The DREAM Implant: A Lightweight, Modular, and Cost-Effective Implant System for Chronic Electrophysiology in Head-Fixed and Freely Behaving Mice

All experimental procedures were conducted according to the institutional guidelines of the Max Planck Society and approved by the local government's ethical committee (Beratende Ethikkommission nach §15 Tierschutzgesetz, Regierungspräsidium Hessen, Project approval code: F149-2000).

Figure 1: Implant design. (A) 3D rendering of the implant superimposed onto a mouse skull with a silicon probe connected to a probe connector. The central aperture of the headplate is approximately 10 mm for scale. The height of the drive is approximately 17 mm. The copper mesh that forms the outside of the Faraday crown, as well as ground/ref wires, is not shown. (B) Same as (A) with a connection to an amplifier board instead of a probe connector. (C) Exploded technical drawing of the implant, showing its components. (D) Rendering of an angled spacer that can be implanted underneath a microdrive, allowing to consistently implant the microdrive at a predefined angle (here: 20°). (E) Rendering of integrated head-fixation mechanism, showing implanted headplate with Faraday crown with the surrounding head-fixation clamp and the dove-tail connection to setup. (F) Image of mouse head-fixed on a treadmill using the implant's integrated head fixation mechanism. Please click here to view a larger version of this figure.

NOTE: Sections 1 and 2 discuss the pre-surgical preparations

1. Preparation of the silicon probe

1. In case of probe reuse, clean the silicon probe according to the recommendations of the probe supplier. Soak the probe in enzymatic cleaner (see Table of Materials) for 5-10 min, then rinse it in demineralized water. Do this as quickly as possible after explantation. A day before (re-)implantation, soak the probe in 70% ethanol for at least 30 min for disinfection.
2. Measure channel impedances to make sure they are within specifications for the recorded signal. Follow the protocol for testing noise levels from the Neuropixels user manual²², measure impedance via the desired recording software (e.g., https://open-ephys.github.io/gui-docs/User-Manual/Plugins/Acquisition-Board.html#impedance-testing)  and follow the target channel impedances from the silicon probe manufacturer or datasheet. If impedances are too high, consider recoating the electrode sites²³.
3. Solder a 0.05" solder tail socket (see Table of Materials) to the ground (GND) wire of the probe. Connect the socket to the GND pin (next step) during surgery.
   NOTE: In this protocol, a separate reference (REF) pin is not used, as GND and REF are shorted on the headstage used. Therefore, only the GND pin will be mentioned in the remainder of the protocol. If a separate REF is used, repeat the following step for the REF pin.
4. To prepare the GND pin, repeatedly insert the pin side of a 0.05" solder tail socket (see Table of Materials) into the GND 0.05" solder tail socket until insertion is largely effortless. Using gold-plated pins can reduce the need for this smoothing step. This makes sure that the GND pin and socket can easily be connected during surgery without the need to apply excessive pressure, reducing the risk of injuries to the animal and probe damage.
5. If an implantable pre-amplifier for the silicon probe is used, prepare them for chronic implantation following the supplier's procedures. Then attach the amplifier/connector to the ring of the Faraday cage by using silicone plaster to glue it to the area of the Faraday ring designed to hold the amplifier (see Figure 1).
   NOTE: Preparing the implantable pre-amplifier for the silicon probe for chronic implantation following the supplier's procedures might include coating them in silicon or epoxy to avoid moisture damaging the electronics, as well as repeatedly mating the amplifier connector to reduce mating force when connecting the amplifier to the recording system during recordings. This is especially useful for Omnetics users.

2. Preparation of the microdrive and headgear

1. Turn the screw on the microdrive body so that the microdrive shuttle is almost entirely retracted upwards.
2. Optionally, attach an angled spacer (see Figure 1D) to the bottom of the microdrive with cyanoacrylate glue or dental cement, which can be used to allow for a specific degree of tilt to be used, for example, when recording through cortical layers in a region within the central sulcus, or within deep structures that may require a non-perpendicular approach (for angled spacer, see Table of Materials).
3. Lay the microdrive horizontally onto the microdrive holder (Supplementary Figure 1).
4. Place a small piece of adhesive putty (see Table of Materials) on the microdrive holder at a distance above the microdrive at which the head-stage connector will be placed. This distance depends on the length of the flex cable that connects the silicon probe to the headstage connector.
5. Place a tiny drop of silicone plaster (see Table of Materials) onto the shuttle.
6. Take the silicon probe out of its packaging with the help of a blunt, soft-tipped forceps. Make these by coating standard needle-nose forceps with 3 mm diameter heat-shrink tubing (see Table of Materials). Place the probe with the flex cable first onto the shuttle of the microdrive so that the bottom edge of the flex cable hangs slightly over the bottom edge of the microdrive shuttle.
7. Gently pull the flex cable towards the top of the microdrive until the bottom edge of the flex cable meets the bottom edge of the microdrive shuttle. Make sure to push the flex cable against the left edge of the microdrive shuttle during this step so that it is placed exactly vertically on the microdrive at the end. At this point, ensure that the shanks of the silicone probe do not (or only minimally) protrude past the lower edge of the microdrive (depending on the exact length of the probe shanks and the depth of the targeted brain area).
8. Place the head-stage connector of the probe onto the adhesive putty at the top of the holder to protect the probe from falling off.
9. Use a 27 G syringe needle to apply a small drop of cyanoacrylate glue (see Table of Materials) between the flex cable and shuttle to secure the probe in place. Ensure the glue does not run onto the microdrive or along the flex cable beyond the shuttle (This is very important)
10. Once the flex cable is glued in position, attach the amplifier to the crown ring (see Table of Materials) using silicone plaster. Then, attach the flex cable to the amplifier and cover the connection and cable in a thin layer of silicone plaster.
11. After 5 min, when the plaster is set, store the microdrive and probe safely until further use.
12. Cut pieces of copper mesh (see Table of Materials) into an open donut shape (see cutting pattern in Supplementary Figure 2) to cover the Faraday cage.
13. Fasten the copper mesh cut-out onto the Faraday cage with small drops of epoxy resin (see Table of Materials). For this step, one can also replace epoxy with dental cement.
    NOTE: The Faraday cage contains a space to house a probe connector or amplifier. This space is marked by an X in the design file, and it contains a supporting base for the amplifier/connector, as well as a larger distance between the two adjacent spokes of the cage. To create sufficient space around the amplifier/connector, fix a small amount of extra mesh between the two adjacent spokes, creating a protrusion. This ensures that the amplifier/connector can later be positioned in this 'pocket' without touching the Faraday cage. To ensure secure adhesion with minimal warping, use the crown ring placed directly on the crown to maintain shape and to support the thin spokes of the crown. Furthermore, use soldering helping hands to secure the crown and mesh during drying. If one struggles to maintain the shape of the crown when undergoing the procedure, attempt to epoxy only two of the crown arms at a time to prevent warping.
14. If separate grounding of the Faraday cage is desired, solder a small header pin onto a 30 mm grounding wire (see Table of Materials), then use conductive epoxy to adhere the wire to the copper mesh cut-out.
    NOTE: This step is not adhered to in the lab.
15. At this point, store the prepared parts safely, and perform surgery at a later stage.
    NOTE: Sections 3-6 discuss the implantation of the microdrive and headgear.

3. Surgery: Preparation of probe and workspace

1. Sterilize and place surgical instruments in the surgical workspace following an approved procedure.
   NOTE: This can include using a bead sterilizer, autoclaving instruments, or rinsing with 30% peroxide or 90% ethanol, depending on the approved experimental protocol.
2. Place the ceramic dish used to prepare the dental cement in an ice box, fridge, or freezer, following the instructions in the dental cement kit (see Table of Materials). Use the cooled ceramic dish during cement mixing to increase the time the cement is malleable. Use a cooled dish whenever longer cementing steps are required.
3. If histological verification of probe placement at the end of the experiment is desired, extend the silicon probe right before the surgery by turning the screw on the microdrive counterclockwise and apply a lipophilic dye (see Table of Materials) to the probe by dipping it in a small drop of the dye. Prepare the lipophilic dye from a commercially bought dimethyl sulfoxide (DMSO) or ethanol (EtOH) diluted stock solution (see Table of Materials) by diluting it in a suitable buffer such as PBS at a 1-5 µM concentration.

4. Surgery: Preparation of the animal

1. Follow an approved anesthesia protocol for a 2-4 h rodent surgery under aseptic conditions. This can include general and local anesthesia, analgesia, application of eye ointments, and injections of saline. Here, use injectable anesthesia (ketamine 100 [mg/kg]/medetomidine 0.5 [mg/kg]) together with local analgesia cream and eye ointment (see Table of Materials), and place the animal on a heating pad to regulate body temperature.
2. When the animal is fully anesthetized, move it to a separate non-sterile shaving area.
   1. Ensure that the animal is warmed sufficiently; for example, place it on a heating pad. Remove hair on the top of the skull. Do this with an electric shaver or depilation cream (see Table of Materials) or by repeatedly shaving the top of the head with a scalpel covered in 70% ethanol.
   2. Carefully remove loose hairs to make sure they do not get in contact with exposed tissue later. To remove hairs, use, e.g., tissues wetted with 70% ethanol and/or a squeeze ball pump. If using depilation cream, ensure that this is removed using cotton swabs and saline thoroughly.
3. Disinfect the shaved area multiple times with an iodine-based disinfectant (see Table of Materials) and alcohol using cotton swabs, moving from the center of the head to the sides to brush any remaining loose hairs away from the incision site.
4. Disinfect the fur on and around the head using betadine. This ensures a sterile working area and protects surgical instruments and materials from coming into contact with unsterile fur.
5. Place the animal in a stereotactic frame using ear bars and nose holder (see Table of Materials).
6. Using small surgical scissors (see Table of Materials), cut an almond-shaped opening in the skin on top of the skull, reaching from just the posterior of the lambda suture to between the eyes.
7. Remove the subcutaneous membrane and periosteum by cutting away while still wet, then scratch the skull with a scalpel blade to remove soft membrane tissue on the skull's surface that may impede dental cement's adhesion.
8. Optional: Once the skull has been cleared of membrane tissue, briefly apply a thin layer of 0.5% peroxide and wash it off with water-based iodine disinfectant (e.g., Betadine) before roughening the surface of the skull to improve adhesion of the primer to the skull.
9. Carefully roughen the surface of the skull by scratching a crisscross pattern with the tip of the scalpel turned upside down. This helps dental cement to adhere to the skull later.
   NOTE: Do not scratch too vigorously on top of sutures since this can cause the sutures to rupture and leak intracranial fluid, which impairs adhesion of the dental cement.
10. Alternate between scalpel blade and sterile cotton buds to gently scratch/push away neck muscles attached to the sides of the lambda suture until the muscles have been pushed back to the 'edge' of the skull on top of the cerebellum. This helps to minimize muscle noise in neuronal recordings.
11. Fill a 1 mL syringe with a 27 G needle (see Table of Materials) with small amounts of surgical cyanoacrylate glue (see Table of Materials). Then, glue the skin to the skull edges using the syringe to smear tiny drops of superglue across it. Glue tissue as flat as possible to the skull to leave space for implants. This procedure ensures that skin and muscles do not come in direct contact with parts of the implant, which avoids muscle noise in recordings and improves the adhesion of the dental cement.
12. Apply dental cement primer across the skull for extra adhesion and harden with UV light (see Table of Materials). This improves dental cement adhesion and prevents cranial sutures from leaking and weakening the cranial-cement bond over time.
13. Find the target location for the probe implantation relative to bregma or lambda and outline the craniotomy around it with a surgical marker. Place the headplate on the skull so that the craniotomy lies within it, with space for the microdrive at one side of the craniotomy, as well as for 1-2 grounding pins.
14. Implant the headplate using dental cement. Mix dental cement in the designated cooled ceramic dish (see step 3.2). Ensure the headplate adheres to the skull on all sides, forming a watertight 'well'.
15. With a dental drill (size US ½ HP), drill a small burr hole the width of the header pins prepared in step 1.4 over the brain area(s) to be used as GND/REF. If grounding the Faraday cage is desired, drill another small burr hole close to the edge of the Faraday cage for the Faraday-GND header pin.
    NOTE: For the GND/REF header pin(s), place the craniotomy at a sufficient distance from the edge of the cage so that the header pin itself can be placed within it later without touching the Faraday cage.
16. Clean the craniotomy by gently dripping sterile saline onto it with a syringe and removing it with non-shedding wipes (see Table of Materials). Repeat until all blood and loose tissue is removed.
17. Prepare a 0.7% agar (see Table of Materials) solution in saline, cool it slightly, and introduce it into the craniotomy using a 27 G needle on a 1 mL syringe.
18. Gently insert a GND pin (see step 1.3) into each craniotomy drilled in the previous step. The pin(s) will be surrounded by agar on all sides (see step 4.17). Apply cement around the header pins to secure them and provide electrical isolation.
19. Clean the ceramic dish and place it back in the fridge/freezer.
20. With a dental drill, drill the outline of a larger craniotomy (circular or square) by moving around the edge in steady movements. Ensure that the craniotomy is 1 mm x 1 mm to 2 mm x 2 mm to allow for small adjustments to the placement of the probe to avoid blood vessels without exposing too much of the cortex. If possible, avoid placing craniotomies over sutures. Drill in rounds of 20-30 s, and cool down the skull with saline between drilling rounds.
    NOTE: When beginning drilling, it is useful to mark out the leading edge of the microdrive with a marker, hence ensuring that when drilling, a straight edge can be formed in parallel to the microdrive leading edge. This improves the chances of avoiding cement in the craniotomy when fixing the microdrive in place, as well as improving adhesion, preventing microdrive overhang over the craniotomy and allowing for greater lateral maneuverability when placing the microdrive in relation to the final recording site position.
21. After a few initial rounds of drilling, test the resistance of the drilled-out portion of the bone by gently pushing on it with fine forceps (size 5 or finer; see Table of Materials).
    1. Keep testing between drilling rounds until the bone begins to 'bounce' underneath the forceps when pushed. When this is the case, add a drop of saline on top of the craniotomy to soften up the bone, then use the forceps to gently remove the drilled-out piece of bone.
    2. If the bone cannot be removed gently, do another round of drilling, focusing on the points where the bone is still attached more strongly. In general, aim to remove the skull with gentle pressure from the forceps before it has been entirely drilled through since this typically minimizes tissue damage.
       NOTE: Ensure the surface of the dura is moistened regularly, both during drilling to reduce temperatures and following bone flap removal. This improves the chances of easy probe insertion by preventing the dura from drying out and becoming more challenging to penetrate. If the dura proves to be too tough to penetrate, or blunt or multi-shank probes are being used, a durotomy is performed by lifting the dura with a 27 G needle and performing a small incision under saline immersion to prevent the dura from sticking to the brain surface.
22. Cover the craniotomy with a hemostatic sponge (see Table of Materials) soaked in cool, sterile saline to protect the dura and brain.

5. Surgery: Probe implantation

1. Attach the custom microdrive holder (see Table of Materials) to the arm of the stereotactic apparatus. If the microdrive was removed from the microdrive holder after probe preparation, place the microdrive with the attached silicon probe into the microdrive holder.Angle the stereotax arm as required to reach the desired target brain area. Place the crown ring with the attached amplifier onto the three vertical pins at the rear of the microdrive holder (see Supplementary Figure 1).
2. Lower the microdrive to within ~0.5 mm of the craniotomy, then use forceps to connect the GND/REF header pin(s) attached to the probe to the corresponding GND/REF pin(s) implanted on the skull (see steps 4.14-4.15). See Supplementary Figure 3 and Supplementary Figure 4 for examples of drive, craniotomy, and GND/REF pin placement.
3. Once in place, optionally secure the pin(s) with a drop of conductive silver epoxy (see Table of Materials) for a more robust connection. Once silver epoxy is cured, cover the connected pins in a small amount of dental cement (see Table of Materials) to ensure the connection stays stable over long periods and that there is no electrical connection with the surrounding tissues and/or implant elements.
4. Remove the hemostatic sponge from the craniotomy (see step 4.22).
5. Position the stereotactic arm with the microdrive over the craniotomy.
   NOTE: If the probe is retracted, make sure that the microdrive is placed in a way that the probe would touch down on a part of the craniotomy that does not contain large blood vessels.
6. Lower the microdrive, if necessary, by adjusting the location and angle until the probe shank touches the dura or brain surface (see step 4.21) in the target area.
7. Mix dental cement in the designated ceramic dish (see step 3.2), and cement the base of the microdrive in place, focusing on the three sides of the microdrive base that are not facing the electrode. Ensure the cement does not touch the microdrive above the removable 'base' (see Figure 1D).
   1. Make sure that any space between the base and skull is covered fully with dental cement. Clean the ceramic dish and put it back in the fridge/freezer. Wait for the cement to cure, approximately 10-15 min.
      NOTE: A small gap is left between the microdrive base and skull, and the cement is used in its most fluid form to fill it. Once the cement has thickened slightly, the cement between the walls of the microdrive base and the skull is built up. Very small amounts of cement are always used, as the flow of the substance can be unpredictable, and larger volumes may flow into undesired regions. Small amounts of hemostatic sponge dipped in saline can be used to cover portions of the craniotomy. If cement should accidentally flow onto the craniotomy, remove the cement with forceps once it enters a film-like consistency.
8. Lower the silicon probe onto the brain, carefully monitoring the probe position through a microscope. When the probe shanks touch the brain, lower the probe quickly by ~250 µm (one full turn of the screw is 282 µm) to ensure that the probe breaks through the resistance of the dura/cortical surface.
   1. Verify this visually. If the probe has not broken into the cortex, wait for 5 min, then attempt to etch through the dura with the shank tip by repeatedly raising and lowering the probe by a few tens of micrometers while the dura/cortex is under tension from the probe tip.
9. Once the probe has broken through the surface of the cortex, gradually lower it at a slower pace (100-200 µm /min) until either the target coordinates are reached or the probe has moved by more than 1000 µm. If the target requires the probe to move by more than 1000 µm, advance the probe in steps of maximally 1000 µm/session over the following recording sessions until the target coordinates are reached.
   NOTE: Skip this step if monitoring neuronal signals while lowering the silicon probe is preferred. Steps for this are described in section 7.
10. Prepare silicone elastomer according to instructions (see Table of Materials) and dispense a small drop into the craniotomy using a 1 mL syringe (see Table of Materials).
11. Once dry, cover the silicone elastomer with a 50/50 mix of bone wax and mineral oil. This step further protects the probe and prevents the accumulation of debris and dry plasma over the craniotomy, making extraction simpler and safer. Exercise caution, as working around the probe while it is lowered can lead to breakage.

6. Surgery: Implantation of Faraday cage

1. When the dental cement has fully solidified, loosen the microdrive holder by loosening the lateral screw fixating the drive with an Allen key (see Supplementary Figure 1). Gently retract the holder by ~1 cm so that the microdrive is free-standing, but the probe amplifier/connector remains fixed to the implant holder without stretching the flex cable.
2. Place the pre-made crown and Faraday mesh around the headplate by stretching the cage at the opening and slotting it over the microdrive and Flex-cable horizontally, then fix it onto the headplate with dental cement.
   NOTE: Make sure to close all spaces between the Faraday cage and skull with dental cement to protect the implant from contamination.
3. Put the Faraday crown ring (see Table of Materials) with probe connector/headstage over the crown, aligning the integrated holder for the probe amplifier/connector with the area marked by an indented 'X' on the Faraday crown (see step 2.13).
4. Secure the ring to the Faraday cage with a small drop of cyanoacrylate glue or dental cement at each spoke-ring junction.
5. Once the Faraday ring with integrated probe amplifier/connector is secured in place, fully retract the stereotactic arm with the microdrive holder. See Supplementary Figure 3 for a step-by-step guide on the assembly of these components.

7. Post-surgery test recording

1. Connect the probe amplifier/connector to the recording hardware and start a recording.
2. If the probe has not yet reached its target location during the initial insertion (see step 5.9), slowly turn the microdrive screw counterclockwise to lower the probe while monitoring neuronal signals.
   NOTE: Signals should change a) when electrodes touch the layer of silicone elastomer above the craniotomy, and b) when the electrodes begin to move into the brain (see step 7.2). High-frequency neuronal activity will be registered by electrodes that are fully inserted in the brain, while electrodes that are in contact with the CSF on the brain surface will typically show a low-pass-filtered neuronal population signal without spiking activity (akin to an EEG trace), and recording sites in air will register increased electrical noise. It is possible to additionally verify the probe insertion depth by measuring the impedance of individual channels after the test recording. Channels in contact with air should show high impedance (indicating an open circuit) and impedances like the ones measured before surgery for the channels touching CSF or already in the brain. Advance the silicon probe by a maximum total distance of approximately 1000 µm per session, with a maximum speed of approximately 75 µm/min (see step 5.5).
3. When neural local field potentials are visible across the probe and/or the probe is advanced by a maximum of 1000 µm, end the test recording and disconnect the head-stage connector.

8. Recovery

1. Cover the Faraday cage with self-adherent veterinary wrap (see Table of Materials).
2. End anesthesia and let the animal recover for a few days following approved experimental guidelines.
3. If the electrodes on the silicon probe are not yet at the desired target location, turn the screw of the microdrive in small steps with a maximum of four full turns (or ~1000 µm) per session. If necessary, repeat this procedure over several days until the target is reached. Combining the probe movement with simultaneous recordings to evaluate electrophysiological activity in areas transversed is recommended.

9. Behavioral experiments and chronic recordings

1. For chronic head-fixed recordings during task performance, attach the headplate at the base of the Faraday cage to the head-fixation clamp by manually opening the clamp and clamping the implanted headplate (see Figure 1C, E, F).
   NOTE: If head fixation is not needed, this implant system can also be used for freely moving recordings. For freely moving recordings, skip steps 9.1 and 9.7.
2. Remove the self-adherent veterinary wrap from the implant.
   NOTE: To minimize discomfort for the animal, it is suggested that a simple, rewarding behavioral task be started before this step as a distraction while the experimenter works with the implant.
3. Attach amplifier/connector to recording equipment.
4. Conduct neuronal recordings as the animal performs the task.
   NOTE: If the goal is to maximize the number of extracellular units recorded, move the shuttle by a few tens of micrometers whenever the neural yield in a location decreases. Note that after moving the probe, the signal can take minutes to hours to stabilize. Therefore, it might be beneficial to move the probe at the end of a session so that the signal can recover until the start of the next session.
5. Disconnect the recording equipment and cover the implant in a new veterinary wrap at the end of the behavioral recording.
6. Open the head-fixation clamp to detach the animal from the head fixation.

10. Probe recovery

1. At the end of the final recording, retract the silicon probe as far as possible onto the microdrive by turning the screw clockwise. Do this while the animal is head-fixed and behaving or with the animal anesthetized in the surgical setup. Chart the exit of the probe from the brain by monitoring neuronal signals simultaneously and checking for the signature of electrodes being immersed in the brain, touching the brain surface, or in contact with air (see step 7.3).
   NOTE: Depending on the histology protocol and probe, electrolytic lesions are performed before retracting the probe to determine the exact location of some electrodes on the probe. If monitoring the probe exit via neuronal recording is not necessary, retracting the probe once the animal has been terminated is also possible.
2. Terminate the animal following approved guidelines (this includes perfusing the animal if fixating the brain for subsequent histology is planned).
3. Wait for ~ 10 min after the animal has died. Then, head-fix the animal in the stereotax, making sure that the animal's head cannot move during explant to prevent probe breakage.
4. Apply a drop of saline on top of the craniotomy and let it soak for a few minutes to soften dried biological tissue on the probe shank and lower the chance of shank breakage.
5. Place the stereotactic holder approximately 0.5 cm above the microdrive. Then cut the upper end of the spokes of the Faraday cage with small surgical scissors (see Table of Materials) to free up the Faraday ring holding the amplifier/connector and transfer the ring back onto the vertical pins at the top of the stereotactic holder (see step 5.1 and Supplementary Figure 1).
6. Carefully cut away the copper mesh with the same surgical scissors by cutting out u-shaped areas of mesh between the spokes of the Faraday crown. Then, cut off the crown's plastic spokes at the base.
   NOTE: Avoid bending the printed plastic spokes as they are being cut, as they may snap and send plastic debris flying toward the probe.
7. Lower the stereotactic holder until the microdrive can be fixated in the holder using the holder's lateral screw, fixate the microdrive, then loosen the T1 screw that connects the microdrive body to the microdrive base.
8. Slowly retract the stereotactic arm with the implant holder to lift the microdrive off its base. Ensure that the microdrive separates from the base at a perpendicular angle (i.e., 'vertically' from the base).
   NOTE: If the microdrive body and base do not separate easily, verify that the movement of the stereotactic arm is not at an angle compared to the microdrive orientation. If necessary, the holder and microdrive are re-aligned to each other by slightly loosening the fixation of the animal's head and repositioning it accordingly. Correct alignment is one of the crucial aspects for easy recovery of the microdrive. Also, check whether there is any residual dental cement connecting the microdrive and microdrive base (see step 5.5). If so, the cement is carefully scraped off with a scalpel and/or dental drill depending on the amount of cement used.
9. Raise the stereotactic arm with the attached probe to create sufficient space below it.
10. Remove the animal from the stereotax, and prepare the brain by following an approved histology protocol if desired. Recover the implanted microdrive base and clean it by soaking it in acetone for several hours for later reuse.
11. Place a clean microdrive base on adhesive putty (see Table of Materials), then lower the microdrive onto the base and tighten the screw. To prevent breakage, monitor the probe position under a microscope throughout the process. This step can be completed at a later time if the implanted microdrive base needs to be cleaned for reuse first.
    NOTE: This protocol calls for the use of adhesive putty as a platform for the base, which is vital as it both secures the base whilst also having a degree of give, ensuring the base does not slip and collide with the probe. The putty should be shaped into a vertical' cliff face' on the side of the microdrive base where the probe will be lowered. This ensures that if the probe is lowered past the base, it does not make contact with the putty underneath. The putty 'tower' should also be tall enough that if it is lowered past the microdrive base, the probe does not make contact with the table surface on which the putty is placed. Finally, secure the putty well to the surface to prevent it from slipping or falling. When lowering the microdrive onto the microdrive base held by the putty, ensure a side profile view from the microscope to monitor the progress so that as the probe is lowered, it does not collide with either the base or the putty.
12. Clean and sterilize the probe following the manufacturer's instructions. For the most commonly available probes, soak them in an enzymatic cleaner (see Table of Materials) for 12 h, then rinse in demineralized water and sanitize in alcohol. Do this by lowering the probe into a large beaker containing the enzymatic cleaner while still attached to the microdrive holder on the stereotactic arm.
    NOTE: If desired, measure the impedances of the electrodes on the probe after cleaning to monitor the potential degradation of individual electrodes.
13. Store the microdrive with the cleaned probe safely until the next experiment.