,12. In contrast, ERG responses in rods appear between 11 and 21 dpf¹³. Therefore, zebrafish larvae at 4-7 dpf serve functionally as an all-cone retina. However, the native photopic ERG response of 4-7 dpf larvae is dominated by the b-wave. Application of pharmacological agents, such as L-(+)-2-amino-4-phosphono-butyric acid (L-AP4), an agonist for the metabotropic glutamate (mGluR6) receptor expressed by the ON bipolar cells, effectively blocks the generation of the b-wave and reveals the isolated cone mass receptor potential, (the “a-wave”)^14-17.

Here we describe a simple and reliable method for ERG analysis using commercially available ERG equipment designed for use with mice that have been adapted for use with zebrafish larvae. This system can be utilized on zebrafish larvae of varying genetic backgrounds, as well as those treated with pharmacological agents, to aid researchers in the identification of signaling pathways that contribute to visual sensitivity and light adaptation¹⁶. The experimental procedures outlined in this protocol will guide investigators in the use of ERG analysis to answer a variety of biological questions pertaining to vision, and demonstrate the construction of a flexible ERG setup.

Protocol

Animal upkeep and experimental protocols were approved by the Institutional Animal Care and Use Committees of the University of North Carolina at Chapel Hill, and meet all requirements of the NIH Office of Laboratory Animal Welfare and the Association for Assessment and Accreditation of Laboratory Animal Care International.
NOTE: To obtain larvae for ERG analysis, published protocols for standard zebrafish husbandry and maintenance were employed¹⁸. Larvae are obtained through natural breeding and housed under a 14 hr light/10 hr dark cycle. This protocol has been optimized for larvae at 5-7 days post-fertilization (dpf), but could ideally be performed on older fish with small modifications to the procedure. Here, use the TL strain of wild-type zebrafish larvae at 5 dpf.

1. Micropipette Production

1. Pull several micropipettes using 1.5 x 0.86 mm (outer diameter by inner diameter) fire-polished borosilicate glass capillaries with filament (melting temperature, 821 °C) and a P-97 Flaming/Brown Micropipette Puller fitted with box heat filament. Use the program for fashioning micropipettes described in Table 1.
2. Check each micropipette under a microscope with an appropriate graticule ruler to ensure that tips are 10-15 µm in diameter and has a smooth tip opening (i.e., no jagged edges).
3. Carefully store micropipettes to prevent tip damage and exposure to dust. Storage options include Petri dishes with lab tape, foam-lined boxes, or commercially available micropipette storage containers.
   NOTE: Other micropipette pullers and glass capillaries can be used as long as the correct diameter of the micropipette and a high-quality tip is achieved.

Pressure	Heat	Pull	Velocity	Time
500	560	-	30	200
500	450	-	30	200
500	410	55	40	200

Table 1: Program for the production of micropipettes using a P-97 Flaming/brown Micropipette Puller fitted with a box heat filament. Micropipettes are made using 1.5 x 1.0 mm² (outer diameter by inner diameter) fire-polished borosilicate glass capillaries with filament (melting temperature, 821 °C).

2. Buffer Preparation

1. Use filtered, oxygenated goldfish Ringer's buffer¹⁹ in the microelectrode capillary and to saturate the polyvinyl alcohol (PVA) sponge onto which the larvae are placed for experiments. Alternatively, use E3 embryo media or Hank's Balanced Salt Solution.
2. Prepare 10x goldfish Ringer's solution as described in Table 2. Adjust to pH 7.8, and sterilize using a 0.22 µm filter and store the 10x stock at 4 °C.
3. Create a working solution on the day of the experiment by diluting the 10x Ringer's solution to 1x with deionized, distilled water. Filter using a 0.22 µm filter system. Oxygenate by bubbling with 95% O₂/5% CO₂ gas for 10 minutes. Cap tightly afterwards to ensure that the solution remains oxygenated.

NaCl	1.25 M
KCl	26 mM
CaCl2	25 mM
MgCl2	10 mM
glucose	100 mM
HEPES	100 mM

Table 2: Preparation of 10x goldfish Ringer’s solution.

3. Electroretinogram Platform

1. Perform ERG experiments on an anti-vibration table inside a Faraday cage to improve the signal to noise ratio. Attach a custom steel platform to the anti-vibration table using hex nuts. Place a movable plastic platform with a viscoelastic urethane polymer shock-absorbing bottom on the table under the light source.
2. Position the camera with a magnetized stand, aimed down at the movable plastic platform. Position the micromanipulator (which will hold the recording microelectrode) with a second magnetized stand to the right of the movable plastic platform. Ensure that the camera and micromanipulator will not be disturbed by the movement of other equipment and that they do not block illumination from the light source.
3. Connect the camera to a video monitor and position it to view the eye of the larva for placing the electrode in the proper position.
4. Ensure that the setup is properly grounded with copper wire. To check the noise, place the reference electrode and tip of the recording microelectrode in a 35 mm Petri dish filled with Ringer's solution. Check the electrical noise levels of the setup with an oscilloscope or a built-in feature of the ERG apparatus. Noise levels should be no more than ±10 µV from baseline.

4. Sponge Preparation

1. Cut a small rectangle of dry PVA sponge that will fit snugly in a 35 mm Petri dish. The thickness of the sponge should not be greater than the depth of the dish. Use a utility knife with a clean razor blade for cutting.
2. Make an additional cut into the sponge to accommodate the reference electrode (either a shallow cut lengthwise on the bottom of the sponge or a butterfly cut vertically through one of the smaller ends).
3. Use a chemically resistant marker to mark a small dot on the sponge (where the larva will be placed) that can be used for positioning the camera.
4. Soak the PVA sponge in Ringer's solution until saturated. Remove and blot quickly on a paper towel 2-3 times. Place the sponge in a clean 35 mm Petri dish.
5. Position the Petri dish containing the sponge on the plastic platform such that the mark can be visualized by the camera.

5. Electrode Preparation

NOTE: The zebrafish setup consists of a reference electrode in contact with the Ringer's solution-saturated PVA sponge and a recording electrode in contact with the cornea. The reference electrode consists of an Ag/AgCl pellet. The recording electrode is a pulled glass micropipette filled with Ringer's solution and held by a microelectrode holder containing an Ag wire.

1. Chloride the electrodes by soaking them in 6-9% sodium hypochlorite (bleach) for 5 min (the recording microelectrode wire) or 15 min (the reference electrode). Air dry on a Kimwipe for 5 min.
   1. Depending on the style of cut made in Step 4.2, place the Ag/AgCl pellet of the reference electrode into (for the vertical butterfly cut) or under (for the shallow cut lengthwise on the bottom) the sponge. Attach the reference electrode lead to the recording system.
   2. Alternatively, if the ERG setup has space constraints or there are particularly strong photovoltaic artifacts from the Ag/AgCl electrode, connect the reference electrode to the sponge via an agar salt bridge to move the electrode out of the light path.
2. Attach ~40 cm of appropriately sized tubing to a 5 ml non-Luer lock syringe. Fill the syringe with Ringer's solution. Microelectrode holders possessing pressure ports typically ship with adaptors to accommodate tubing with inner diameters of 1/16”, 3/32”, 1/8” or 5/32”.
3. Fill a 1 ml non-Luer lock syringe with Ringer's solution and, using a Micro-fil, carefully fill the microelectrode holder. Prevent the formation of bubbles.
4. Attach the 5 ml syringe to the pressure port of the microelectrode holder with tubing and use it to ensure that the microelectrode holder is full of Ringer's solution. Using the Micro-fil and 1-ml syringe filled with Ringer's solution, fill the micropipette glass from the tip and ensure that no bubbles are present.
5. Attach the glass micropipette to the microelectrode holder, being careful to keep the electrode wire straight. Once secured, use the 5 ml syringe to carefully force Ringer's solution through the microelectrode until a tiny amount of solution is visible on the tip. Occasional application of pressure to the syringe (when not applied to cornea) will prevent formation of air bubbles, as well as occlusions due to dust or salt accumulation, in the micropipette tip.
   1. If the solution comes out as a stream, replace the glass micropipette, as the tip opening is too large or is damaged.
6. Carefully place the recording microelectrode in the micromanipulator and attach the lead to the recording system.

6. Electroretinogram Analysis

NOTE: Due to the cone dominance of the larval retina, high quality ERG results can be obtained when preparations for recording are performed under low levels of indirect white light (<1 lux) or with short periods (<1 min) of higher intensity (≤250 lux) working light. A short period of dark adaptation is still required prior to recording (see step 6.7). However, experiments can be performed under dim red or infrared light using an infrared-sensitive camera. All experiments were performed in filter-sterilized (0.22 µm) system water from the UNC Zebrafish AquaCulture Facility but alternative embryo media can be used.

1. Cut paper towel squares measuring approximately 1 cm².
2. If measuring isolated cone mass receptor potential, incubate 3-5 larvae in system water with 500 µM (±)-2-Amino-4-phosphonobutyric acid (APB) for 5 min.
   NOTE: While APB is a racemic mixture of the active (L) and inactive (R) forms of AP4, it is as effective as L-AP4 and is less expensive.
3. Anesthetize 3-5 larvae in system water with 0.02% (w/v) Tricaine until unresponsive, about 1-2 min.
4. Use a Pasteur pipette and pipette pump to carefully transfer individual larvae onto paper towel squares under a dissecting stereoscope using minimal illumination (≤250 lux for <1 min). Check the position of each larva and choose a candidate that is dorsal side up with an unoccluded eye.
   1. For extended recordings (>30 min), keep the larva moist by glazing the body up to but not including the head with 3% methylcellulose using a fine camel-hair brush.
5. Using forceps, transfer the paper towel square with the larva to the damp PVA sponge.
   1. For extended recordings (>30 min), apply a continuous stream of water-saturated 100% O₂ gas over the larva by bubbling the gas through an airstone in a side-arm flask containing distilled water. Position the tubing exiting the flask side-arm that conveys the humidified oxygen near the larva's head.
      NOTE: Step 6.4.1 and step 6.5.1 will prolong the life of the fish¹⁶.
6. Under minimal illumination, use the micromanipulator and camera to position the microelectrode tip at the midpoint between the nasal and caudal ends of the eye and gently press onto the dorsal limit of the cornea.
   NOTE: Misplacement of the electrode tip to the far distal areas of the cornea can result in ERG waveforms of reversed polarity.
7. Allow larva to dark-adapt for 5-10 min.
8. Record test flash responses to light provided from an LED light source or optical stimulator using the available stimulation and recording equipment. Adjust protocol parameters such as flash intensity, flash length, flash color, background intensity and color and filter settings to fit the experiment.
9. When finished with the experiment, euthanize larvae according to AVMA/IACUC guidelines.

Results

Typically, ERGs are recorded from zebrafish larvae at 5 dpf, since a number of studies have published ERG recordings at this stage^9,16,20. Larval responses were measured under dark-adapted conditions with no background illumination using a 20 msec stimulus of white LED light. We utilized a commercially available ERG system consisting of a Ganzfeld light stimulator and computer controller/recorder. The stimulator uses a tightly controlled proprietary pulse width modulation (PWM) system to control the...

Discussion

In this protocol a simple procedure for ERG recordings of larval zebrafish is detailed. This procedure allows for a quick and comprehensive assay of visual function.There are several critical steps throughout the procedure that should be kept in mind. The zebrafish larvae should be healthy before the experiment to prevent death during potential drug treatments and ensure prolonged livelihood during the ERG recordings. In addition, it is important that the larvae utilized in experiments are closely age-matched. This is du...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Faraday cage	80/20 Inc	custom	Custom designed aluminum "Industrial Erector Set" for Cage framework
PVA sponge	Amazon	B000ZOWG1C	Provides a soft, moist platform for placement of zebrafish larvae
150 ml Sterile Filter systems	Corning	431154	Filtering solutions to prevent small articulates from blocking micropipettes
Espion E2	Diagnosys, LLC	contact	Modular electrophysiology system capable of generating visual stimuli for any stimulator and digital recording and analysis of responses using propietary software, more information at http://www.diagnosysllc.com
Colordome	Diagnosys, LLC	contact	Light stimulator with RGB LED and Xenon light sources for Ganzfeld ERG, more information at http://www.diagnosysllc.com
Micromanipulator	Drummond	3-000-024-R	Holding and positioning the recording microelectrode
Magnetic ring stand	Drummond	3-000-025-MB	Holding and positioning of the camera and refrence electrode
Lead extensions	Grass Technologies	F-LX	Spare female to male 1.5 mm lead cables for connecting electrodes
Male Pin to Female SAFELEAD Adaptor	Grass Technologies	DF-215/10	Connecting 2 mm pins to 1.5 headboard pins
Window screen frame (metal) and spline	Lowes or Home Depot	various	For attaching copper mesh to Faraday cage framework
Steriflip 50 ml filters	Millipore	SCGP00525	Filtering solutions to prevent small articulates from blocking micropipettes
BNC adaptor	Monoprice	4127	Connecting camera to BNC cable
BNC cable	Monoprice	626	Connecting camera to video adaptor
Camera lens	Navitar	1582232	Visualizing the positioning of the recording microelectrode onto the larval cornea
Camera coupler	Navitar	1501149	Visualizing the positioning of the recording microelectrode onto the larval cornea
Luna BNC to VGA + HDMI Converter	Sewell	SW-29297-PRO	BNC to VGA adaptor allowing camera image to project on computer monitor
APB	Sigma	A1910	mGluR6 agonist, blocks b-wave allowing analysis of the isolated cone mass receptor potential
Borosilicate glass	Sutter	BF-150-86-10	Fire- polished borosilicate glass (metling temperature = 821°C) with filament and dimensions of 1.5mm x 0.86 mm (outer diameter by inner diameter)
P97 Flaming/Brown puller	Sutter	P97	For pulling glass micropipettes
Sorbothane sheet	Thorlabs	SB12A	Synthetic viscoelastic urethane polymer, placed under Passive Isolation Mounts and ERG platform to absorb shock and prevent slipping, can be cut to size
Breadboard	Thorlabs	B2436F	Vibration isolation platfrom for ERG stimulator and zebrafish specimen
Passive Isolation Mounts	Thorlabs	PWA074	Provides vibration isolation to breadboard
Copper mesh	TWP	022X022C0150W36T	To line Faraday Cage
Pipette pump	VWR	53502-233	Used with Pasteur pipettes to carefully transfer zebrafish larvae
Pasteur pipettes	VWR	14672-608	Used with Pipette pump to carefully transfer zebrafish larvae
Camera	Watec	WAT-902B	Visualizing the positioning of the recording microelectrode onto the larval cornea
Tricaine (MS-222)	Western Chemical	Tricaine-S	Pharmaceutical-grade anesthetic,
Micro-fil	WPI	MF28G-5	Filling microelectrode holder and microelectrode glass
Microelectrode holder	WPI	MEH2SW15	Holds glass microelectrode, connects to ERG equipment
Reference Electrode	WPI	DRIREF-5SH	Carefully break off last centimeter of casing to drain electrolyte and expose sintered Ag/AgCl pellet electrode
Reference Electrode (alternative)	WPI	EP1	Alternative to DRIREF-5SH. Ag/AgCl electrode that must be wired/soldered to connecting lead
Low-noise cable for Microelectrode holder	WPI	13620	Connecting recording microelctrode holder to adaptor/headboard