A Procedure for Implanting Organized Arrays of Microwires for Single-unit Recordings in Awake, Behaving Animals

Summary

Implanting organized arrays of microwires for use in single-unit electrophysiological recordings presents a number of technical challenges. Methods for performing this technique and the equipment necessary are described. Also, the beneficial use of organized microwire arrays to record from distinct neural subregions with high spatial selectivity is discussed.

Abstract

In vivo electrophysiological recordings in the awake, behaving animal provide a powerful method for understanding neural signaling at the single-cell level. The technique allows experimenters to examine temporally and regionally specific firing patterns in order to correlate recorded action potentials with ongoing behavior. Moreover, single-unit recordings can be combined with a plethora of other techniques in order to produce comprehensive explanations of neural function. In this article, we describe the anesthesia and preparation for microwire implantation. Subsequently, we enumerate the necessary equipment and surgical steps to accurately insert a microwire array into a target structure. Lastly, we briefly describe the equipment used to record from each individual electrode in the array. The fixed microwire arrays described are well-suited for chronic implantation and allow for longitudinal recordings of neural data in almost any behavioral preparation. We discuss tracing electrode tracks to triangulate microwire positions as well as ways to combine microwire implantation with immunohistochemical techniques in order to increase the anatomical specificity of recorded results.

Introduction

Electrophysiological recordings allow scientists to examine the electrical properties of biological cells. In the central nervous system, where electrical impulses serve as a signaling mechanism, these recordings are of particular importance for understanding neural function^1-2. During single-unit recordings in behaving animals, a microelectrode that has been inserted into the brain is able to record changes in a neuron's generation of action potentials over time.

While many techniques allow one to record brain activity, single-unit electrophysiology is one of the most precise methods by allowing resolution at the single neuron level. When a high degree of spatial specificity is desired, microwires can be used to target discrete sub-nuclei or ensembles of cells within the brain3. Single-unit recordings also benefit from high temporal resolution as recordings are accurate at the microsecond level. And, in vivo awake recordings allow intact circuit interactions, with the natural milieu of afferent and efferent projections, systemic chemical and hormonal influences, and physiological parameters. Neural signals are derived from sensory input, motor behaviors, cognitive processing, neurochemistry/pharmacology, or some combination. Accordingly, the segregation of sensory, motor, cognitive, and chemical influences necessitates well-conceived experiments with effective contingencies and controls that may allow for the assessment of each of the aforementioned influences. All in all, recordings in behaving animals allow experimenters to observe the integration of multiple sources of information within a functioning circuit and to derive a more comprehensive model of circuit function.

Single-unit recordings also suffer from a number of disadvantages of which any experimenter should be aware. First and foremost, recordings can be difficult to conduct. Indeed, properties of the headstage amplifiers and the implanted microwires that allow for spatial and temporal specificity in these recordings also makes recordings susceptible to the influence of extraneous electrical signals (i.e. electrical "noise"). Accordingly, the ability to troubleshoot problems in an electrophysiological system necessitates a well-developed technical understanding of electrophysiological principles and apparatus. It is also important to note that, under certain circumstances, recorded electrical signals in extracellular recordings can represent the summation of multiple neural signals. Moreover, the generalizability of single-unit activity to population activity within a target region can often be limited by the degree of cellular heterogeneity within the target region (but see Cardin⁴). For example, electrodes might be biased towards recording high amplitude output neurons in lieu of other cells. The interpretability of single-unit recordings is increased by combining recordings with other techniques including, but not limited to, electrical (orthodromic or antidromic), chemical (e.g. iontophoretic or designer receptor) or optogenetic stimulation⁴, temporary neural inactivations, sensorimotor examinations⁵, disconnection procedures, or immunohistochemistry³.

In the protocol that follows we will enumerate the materials and steps necessary to implant an organized microwire array in the rat (although the protocol can be adapted for use in other species). The procedure and style of fixed arrays used in our laboratory have proven reliable for longitudinal recordings and can sustain recordings of the same neuron for over one month's time ^6-8. This makes this procedure ideal for examining phasic responses to experimental stimuli, plastic changes in neural responses, or mechanisms of learning and motivation.

Protocol

The utmost care must be taken to maintain aseptic conditions (as described in the Guide for the Care and Use of Laboratory Animals⁹) while preparing for and conducting the following procedure. The following protocol is in compliance with the Guide for the Care and Use of Laboratory Animals and has been approved by the Institutional Animal Care and Use Committee, Rutgers University. It is estimated that the subsequent procedures will require 3-6 hr to complete.

Implanting the Microwire Array:

1. Place animals under anesthesia using 50 mg/kg sodium pentobarbital (i.p) and administer 10 mg/kg of atropine methyl nitrate (IP; Glycopyrrolate may be substituted) and 0.25 mg penicillin (300,000 U/ml i.m.) to maintain respiratory function and prevent infection, respectively.
   Note: With the 3-6 hr length of this implantation surgery, sodium pentobarbital is used because it is cost-effective and limits human exposure to anesthetics (as might occur with prolonged use of gas anesthetics) while still providing long-lasting anesthesia. Substitution of other anesthetics is acceptable.
2. Verify that the anesthesia has taken effect using the tail pinch test before proceeding.
3. As necessary, give alternating injections of ketamine hydrochloride (60 mg/kg IP) and sodium pentobarbital (5-10 mg/kg IP) to maintain anesthesia throughout the surgery.
4. Shave the scalp using a #22 scalpel blade.
5. Disinfect the shaved scalp with povidone iodine.
6. Give subcutaneous injections of bupivacaine (~1 mg/kg SC spread over 4 injection sites) to locally anesthetize the scalp. Allow 5-10 min for the local anesthetic to take effect.
7. Place an ophthalmic lubricant over the eyes to maintain moisture during anesthesia.
8. Secure the animal into the ear bars and nose clamp of a stereotaxic apparatus.
9. Use visual landmarks (e.g. a horizontal bar on the stereotaxic frame) to approximately level the animal's head. This step is only meant to approximately level the skull.
10. Make an incision along the midline of the scalp using a #11 scalpel blade mounted on a scalpel holder. The incision must extend from just behind the ears to the posterior portion of the nasal bone.
11. Using a dissection spatula, clear the skull of all remaining tissue until both lateral skull ridges and the posterior skull ridge have been reached.
12. Pull back the skin around the incision using a number of hemostats (6x).
13. Clean the skull of any blood and allow it to dry. If any remaining bleeding occurs, terminate the residual bleeding with a small cauterizing tool. Ensuring that the skull remains clean and dry allows the dental acrylic used in subsequent steps to bind to the skull permanently.
14. Mark bregma and lambda (Figure 2D) by interpolating the intersection of the skull sutures. A dissecting microscope is necessary to accurately mark these positions.
15. Attach a small pointed item (e.g. a pin or dental drill bit) to a stereotaxic arm and lower it to determine the dorsal/ventral (DV) coordinate of bregma and lambda.
16. Adjust the nose clamp until the DV coordinates for bregma and lambda are within 100 µm (0.1 mm) of each other.
17. Once leveled, record the anterior/posterior (AP), medial/lateral (ML) and DV coordinates of bregma along with the position of the nose clamp. Double check the coordinates for accuracy, as the remainder of the surgery depends on the precision of these coordinates.
18. Use the measured coordinates to calculate the ML and AP coordinates for the four corners of the "skull window" relative to bregma (i.e. craniotomy; Figure 1). The skull window is a precisely positioned rectangle through which the microwire array will pass.
19. Use the calculated coordinates to mark the skull window coordinates on the skull using a pointed stereotaxic attachment and fountain pen ink. To obtain precise marks, apply only a small amount of ink to the tip of the marking tool using a cotton applicator.
20. Drill out the skull window by removing the bone in a series of small layers.
    1. Start by drilling the marked corners of the window where the marks have been placed.
    2. Next, connect the corners and outline the window.
    3. Finally, clear out the area within the outline down to the depth of dura mater. The hole must be wider (i.e. beveled) below the superficial layers of the skull to ensure that the window maintains an appropriate width from top to bottom.
21. Use microforceps to carefully remove any remaining bone chips, debris, or dura mater inside the skull window. This step is extremely important, as these bits of material can compromise the integrity of the array during lowering. Once cleared, one must keep the window moist with bacteriostatic saline for the remainder of the surgery.
22. Place markings on the skull for 5 skull screws and 1 ground wire (Figure 1). These positions will vary depending on the targeted brain region. The headstage will be most secure if one screw is placed on each of the 5 skull bones. Place both the skull screws and ground wire in locations that will not interfere with the microwire array placement.
23. Drill holes for the skull screws and secure the screws in place. Screws must be lowered until only 3-4 threads are showing (or, if using screws other than those recommended, deep enough to traverse the thickness of the skull to promote array stability but not too deep so as to damage cortex). Clean the threads of the screws after placement.
24. Drill a hole for the ground wire. Lower the wire slowly (over 1-2 min) to the target DV coordinate and fix the wire in place using dental acrylic.
25. Add acrylic around the threads of the skull screws. Before the acrylic dries, remove any excess cement that flows away from the ground wire or skull screws. Allow 15 min for the dental acrylic to dry/harden.
26. Attach the array to the stereotaxic arm. Level the array and orient it so that it will squarely pass through the confines of the skull window.
27. Place saline in the skull window so that it is level with the skull. Lower the array until it creates a dimple in the saline. This coordinate is used as skull level for the array. Use this value to calculate the final DV coordinate of the array.
28. Lower the array slowly until it reaches its final DV coordinate. Stop every 1mm of lowering and wait for several minutes in order to allow brain tissue to recover from dimpling and to dissolve the bottom portion of the polyethylene glycol (PEG) on the array (PEG is used to temporarily keep the wires in their conformation while lowering, and dissolves slowly in saline).
29. When the array is 1mm away from the target, lower it more slowly until the final target is reached. Lowering at 0.1-0.2 mm at a time before allowing the tissue to rest for a period of 5 min is intended to preserve the tissue at the target site as well as proximal synaptic connections. If the equipment is available, precision lowering of the array can be assisted using a motorized manipulator.
30. Dissolve the remaining PEG and use dental acrylic to cement the microwires in place. Add multiple layers of cement to ensure that the wires are secure and then allow 15-20 min for the cement to harden before proceeding. This ensures that the wires will not be jostled from their final placement.
31. Build the remainder of the animal's headstage using dental acrylic. Use the acrylic to encase the skull screws, ground wire, and connector for the microwires. Allow the acrylic hat to harden sufficiently before proceeding.
32. Suture the scalp incision using absorbable sutures and administer 2 ml of bacteriostatic saline (s.c.) to restore hydration from surgery.
33. Remove the animal from the ear bars and place the subject in a clean area. Observe the animal frequently during the post-operative recovery until thermoregulation and locomotion have recovered. Following recovery from anesthesia, move the animal into single-housing for the remainder of post-surgical recovery.
34. Give animals daily postoperative monitoring and care in the days following the procedure. Recovery from this procedure is optimal when seven or more days are allowed.
35. Give animals injections of Carprofen (5mg/kg) and Enrofloxacin (5-10 mg/kg) or their equivalents during recovery per the schedule recommended by the attending veterinarian.

Results

A list of Equipment used by this laboratory for recording electrophysiological signals can be found in Table 3. Following recovery from surgery, single-units are recorded by plugging a unity-gain headstage into the implanted connector. This headstage is connected via a cable to a commutator, which is capable of free rotation without breaks in the electrophysiological recording through the use of electrical slip rings. The commutator allows subjects to freely move while recording during behavior, which is...

Discussion

Extracellular recordings represent a powerful experimental technique that can be incorporated into nearly any experimental preparation in neuroscience. Wires that have been implanted in organized arrays can be tracked as their shafts pass through the brain and into their target region (Figure 5A). When a small, post-experimental lesion is created at the noninsulated microwire tip to create a small iron deposit from the stainless steel wire, one can precisely mark the location of the uninsulated microwire...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Table 1. List of Surgical Materials.
Gauze	Fisher (MooreBrand)	19-898-144
Cotton Swabs	Fisher (Puritan)	S304659
Nembutal (Pentobarbital)	Sigma Aldrich	P3761
Atropine Methyl Nitrate	Sigma Aldrich	A0382
Baytril (Enrofloxacin)	Butler Shein (Bayer)	1040007
Ketamine Hydrochloride	Butler Shein	SKU# 023061
Betadine (Povidone-Iodine)	Fisher (Perdue)	19-066452
Stereotax	Kopf	Model 900
Cauterizing Tool	Stoelting	59017
Dissecting Microscope	Nikon	SMZ445
Dental Drill	Buffalo	37800
Bacteriostatic Saline	Bulter Schein	8973
Jewlers Screws	Stoelting	51457
Microwire Array	Microprobes	Custom (Flexible)
Ground Wire	Omnetics	Custom Plug
Dental Acrylic	Fisher (BAS)	50-854-402
Absorbable Sutures	Fisher (Ethicon)	NC0258473
Puralube (Opthalamic Ointment/Lubricant)	Fisher (Henry Schein)	008897
Table 2. List of Surgical Instruments.
2x Microforceps	George Tiemann Co.	#160-57	Multi-use (e.g. clearing debris in skull window)
2x Forceps	George Tiemann Co.	#160-93	Multi-use (e.g. tying sutures)
6x Hemostats	George Tiemann Co.	#105-1125	Clamp and open incision
1x Small scissors	George Tiemann Co.	#105-411	Cut sutures after tying
1x Tissue forceps	George Tiemann Co.	#105-222	Holding tissue while suturing
1x Needle holder	George Tiemann Co.	#105-1259	Holding suture needle
1x Scalpel holder (with #11 blade)	George Tiemann Co.	#105-80 (w/ #105-71 blade)	Making skull incision
1x #22 Scalpel blade	George Tiemann Co.	#160-381	Shaving scalp
1x Surgical Spatula	George Tiemann Co.	#160-718	Scraping skull to clear tissue on skull
Machine/Jewelers Screws	Various	N/A	0/80 x 1/8”
Table 3. List of Equipment for Recording Electrophysiological Signals.
Microwire Array & Connector	Micro Probe, Inc. (Gaithersburg, MD)	N/A (Part No. based on array characteristics)	Cranially implanted in target recording region. Arrays are customized based on desired wire spacing, length, etc.
Unity-Gain Harness/Headstage	M.B. Turnkey Designs (Hillsborough, NJ)	Proj 1200	Initial amplification of neural signal; allows for propagation of small neural signals.
Commutator (and Optional Fluid Swivel)	Plastics One, Inc. (Roanoke, VA)	SL18C	Allows animals to freely rotate while propagating electrical signal to preamp
Pre-amplifier	M.B. Turnkey Designs (Hillsborough, NJ)	Proj 1198	Differentially amplifies neural signals against a reference electrode.
Filter and Amplifier	M.B. Turnkey Designs (Hillsborough, NJ)	Proj 1199	Band-pass filters and further amplifies the differentially amplified signal.
Acquisition Computer	EnGen (Phoenix, AZ)	N/A (Custom Build)	Runs software and hardware for behavioral and neural data acquisition.
A/D Card	Data Translation (Marlboro, MA)	DT-3010	Digitizes neural signals for computer sampling.
Digital I/O Card	Measurement Computing (Norton, MA)	PCI CTR-05	Acquires behavioral inputs and outputs