Juxtacellular Monitoring and Localization of Single Neurons within Sub-cortical Brain Structures of Alert, Head-restrained Rats

Summary

This protocol describes the design and surgical implantation of a head-restraining mechanism to monitor neuronal activity in sub-cortical brain structures in alert rats. It delineates procedures to isolate single neurons in the juxtacellular configuration and to efficiently identify their anatomical locations.

Abstract

There are a variety of techniques to monitor extracellular activity of single neuronal units. However, monitoring this activity from deep brain structures in behaving animals remains a technical challenge, especially if the structures must be targeted stereotaxically. This protocol describes convenient surgical and electrophysiological techniques that maintain the animal’s head in the stereotaxic plane and unambiguously isolate the spiking activity of single neurons. The protocol combines head restraint of alert rodents, juxtacellular monitoring with micropipette electrodes, and iontophoretic dye injection to identify the neuron location in post-hoc histology. While each of these techniques is in itself well-established, the protocol focuses on the specifics of their combined use in a single experiment. These neurophysiological and neuroanatomical techniques are combined with behavioral monitoring. In the present example, the combined techniques are used to determine how self-generated vibrissa movements are encoded in the activity of neurons within the somatosensory thalamus. More generally, it is straightforward to adapt this protocol to monitor neuronal activity in conjunction with a variety of behavioral tasks in rats, mice, and other animals. Critically, the combination of these methods allows the experimenter to directly relate anatomically-identified neurophysiological signals to behavior.

Introduction

Monitoring neuronal activity in an alert animal actively engaged in a behavioral task is critical for understanding the function and organization of the nervous system. Extracellular recording of the electrical activity from single neuronal units has long been a staple tool of systems neuroscience and is still widely in use at present. A variety of electrode types and configurations are available depending on the scientific and technical demands of a particular experiment. Chronically implanted microdrives or electrode arrays are often used in freely moving animals, including birds, rodents, and non-human primates^1-4. Alternatively, acute penetrations with metal or glass microelectrodes via an external micromanipulator are often used to record from anesthetized or head-restrained animals. Glass micropipette electrodes have the advantage that they can be used in the juxtacellular or “cell attached” configuration to unambiguously isolate the activity of single neurons without the complications of post-hoc spike sorting⁵. These electrodes further permit recording from anatomically-identified cells or locations, as they can be used to inject small deposits of dye or neuroanatomical tracers, or even to fill the individual recorded cell. This configuration has been successfully applied in rats, mice and birds^6-10. The presently described technique focuses on juxtacellular monitoring and extracellular dye deposits in alert, head-restrained rats. Note that unlike single cell juxtacellular fills, these dye deposits do not provide information about cell morphology or axonal projections¹¹, but they enable exact anatomical localization to approximately 50 μm and, critically, have a significantly higher yield in alert animals. Information regarding single-cell juxtacellular fills is nonetheless provided as an alternative strategy for anatomical labeling.

In brief, the protocol consists of three major phases. In the first phase, the rat is acclimated to body restraint in a cloth sock (Figure 1) over a period of 6 days. In the second phase, a head restraint apparatus (Figure 2) and recording chamber are surgically implanted such that the rat can be maintained in the stereotaxic plane during multiple subsequent recording sessions (Figure 3); this procedure enables the experimenter to target particular sub-cortical regions of the brain for electrophysiological study based on standard reference coordinates¹². The third phase involves placing the rat in an appropriate jig for conducting the behavioral and electrophysiological experiments (Figure 4), constructing the electrode from a quartz capillary tube (Figure 5), making juxtacellular neuronal recordings that unambiguously isolate single units^6-9, and marking the anatomical location of the recording site with Chicago Sky Blue dye (Figures 6 and 7). The recordings are performed with simultaneous behavioral monitoring; however, the technical details of the behavior will depend on the scientific goals of each experiment and are thus beyond the scope of a single protocol. After completion of the experimental procedure, which can be repeated on multiple days, the animal is euthanized. The brain is extracted and processed according to standard neuroanatomical techniques using either bright field or fluorescence microscopy.

Protocol

Experimental protocols were carried out on female Long Evans rats (250 - 350 g) in accordance with federally prescribed animal care and use guidelines and were approved by the Institutional Animal Care and Use Committee at the University of California San Diego.

1. Acclimating the Rat to Body Restraint

NOTE: Place the rat on a restricted diet. Feed the rat once per day immediately after each daily handling session to acclimate the rat to the restraint (described below). Provide enough feed to maintain the animal at 80% of its initial weight. This amount is approximately 6 grams of feed per day for a 250 g female Long Evans rat.

1. Acclimate the rat to being handled by humans. Gently restrain the rat by hand for periods of approximately 5 sec every 30 sec. Do this for 15 min a day on 2 consecutive days. Monitor the rats daily for signs of stress during the training period. Signs include struggling in response to restraint, vocalization, and tooth-chattering.
2. On the 3^rd day, place the rat in a body-constraining cloth sock (Figure 1) for 15 min a day on two consecutive days.
3. On the 5^th day, place the rat inside the sock, and place the sock inside a rigid tube on the experimental jig (Figure 4). Leave the rat in the tube for 15 min a day on 2 consecutive days.
4. Feed the rat immediately after each training session. At the end of the last session (6^th day) give unrestricted access to food. Select for implantation only those rats which do not show signs of excessive stress on the last training day.
   NOTE: Though selection of rats for implantation is subjective, in our hands over 90% of rats were found to be responsive to habituation and able to undergo implantation.

2. Implanting the Recording Chamber and Head-restraint Mechanism

1. Make a reference wire by soldering bare stainless steel wire to a pin connector. Cut the wire so that 3-5 mm remains uncovered by the pin. Autoclave the reference wire, along with all surgical tools.
2. Administer ketamine/xylazine anesthesia. Administer 95 mg/kg ketamine mixed with 5 mg/kg xylazine intraperitoneally. The initial dose lasts for 1½ to 2 hr. Supplement the anesthesia every 30-45 min thereafter, as needed. Lubricate the animal’s eyes with an ophthalmic ointment.
3. Check the toe withdrawal reflex to determine the plane of anesthesia, and maintain anesthesia as necessary.
4. Place the rat in a stereotaxic holding frame with ear bars (Figure 3A). Shave the hair on the head and disinfect the wound site with povidone-iodine (10% solution). Take care to insert the ear bars appropriately so that they do not damage the ear drum. Ear bars with blunt tips are preferable.
5. Make an incision with a scalpel approximately along the midline of the cranium from the rostro-caudal level of the eyes to the back of the ears. Use scissors to cut and remove a 2-3 mm strip of skin on either side of the incision.
6. Scrape away the periosteum to expose the surface of the cranium out to the lateral ridges. Cover the exposed cranium with thin layer of superglue.
7. Drill a small hole, with a slightly smaller diameter than 0-80 screws, using a ½ mm diameter drill burr (see Materials section). Drill the hole immediately posterior to where the bregma suture meets the lateral ridge, and at a 30-45° angle into the cranium, so that the screw can go in with the top more lateral than the bottom.
8. Screw in a 0-80 flat-bottom machine screw to the hole (approximately 3 turns), at 30-45° angle. Be careful not to screw in too deeply to avoid damaging the underlying brain tissue.
9. Repeat this procedure for 6 additional screws in the configuration shown (Figure 3B). Apply superglue to the base of all screws.
10. Place a syringe needle in the stereotax manipulator. Measure from the bregma suture, and make a dent in the cranium with the needle near, but outside of the desired craniotomy location, as a reference mark.
    NOTE: In this demonstration, the stereotaxic coordinates of the mark are 3 mm posterior and 1 mm lateral to the bregma suture.
11. Mark the dent with a permanent marker. Note the stereotaxic coordinates of the mark, as it will serve as a stereotaxic reference point (Figure 3B).
12. Make a craniotomy centered on the desired coordinates (3 mm posterior and 3 mm lateral to bregma in this example). Leave the dura mater intact. Cover the craniotomy with modified artificial cerebral spinal fluid (125 mM NaCl, 10 mM glucose, 10 mM HEPES, 3.1 mM CaCl₂, 1.3 mM MgCl₂, pH 7.4)¹³ (Figure 3C).
13. Cut a 0.2 ml centrifuge tube to 4 - 5 mm in length, and cut off the cap. Place the tube on the cranium and center it over the craniotomy.
14. Apply dental cement around the bottom of the tube to seal the base of the tube to the cranium. Be careful not to leak cement into the exposed craniotomy (Figure 3D).
15. Drill another small hole (about ½ mm diameter) in the contralateral cranial plate and carefully insert the reference wire. Construct the reference wire by soldering a pin that interfaces with the recording amplifier to the end of the wire (see Equipment section). Do not move the reference wire once it is in the brain as this may cause damage.
16. Apply superglue to the hole in which the wire was inserted. This will seal the pin and wire in place temporarily.
17. Mix the two parts of the silicone gel kit (see Materials section) in approximately equal portions. Wait two minutes for mixing and fill the centrifuge tube approximately ⅓ full with gel.
18. Attach a right angle post clamp (see Materials section) to the head restraint bar. Attach the head plate (Figure 2A) to the holding bar (Figure 2B) at a 45° pitch angle so that the bottom of the plate is toward the nose of the animal. It is helpful to use an inclinometer to set the pitch angle to match that of the experimental jig (Figure 4).
19. Attach the holding bar to the stereotax manipulator arm so that the bar is parallel to the ear bars. Lower the bar and plate so that the plate is posterior of the lambda suture and anterior to the caudal-most screw.
20. Grasp the front head-bolt (an 8-32 stud or screw, see Materials section) at approximately 45° angle with a helping hands arm and clamp, so that the head of the screw faces down and towards the tail of the animal. Lower the screw so that it touches the anterior 0-80 screws (Figure 3F, G).
21. Secure the bolt and plate in place with dental acrylic. Apply dental acrylic around the bone screw heads, reference pin, and around the sides of the centrifuge tube. Wait approximately 10 min for the dental acrylic to dry (Figure 3H).
22. Apply a layer of dental cement around the edges of the dental acrylic, cementing the skin to the implant. Wait for the cement to dry.
23. Poke several holes in the cap of the centrifuge tube, and place the cap on the tube.
24. Remove the helping hands clamp. Then carefully remove the head-holding bar from the stereotax, and then remove head plate from the bar (Figure 3I).
25. Remove the animal from the stereotax and administer post-operative care and monitoring in accordance with all applicable rules, regulations, and laws (e.g., Buprenorphine 0.02 mg/kg, every 8-12 hr for a minimum of 24 hr). If inflammation develops around the edges of the implant, apply a thin layer of antibiotic ointment to the affected site daily until resolved.

3. Juxtacellular Monitoring of Neuronal Units

1. Pull quartz capillary tubing on a carbon dioxide laser micropipette puller (see Equipment section) with tip diameters of less than 1 μm (Figure 5A).
   NOTE: the heating parameters will vary according to the particular instrument, and that the required neck diameter will vary depending on the brain structure of interest. For recordings in rat thalamus, micropipettes have necks ranging from 5-7 mm.
2. Secure the pipette in place under a differential interference contrast (DIC) microscope equipped with long working distance objectives. Use modeling clay to hold the pipette in place on the microscope’s stage.
3. Slowly move a glass block (0.5 - 1 cm thick; a piece of glass used for making ultra-microtome knives is convenient) into the field of view with the pipette tip. Using the stage micromanipulator, gently touch the pipette tip to the glass, causing it to break. Repeat as necessary until the pipette tip outer diameter is between 1-3 μm (Figure 5B,C). Ensure that these micropipettes have impedances between approximately 5-15 MΩ.
4. Prepare extracellular saline (135 mM NaCl, 5.4 mM KCl, 1.8 mM CaCl₂, 1 mM MgCl₂, 5 mM HEPES, pH 7.2 with NaOH)⁸ and add 2% (w/v) Chicago Sky Blue. Using a syringe with a 30 G needle or smaller, fill the back end of the pipette with the solution. Alternatively, for single cell juxtacellular labeling add 2% (w/v) Neurobiotin or biotinylated dextran amine (BDA-3000 or BDA-10000) to the saline solution in place of Chicago Sky Blue.
5. Place the rat inside the cloth sock inside the rigid tube on an appropriate experimental jig (Figure 4).Wait a minimum of 72 hr after surgery before placing the animal on the head-restraint jig. Tighten the drawstring around the upper sternum, rather than the neck, to avoid obstruction of the airway. If breathing appears to be labored or obstructed, loosen the drawstring or move it posteriorly. Carry out this procedure while the rat is alert.
6. Attach the head-restraining plate on the rat to the corresponding piece on the jig. To do this, first pin the head-restraint plate in place, and then secure it using a 4-40 screw. Be sure to wait until the animal is calm in order to avoid applying excessive torque to the head. Next, place an 8-32 nut on the head-restraining bolt that is implanted on the rat. Then screw in a threaded stainless-steel rod to the head-bolt. Affix the rod to the experimental jig in such a way that it can be tightened in place (Figure 4).
   NOTE: Securing the animal with head-bolt as well as the head-restraint plate minimizes bending of the apparatus and improves recording stability. In most cases recording can commence on the first day the animal is head-restrained. However, if the animal fidgets excessively or vocalizes, provide one day of head-restraint habituation training before commencing any recordings. In this case, leave the animal on the jig for 15 min and then place it back in its cage. Repeat this step the following day and continue with the protocol.
7. Open the recording chamber and remove the silicone gel. Clean the craniotomy using fine forceps if tissue has re-grown in the craniotomy.
8. Attach the pipette to the motorized micromanipulator, and plug in the headstage pre-amplifier.
   NOTE: In the present demonstration, a small relay circuit on the headstage is used to switch between the amplifier lead wires and an external current source with high compliance (Figure 6A-C). Note that some amplifiers have an appropriate built-in high compliance source (see Discussion and Materials sections).
9. Use the motorized micromanipulator to move the tip of the pipette to the stereotaxically identified mark in the recording chamber. Note the coordinates of this location. Be careful not to break the tip of the pipette on the cranium.
10. Move the pipette to the desired recording location in the anterior-posterior and medio-lateral axes. Then advance the pipette ventrally through the dura until it is approximately 500 μm dorsal to the intended recording location.
11. Slowly advance the pipette while listening for spiking events on an audio monitor of the amplified voltage recorded between the pipette tip and the reference wire. Once spiking events are identified, continue to move the pipette 0-100 μm until positive going voltage deflections greater than approximately 500 μV are observed.
    NOTE: The electrode resistance will increase by a factor of approximately 1.5-10 when this occurs.
12. Begin recording once a unit has been isolated as according to step 3.11, along with all other behavioral and physiological measures of interest. In the present demonstration, self-generated vibrissa movements are monitored with high speed videography (see Materials section).
13. To label the recording site, switch the electrode leads to connect to the current source. There are several ways to do this, using either a relay circuit (Figure 6A-C), a built-in current source on the amplifier, or manually (Figure 6D, see step 3.8 and Discussion). Pass -4 μA with 2 sec pulses at 50% duty cycle for 4 min to iontophoretically inject Chicago Sky Blue through the pipette.
14. Kill and perfuse the animal after several labeling procedures according to standard practice. Section the brain and counterstain as necessary to identify the anatomical location of the recording site. Counterstain the tissue for cytochrome oxidase reactivity as per¹⁴. Alternatively, identify the Chicago Sky Blue deposits using fluorescence microscopy.
    NOTE: To accurately differentiate between different labeled units it is important that all labels are made with the same pipette on the same day, without unclamping the pipette between labels. In this case recording sites can be differentiated by their relative locations in post-hoc histology along with the noted manipulator coordinates of each site (see step 3.9). For unambiguous identification, mark the labels at least 200 μm apart from one another, and no more than 3-5 labels per brain region.

Results

Neuronal units in ventral posterior medial (VPM) thalamus encode the phase of vibrissa movement during self-generated whisking^15,16. Figure 7A shows sample spiking activity of a VPM thalamic unit as a rat is actively whisking. Figure 7B shows a histogram of spike times aligned to the instantaneous phase of vibrissa motion¹⁷. There are more spikes during the retraction phase of whisking. After the recording, the location of the unit was labeled via iontophoresis of C...

Discussion

Construction of the experimental jig

The description of the mechanical parts used to build the experimental jig (Figure 4) is omitted from the protocol, as it can be constructed in a variety of ways. In this demonstration standard opto-mechanical parts and support clamps are used to mount the head restraint bar and the body restraint tube (see Materials section). Similar opto-mechanical parts can be used to mount the electrode ho...

Materials

Name	Company	Catalog Number	Comments
Ketaset (Ketamine HCl)	Fort Dodge	N/A
Anased (Xylazine solution)	Lloyd Laboratories	N/A
Betadyne (Povidone-Iodine)	CVS Pharmacy	269281
Loctite 495	Grainger Industrial Supply	4KL86	20 - 40 cp cyanoacrylate
Vetbond	3M	1469SB
Grip cement powder	Dentsply Intl	675571	For the base of the recording chamber
Grip cement liquid	Dentsply Intl	675572	For the base of the recording chamber
Silicone gel	Dow Corning	3-4680
Jet denture repair acrylic powder	Lang Dental Manufacturing Co.	N/A	For securing the head restraint apparatus to the cranium
Ortho-Jet Fast curing orthodontic acrylic resin liquid	Lang Dental Manufacturing Co.	N/A	For securing the head restraint apparatus to the cranium
Chicago sky blue	Sigma	C8679
Paraformaldehyde	Sigma	158127	For perfusion and tissue fixation
Phosphate-buffered saline	Sigma	P3813	For perfusion and tissue fixation
Cytochrome C	Sigma	C2506	For cytochrome-oxidase staining, Figure 7
Diaminobenzidine	Sigma	D5905	For cytochrome-oxidase staining, Figure 7
Rat sock	Sew Elegant (San Diego, CA)	N/A	Custom made, Figures 1, 4
PVC tube 2 ½”	U.S. Plastic Co.	34108	Figure 4
Subminiature D pins & sockets	TE Connectivity	205089-1	Figure 3
Stainless steel music wire 0.010” diameter	Precision Brand Products, Inc.	21010	Figure 3
Stereotaxic holding frame	Kopf Instruments	Model 900	Figure 3
Stereotaxic ear bars	Kopf Instruments	Model 957	Figure 3
Stereotaxic manipulator	Kopf Instruments	Model 960	Figure 3
½ mm drill burr	Henry Schein	100-3995
Quiet-Air dental drill	Midwest Dental	393-1600
Stainless steel 0-80 ⅛” screw	Fastener superstore	247438	Figure 3
0.2 ml centrifuge tube	Fisher Scientific	05-407-8A	Figure 3
Custom head-holding bar	UCSD SIO Machine Shop	N/A	Custom made, Figures 2 - 4
Custom head-holding plate	UCSD SIO Machine Shop	N/A	Custom made, Figures 2 - 4
Right angle post-clamp	Newport	MCA-1	Figures 3, 4; standard opto-mechanical parts for the experimental jig (Figure 4) are also from Newport Corp.
8-32 ¾” screw	Fastener Superstore	240181	For head-restraint, Figure 3
4-40 ¼” screw	Fastener Superstore	239958	For head restraint, Figures 3, 4
Quartz capillary tubing	Sutter Instruments	QF-100-60-10	Figure 5
Carbon dioxide laser puller	Sutter instruments	P-2000
Motorized micromanipulator	Sutter Instruments	MP-285
Microelectrode amplifier	Molecular Devices	Multiclamp 700B	Alternate part: Molecular Devices Axoclamp 900A
Microelectrode amplifier head stage	Molecular Devices	CV-7B	Alternate part: HS-9Ax10 with Molecular Devices Axoclamp 900A
Isolated pulse stimulator	A-M Systems	Model 2100	Alternate part: HS-9Ax10 with Molecular Devices Axoclamp 900A
Audio monitor	Radio Shack	32-2040
Pipette holder	Warner Instruments	#MEW-F10T	Alternate parts: see Discussion, Figure 6A
Electrode lead wire	Cooner wire	NEF34-1646	(optional), Figure 6D
Relay for amplifier head-stage	COTO Technology	#2342-05-000	(optional) Used with a custom-made printed circuit board (UCSD Physics Electronics Shop), Figure 6A-C
Digital video camera	Basler	A602fm	(optional) For behavioral monitoring, Figure 7
Puralube vet ointment	Amazon.com, Inc	NC0138063