Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques

Neuroscience

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Summary

We will demonstrate how to prepare retinal slices from the mouse eye and record light responses in retinal neurons. The entire procedure is conducted in dark-adapted conditions.

Cite this Article

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Hellmer, C. B., Ichinose, T. Recording Light-evoked Postsynaptic Responses in Neurons in Dark-adapted, Mouse Retinal Slice Preparations Using Patch Clamp Techniques. J. Vis. Exp. (96), e52422, doi:10.3791/52422 (2015).

Abstract

The retina is the gateway to the visual system. To understand visual signal processing mechanisms, we investigate retinal neural network functions. Retinal neurons in the network comprise of numerous subtypes. More than 10 subtypes of bipolar cells, ganglion cells, and amacrine cells have been identified by morphological studies. Multiple subtypes of retinal neurons are thought to encode distinct features of visual signaling, such as motion and color, and form multiple neural pathways. However, the functional roles of each neuron in visual signal processing are not fully understood. The patch clamp method is useful to address this fundamental question. Here, a protocol to record light-evoked synaptic responses in mouse retinal neurons using patch clamp recordings in dark-adapted conditions is provided. The mouse eyes are dark-adapted O/N, and retinal slice preparations are dissected in a dark room using infrared illumination and viewers. Infrared light does not activate mouse photoreceptors and thus preserves their light responsiveness. Patch clamp is used to record light-evoked responses in retinal neurons. A fluorescent dye is injected during recordings to characterize neuronal morphological subtypes. This procedure enables us to determine the physiological functions of each neuron in the mouse retina.

Introduction

The retina is one of the unique parts of the nervous system. As an accessible part of the brain, its synaptic architecture has been well characterized. In addition, the functions of this neural network can be examined with a physiological stimulus: light. If the retinal tissue is isolated in a dark room with appropriate procedures, neurons in the tissue will respond to light. This preparation has been used to study visual signal processing and elucidate various synaptic mechanisms and neural network functions, as well as disease mechanisms.

Light responses in retinal neurons have been recorded for decades. Early studies used sharp electrodes to make intracellular recordings from mudpuppy retinal neurons1. In the 1980s, the patch clamp technique was invented2, and soon became a popular method among vision researchers3,4. Single cell recordings from lower vertebrates, including mudpuppy and fish retinal neurons, were popular methods that contributed to the elucidation of visual signal processing mechanisms5,6.

After genetic mutation techniques were developed, the mouse retina became a more popular model for vision researchers7-9. The mammalian retina is more attractive than that of lower vertebrates because it is evolutionarily closer to the human retina, and there is an opportunity to use disease models. However, mouse retinal cells are small and fragile10, and making retinal preparations and conducting patch clamp recordings in a dark room is challenging. As technology has improved, diverse approaches have become available to study visual signaling mechanisms such as imaging studies11 and the electroretinogram (ERG)12. Nevertheless, single cell recording with the patch clamp method is still important because it is highly temporally and spatially sensitive compared to other methods. Therefore, we have continuously conducted patch clamp recordings and improved our methods to investigate visual signal processing in mouse retinal slice preparations13-15.

In this video tutorial, the protocols are presented with important tips. Good recordings can only be achieved with good preparation. Practicing animal dissection and building a sturdy patch clamp rig will enable most researchers to achieve successful recordings.

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Protocol

Ethics Statement: Procedures involving animal subjects were approved by the Institutional Animal Care and Use Committee (IACUC) at Wayne State University.

1. Preparation of Experimental Solution

  1. Prepare the dissecting solution 1 day to 1 week before the actual experiment. Use a HEPES buffer solution for retinal dissection because of its strong buffering ability at lower temperatures16. Mix all chemicals as follows (in mM): 115 NaCl, 2.5 KCl, 2.5 CaCl2, 1.0 MgCl2, 10 HEPES, 28 glucose. Adjust pH to 7.4 with NaOH. Keep the solution in a refrigerator up to 1 week.
  2. The recording solution is Ames’ medium, which is an artificial cerebral spinal fluid (aCSF) designed for retinal preparations17. By the day of the experiment, weigh out Ames’ powder (4.4 g for 500 ml solution) in a tube. Also, weigh out NaHCO3 (0.95 g for 500 ml solution) and mix with the Ames’ powder.
  3. Prepare the intracellular pipette solution for patch clamp recordings by the day of the experiment. Mix all chemicals as follows (in mM): 111 K-gluconate, 1.0 CaCl2, 10 HEPES, 1.1 EGTA, 10 NaCl, 1.0 MgCl2, 5 ATP-Mg, and 1.0 GTP-Na. Adjust pH to 7.2 with KOH. Filter the pipette solution with a conventional syringe filter. Make ~500 µl aliquots and store them in a freezer or a deep freezer.

2. Preparation for the Day of Experiment

  1. The night before the experiment, place a mouse in a cage (C57BL/6J or similar background, 4 - 8 weeks old, male) in a dark space for O/N dark adaptation.
  2. Preparation for dissection:
    1. Fill dissecting solution (100 - 200 ml) in a glass beaker, place it on ice, and bubble with oxygen for at least 10 min.
    2. Start oxygenation of the dark box for retinal preparation storage.
    3. Prepare plastic coverslips with grease rails for retinal slice preparations and place each of them in a 35 mm plastic dish.
    4. Attach a new razor blade to a chopper (tissue slicer).
    5. Cut a filter membrane into two halves. This is for retinal slicing.
    6. Align the dissecting tools and transfer pipettes near the dissecting microscope. The work area must be well organized so that each tool can be easily accessed in the dark.
  3. Preparation for patch clamp recordings:
    1. Dissolve Ames’ powder and NaHCO3 in distilled water (Ames 4.4 g and NaHCO3 0.95g/500 ml ddH2O). Bubble the Ames’ medium with mixed oxygen (95% O2 and 5% CO2) for at least 30 min while heating to a recording temperature level (~35 °C). Adjust the pH to 7.4 with NaHCO3 while still bubbling.
    2. Thaw the internal pipette solution during the dissection. After it is thawed, add 0.01% sulforhodamine B for intracellular staining.

3. Retinal Dissection

  1. In a dark room, euthanize the mouse using carbon dioxide and bilateral pneumothorax. After 2 - 3 min of using the carbon dioxide, the mouse will lose consciousness.
  2. When the mouse no longer responds to a tail pinch, quickly enucleate the eyes and place them in a cold dissecting solution in a plastic dish. Perform the bilateral pneumothorax to ensure the animal’s death. Quickly store the eye in the dish in the dark box. Use an infrared viewer to conduct the procedure in the dark.
    NOTE: Alternatively, isoflurane, decapitation, or cervical dislocation can be used for euthanasia. The method of euthanasia should follow the guidelines of the National Institute of Health (NIH) and/or the Institutional Animal Care and Use Committee.
  3. In a dark room, adjust the dissecting microscope and infrared illumination. Under the microscope, transfer an eye from its dish into a 10 cm plastic dish with the cooled dissecting solution. Continuously bubble oxygen through tubing placed on the bottom of the plastic dish. Adjust the flow volume so it is not too low, but ensure that it is not high enough to disturb the dissection.
  4. Cornea and lens removal:
    1. Make an incision on the top of the cornea with a micro-surgical knife. Holding the optic nerve at the other end of the eyeball prevents the eye from moving. Extend the cut using a pair of surgical scissors. Cut out the cornea with some sclera around it.
    2. Grab the lens with a fine forceps and slowly pull the lens out. If the vitreous is too sticky to remove the lens, cut the vitreous with a pair of scissors.
    3. Once the eye-cup is made, gently pour cold, oxygenated dissecting solution into the eyecup with a small transfer pipette.
      NOTE: The eyeball quickly deflates after an incision is made in the cornea. It is important to maintain the shape of the eye as deforming the eye during this procedure will damage the retina and cause retinal detachment.
  5. Identifying the dorsal and ventral sides of the retina:
    NOTE: Because of uneven distribution of the green and the ultraviolet (UV) cones18, identifying a specific retinal area is important for light response recordings.
    1. Identify the line going across the retinal eye cup passing near the optic nerve head. This line passes mostly across the ventral side of the optic nerve; the side including the optic nerve is the dorsal side, while the other is ventral 19. See this landmark clearly under infrared viewers.
    2. Depending on the purpose of the experiment, make a cut on the unnecessary side. This will be useful after isolating the retina from the eye cup.
  6. Vitreous removal:
    1. Optionally, incubate the eye-cup with hyaluronidase (0.5 mg/ml) for 15 min.
    2. Remove the vitreous with extra-fine forceps. The vitreous is mainly attached to the bottom and the outer edge of the eye-cup. Gently remove it without touching or poking the retina. As the vitreous might be tightly attached to the eyecup, remove gently to avoid retinal detachment from the eye-cup. Continue to pull until no tension is felt.
      NOTE: This step is particularly important for dissecting the mouse retina; however, it is not a problem for the retinas of other species such as the rat or salamander.
  7. Isolate the retina from the eye-cup. Grab the sclera and gently peel off the retina using the backside of forceps.
    NOTE: Whole-mount retinal preparations can be made from the isolated retina. Make three to four cuts at the edges and flatten the retina, or cut the retina into several pieces.
  8. Trim the retina and discard the unnecessary half of the retina (step 3.5). In the same plastic dish, cut the remaining retina into two pieces. For each piece of the retina, cut off the edges to make a retinal slab, and trim the folding edges.
  9. Transfer a retinal slab onto a glass plate using a large transfer pipette (~2 ml). Suck up the excess solution with a small pipette, use a piece of filter paper to flatten the slab, and then quickly place a piece of the filter membrane on top of the tissue.
    1. Using a small transfer pipette, place a drop of dissecting solution on the filter membrane. Wait ~15 sec until the solution spreads over the membrane. During this time, the retinal tissue will stick to the filter membrane.
    2. Pour more cold dissecting solution under and around the filter. The retinal tissue with the filter membrane will float. Grab the filter membrane with a forceps and place it in the slicing chamber. Pour dissecting solution over the tissue, and store the preparation in the dark box.
      NOTE: The procedure described here is critical. It is important to perform these steps quickly to ensure that the retinal tissue sticks to the filter membrane and does not dry out.
  10. Using a chopper, cut the retinal tissue into slices. Around 200 μm thickness is feasible for patch clamp studies.
  11. Place a plastic coverslip with grease rails in the slicing chamber. Transfer a retinal slice on top of the grease rails on a plastic coverslip. Rotate the slice 90° so that the transverse section is visible. Press the filter membrane down onto the coverslip, then, cover the sides of the filter paper with grease.
    1. After the slice is immobilized on a plastic coverslip, grab the coverslip and transfer it to a 35 mm plastic dish. Pour cold dissecting solution onto the retinal preparation and submerge the preparation. Store each dish in the dark box, which should be continuously oxygenated.
      NOTE: The entire procedure needs to be done carefully so that the filter membrane is not deformed. Otherwise, the retinal slice easily separates from the filter membrane.

4. Patch Clamp Recordings from Retinal Slice Preparation

  1. Recording preparation:
    1. Prime the perfusion tubes with Ames’ medium. Allow all the bubbles to pass through when filling the tubing.
    2. Make patch clamp recording pipettes with a puller. To fill a pipette with the pipette solution, dip in the backside of a pipette for 1 min until the pipette tip is backfilled. Next, fill ~1/3 of the pipette using a micro-pipette filler. Store each pipette in a moist pipette box.
    3. Turn on the equipment for patch clamp recordings; including the computer, amplifier, CCD camera, and microscope.
  2. Shut off the room light. Place a retinal slice preparation onto the microscope stage chamber using an infrared viewer. After it is immobilized, begin continuous perfusion. Set the perfusion temperature at 33 to 37 °C.
  3. View the slice surface with the CCD camera. Focus on the target location where the target cell types reside. Select a healthy-looking cell for patch clamp recording (i.e., it should have smooth-looking surface and a good cell shape).
  4. Place a recording pipette in a pipette holder. Advance the pipette to the slice preparation. When it is close to the slice preparation (~2 mm above), find the tip of the pipette with the microscope. Once the pipette tip is visible under the microscope, move the tip down towards the target cell.
  5. Set the amplifier. Adjust the pipette at 0 mV/0 pA (zeroing) and start a continuous electric pulse of ~5 mV at ~10 Hz. Check the pipette resistance; ideal resistance is between 5 and 10 MΩ for most retinal neurons.
  6. Start blowing out from the tip. Use a mouthpiece, or use a syringe, to apply positive pressure when the pipette dips into the bath solution. Continuously blow out the internal solution until the tip is on the surface of the target cell.
    1. When the positive pressure makes a small dimple on the cell surface, advance the tip slightly and stop blowing out. Check the pipette resistance, which should increase. If it is continuously increasing, leave it and monitor the resistance until it reaches >1 GΩ (gigaseal). If the resistance does not spontaneously increase, gently apply negative pressure until it becomes a gigaseal.
  7. After the gigaseal is achieved, change the holding potential to -70 mV. Then, intermittently apply negative pressure to rupture the membrane inside the pipette tip. When the whole-cell configuration is made, the pipette resistance can be between 500 MΩ and 1 GΩ, and the capacitive current is seen. Sometimes spontaneous postsynaptic currents can be observed.
  8. Record the I-V relationship from -80 to +40 mV. Different types of voltage-gated channels are activated depending on the cell type.
  9. Record light-evoked synaptic currents or voltages.

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Representative Results

A representative slice preparation is shown in Figure 1. The slice preparation is in a straight position, showing photoreceptors to ganglion cells in a flat surface, and no detachment from the filter paper. If a slice is tilted, only part of the preparation is in focus, which makes it difficult to identify an appropriate cell for patch clamping. For recordings, it is important to select a good soma, which usually has a shiny surface, an even round shape, and no visible dark plaques. After the whole cell configuration was made, the slice was exposed to background light and then to a step-light (Figure 2). Light-evoked excitatory postsynaptic potentials (L-EPSPs) were evoked in response to a step-light (timing is shown in yellow). The peak amplitude and the decay time varied depending on the cell type and subtype. A ganglion cell generated action potentials in response to bright stimuli (Figure 2 D&E). The cell type and its morphological subtype were revealed by sulforhodamine labeling after physiological recordings (Figure 2, right).

Figure 1
Figure 1. Retinal slice preparations. (A) A retinal slice preparation viewed with a 10X objective. A retinal slice preparation (top) was attached to a piece of filter paper (bottom). (B) A four-image compilation showing a retinal slice preparation viewed with a 60X objective. Each cell layer is clearly observed (ONL: outer nuclear layer, OPL: outer plexiform layer, INL: inner nuclear layer, IPL: inner plexiform layer, GCL: ganglion cell layer) Please click here to view a larger version of this figure.

Figure 2
Figure 2. Light-evoked excitatory postsynaptic potentials (L-EPSPs) from retinal neurons. Step-light evoked L-EPSPs (left). The light intensity was 30 - 60% Weber contrast. Background light adaptation level was 4 x 104 photons/μm2/sec. Sulforhodamine B was injected during patch clamp recording to visualize the recorded neurons (right). A recording pipette was still attached to the soma (A) L-EPSPs and sulforhodamine staining from an ON cone bipolar cell. (B) OFF cone bipolar cell. (C) Amacrine cell. (D) ON ganglion cell. (E) OFF ganglion cell. Scale bar indicates 2 or 5 mV as noted. Light stimulation was applied for 1 sec. Please click here to view a larger version of this figure.

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Discussion

Good recordings can only be achieved with good retinal preparations and well-designed patch clamp setups. Although all the steps described above are important, the discussion highlights some critical steps both for the dissection and recordings.

For dissection, two things are especially important: cooling and oxygenation. After enucleating the eye, quickly remove the front part of the eye in a dissecting chamber with oxygen-bubbled, cooled dissecting solution, and pour cold solution into the eye-cup. When the retinal tissue is in the dissecting chamber, cold solution should be poured on the tissue every 10 min. The tissue should be sliced quickly without pulling or poking the retinal tissue during the dissection, which could easily damage its structure. Using good dissecting tools (sharp forceps and scissors) is also crucial. A chopper is equally as effective as a vibratome because it allows for speedy maneuvering. Extra-fine forceps should be used to remove vitreous matter as efficiently as possible because residual vitreous damages the retinal structure when slicing the tissue.

Beginners tend to damage the retina when removing the lens. The retina can detach from the eye-cup, or can even be accidentally removed with the lens. To avoid this issue, the sclera can be cut around the Ora Serrata which is located close to the extra-ocular muscle attachment point. An eye-cup with a wide opening makes it easier to remove the lens. Also, it is important not to deform the eye when removing the front portion. Another critical point is fixing the retinal slice onto a coverslip. This can be facilitated by practicing rotating the filter paper without distorting it to prevent the retinal slice from coming off during the experiment.

A sturdy, well-built rig is crucial for patch clamp recordings. It must be free of electric noise, vibration, and pipette drifting. In addition, light response can be recorded in dark conditions to prevent desensitization. Our cage is made with a metal frame and aluminum panels with black plastic drapes that act as an electric shield while also completely blocking light from entering the cage; computer monitor illumination will not affect the retinal preparation. Use a remote controller for the pipette manipulator and close the cage while setting up the patch clamp configuration, which also avoid illuminating the preparation.

Although we have been tried to simplify the procedures involved, patch clamp study is still a difficult experiment that requires extensive knowledge and training. An automated patch clamp has been developed by some groups20; this system is applicable to dissociated cells and might be an efficient technique for pharmacological screenings.

In summary, patch clamp recording in dark-adapted retinal slice preparations is a useful technique to investigate visual information processing. Compared to other methods, patch clamp offers better temporal and spatial resolution. Additional technical innovations will make the patch clamp method more user-friendly in the near future.

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Disclosures

The authors have nothing to disclose.

Acknowledgments

This work was supported by NIH R01 EY020533, WSU Startup Fund, and RPB grants.

Materials

Name Company Catalog Number Comments
Mice (28-60 days old, male) Jackson laboratory C57BL/6J strain
Ames' medium powder Sigma A1420 excellent
Stereo microscope Nikon SMZ745 excellent
Dissecting tool: forceps Dumont #4, #5, #55 excellent
Dissecting tool: scissors Roboz RS-5605 excellent
Dissecting tool: surgery knife Surgistar 7514 excellent
Razor blade (for chopper) EMS 71970 excellent
Chopper handmade
Infrared viewer Night Owl Optics NOBG1 It shows bright view. Focusing small objects is an issue.
Puller Sutter P-1000 excellent; makes consistent size pipettes.
Dark box Pelican dark box excellent
Patch clamp system Scientifica slice scope 2000 Excellent setup. Most key components are included in one package. Micromanipulators are excellent.
Amplifier Molecular Devices multiclamp 700B Excellent and easy control.
Acquiring software Molecular Devices pClamp software Excellent and easy control.
Light source (LED) Cool LED pE-2 4 channel system Excellent
CCD camera Q-imaging Retiga 2000 Excellent
Faraday cage handmade

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References

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