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Metal microdrive and head cap system for silicon probe recovery in freely moving rodent

  1. Mihály Vöröslakos
  2. Peter C Petersen
  3. Balázs Vöröslakos
  4. György Buzsáki  Is a corresponding author
  1. Neuroscience Institute, New York University, United States
  2. Budapest University of Technology and Economics, Faculty of Mechanical Engineering, Hungary
  3. Department of Neurology, Langone Medical Center, New York University, United States
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Cite this article as: eLife 2021;10:e65859 doi: 10.7554/eLife.65859

Abstract

High-yield electrophysiological extracellular recording in freely moving rodents provides a unique window into the temporal dynamics of neural circuits. Recording from unrestrained animals is critical to investigate brain activity during natural behaviors. The use and implantation of high-channel-count silicon probes represent the largest cost and experimental complexity associated with such recordings making a recoverable and reusable system desirable. To address this, we have designed and tested a novel 3D printed head-gear system for freely moving mice and rats. The system consists of a recoverable microdrive printed in stainless steel and a plastic head cap system, allowing researchers to reuse the silicon probes with ease, decreasing the effective cost, and the experimental effort and complexity. The cap designs are modular and provide structural protection and electrical shielding to the implanted hardware and electronics. We provide detailed procedural instructions allowing researchers to adapt and flexibly modify the head-gear system.

Introduction

Action potentials are the common currency of communication between neurons and they can be detected as voltage fluctuation in the extracellular space (Adrian and Moruzzi, 1939). However, recording from representative ensembles of neurons simultaneously requires electrodes with multiple recording sites. Multi-wire twisted electrodes (tetrodes) and silicon probes offer the possibility to record tens to hundreds of neurons simultaneously from multiple cortical and subcortical structures simultaneously in freely moving animals (McNaughton et al., 1983; Wise and Najafi, 1991; Csicsvari et al., 2003; Buzsáki, 2004; Blanche et al., 2005; Montgomery et al., 2008; Jun et al., 2017). For cost benefits, microwire arrays are a popular choice for neuroscientists, despite the amount of manual labor involved (Edell et al., 1992). Silicon probes, while more expensive, do not require assembly, the tissue-volume displacement is minimal (Buzsáki, 2004; Kipke et al., 2008), recording properties are consistent (site impedance and locations) and geometric configurations (number of shanks, distance, and pattern of recording sites) can be customized to suit the architecture of the particular brain structure under study (Wise and Najafi, 1991; Scholvin, 2016). The affordability of high-channel-count electrophysiology amplifier chips (e.g., RHD-2132 and RHD-2164, Intan Technologies, Los Angeles, CA; Berényi et al., 2014) and integrated designs (Jun et al., 2017) have accelerated the spread of large-scale recordings. Integration of µLEDs into silicon-based electrodes and can offer unique spatiotemporal control of neuronal activity (Wu et al., 2015; Kim et al., 2020).

The most expensive component of the head gear is the recording probe (from $1000 for 32-channel passive recording probes to > $3000 for µLED probes or larger channel count probes). Therefore, reusing silicon probes in freely moving animals is an important current goal (Juavinett et al., 2019). In addition to reducing costs, repeated usage of the same probe/headgear would allow for better consistency in recordings across animals, enhances data reproducibility, and would reduce electrode/headgear preparation for surgery. Achieving this goal requires an integrated design of a reusable microdrive and head gear to increase recording stability and protect/shield sensitive drive and electronic components (Chung et al., 2017; Senzai et al., 2019).

Currently available headgear systems are typically large, reducing the ability to record from multiple brain structures in mice. In contrast to recent progress in recording electrodes, the development of implantation techniques such as the Flexdrive, Shuttledrive, DMCdrive and the Hyperdrive (Voigts et al., 2013; Voigts et al., 2020; Kim et al., 2020; Lu et al., 2018) has lagged behind. Electrodes are either fixed in brain tissue or attached to a microdrive to allow the advancement of the electrode after implantation (Chung et al., 2017; Fee and Leonardo, 2001; Korshunov, 2006; Vandecasteele et al., 2012; Wilson and McNaughton, 1993; Yamamoto and Wilson, 2008). Microdrives and accompanying head gear protection and shielding inevitably add extra weight (weight = 0.12 g - 1 g, drives designed for mice) and volume (skull surface area = 7.68–252 mm2, drives designed for mice) to the implant (Table 1). The weight, volume and footprint of the microdrive can limit comfortable movement of small rodents and can prevent flexible multiregional recordings in mice (Headley et al., 2015). Yet, chronic recordings from freely behaving subjects are essential in many experiments, where the relationship between neuronal activity and movement, perception, learning and memory, decision making, and other forms of cognition are studied to disambiguate overt behavior and hidden variables (Juavinett et al., 2019; Jun et al., 2017; Steinmetz, 2020). An ideal microdrive should have high precision movement, mechanical stability, minimal size, low weight, and the ability for flexible customization. Commercially available microdrives are expensive and hard to customize. Disposable 3-D printed customized drives and head gear have reduced costs (Headley et al., 2015; Chung et al., 2017; Allen et al., 2020). Most importantly, recovery and reimplantation of recording probes are limited with currently available headgears.

Table 1
Summary of microdrive designs used in mice.
Study/CompanyWidth (mm)Length (mm)Height (mm)Footprint (mm2)Weight (g)Travel distance (mm)Easy recovery
Vandecasteele et al., 20124.36.41327.520.68–12no
Janelia Research Campus3.53.8913.30.85no
Janelia Research Campus2.53.8109.50.55no
Cambridge Neurotech2.549100.545no
Neuronexus12.511.58.5143.750.361no
NeuroNex MINT3.27.516240.674.8yes
Chung et al., 20176.265.26932.930.42yes
Juavinett et al., 20191814202521-yes
Voroslakos et. al.3.1515.315.50.877yes

Below, we report the design and testing of an integrated 3D printed headgear system (including microdrives and protective head cap) that is adaptable for multiple recording devices for both mice and rats. Our design reduces surgery time substantially and the small footprint of the metal Microdrive allows targeting multiple brain structures. The fast and reliable recovery of the probe and reuse of the same system in multiple animals decreases costs and experimenter effort.

Results

Recoverable metal microdrive

3D printing has taken science and industries by the storm, offering in-house design customization, fast iterative development, and cheap production using professional printers based on filament extrusion (e.g., MakerBot Industries, New York, NY) and liquid resin (e.g., Form three by Formlabs, Sommerville, MA). Yet, plastic prints have limitations mostly due to the low strength of the materials. Repeated use of plastic threads results in rapid deterioration, which can be prevented by metal-to-metal connection (Figure 1—figure supplement 1). Recently, metal printing has become affordable offering increased strength, with options for printing in aluminum, stainless steel and even titanium with similar printing resolution to plastics. Here, we have taken advantage of this and constructed a 3D printed microdrive from stainless steel (stainless steel 316L, 20 μm resolution), which offers superior strength and form factor compared to plastic prints (Young's modulus of stainless steel: ~180 GPa vs plastic: ~2 GPa). The metal printing allows reuse of the drives with minimal wear, driving the effective cost down.

The microdrive is composed of three metal parts: an arm, a body, and a base (Figure 1A and B) and has a footprint of 15.5 mm2 (width: 3.1 mm, length: 5 mm, height: 15.3 mm). The detachable base allows for easy recovery of probes. The arm/shuttle is mounted on a screw to the drive body, allowing it to move linearly along the vertical axis simply by turning the screw (270 µm/ turn). The constructed microdrive has a total travel distance of 6 mm (maximum distance between arm and bottom of the drive body), allowing one to record from multiple brain regions across days and weeks. Due to its small form factor, multiple probes can be implanted in the same animal (Figure 1C). The drives come with a stereotaxic implantation tool, printed in plastic (clear v4 resin from FormLabs), for user-friendly and reliable implantations and probe recovery (Figure 1—figure supplement 2), consisting of a stereotactic manipulator attachment and a microdrive holder (Figure 1D and E, Figure 1—video 1 and 2).

Figure 1 with 4 supplements see all
Reusable metal microdrive.

(A) The metal microdrive consists of three main parts: a drive body, a movable arm/shuttle, and a removable base. All components are 3D printed in stainless steel. Additional necessary components are a 00–90, 1/2 ‘brass screw, a 00–90 brass hex nut, a 000–120, 1/8’ stainless steel screw fixing the drive to the base, and a male header pin. (B) The assembled drive with dimensions. (C) Schematic showing three microdrives, with silicon probes attached, implanted in a rat to target hippocampus, medial and lateral entorhinal cortices. 3D printed resin head cap is shown in purple. (D) 3D printed stereotaxic attachment and drive holder together with assembly pieces: male header pin, four 00–90 brass hex nuts, three 00–90, 1/4’ and a 3/16’ stainless steel screw. (E) Stereotaxic attachment with the metal drive assembled, and a probe attached, ready for implantation (red circle highlights the temporary soldering joint for the Omnetics connector). The backend of the silicon probe is attached to the arm using cyanoacrylate glue.

The fully assembled steel microdrive weighs 0.87 g (base: 0.23 g, shuttle/arm with nut: 0.16 g, drive body with screw and metal bar: 0.49 g). This weight and dimensions are similar to other commercially available or custom-made electrode microdrives (Table 1). The design files for the microdrive can be submitted to commercial 3D printing companies (e.g., Proto Labs, Maple Plain, MN, https://www.protolabs.com/) allowing for high-quality printing and fast production. The printing costs of the three components can be as low as $170 (base: $50, body: $60, arm: $55), a highly competitive price compared to commercial microdrives.

Inclusion criteria: microdrive systems that used silicon probes in freely moving mice

Mouse cap

To reuse silicon probes in multiple experiments, both the microdrive and the head cap have to be sturdy, easy to disassemble and reassemble. The mouse cap is composed of three parts: a base, a left-side wall, and a right-side wall (Figure 2A). The cap-base is attached to the skull of the animal during anesthesia using a ring of Metabond cement, serving as a base for the rest of the cap. There is no need for skull support screws, making the head cap minimally invasive. The cap has a large internal window shaped as an elongated octagon, following the outer ridge of the skull, giving wide access for various surgical needs (Figure 2B). The sidewalls provide structural support, electrical shielding (by acting as a Faraday cage), and physical protection of the silicon probes, hardware, and electronics. The internal volume allows for great flexibility and can fit two Omnetics preamplifier-connectors, as well as optic fibers. The sidewalls attach to the base using a rail and with three support screws (Figure 2—video 1).

Figure 2 with 1 supplement see all
Mouse cap.

(A) The mouse cap consists of three main 3D printed parts: a base, and two side walls. The pieces are assembled with three 000–120, 1/8’ steel screws, six male header pins, and copper mesh. (B) The base with the left side wall attached. Copper mesh was attached in three pieces to the wall, and a male header pin was soldered across the top of the wall. (C) The fully assembled mouse cap. (D) The implanted headgear with preamplifier and recording cable attached. (E) Wide-band extracellular traces recorded from the prelimbic cortex of the implanted mouse shown in (D) using a multi-shank silicon probe during food pellet chasing exploration (sh-1 and sh-2 denote shank 1 and shank 2 of the silicon probe). (F) Well-isolated single units can be recorded in the anterior cingulate cortex using the mouse cap system and microdrive (n = 31 putative single units with a 4-shank probe; same session as in E). The location of the maximum waveform amplitude of each neuron is shown (0 μm corresponds to the location of the topmost channel of the shank). (G) The average spike waveform for each neuron. (H) Distribution of the spike amplitude across the recorded neurons (n = 31).

The entire cap weighs 2.2 g (base: 0.19 g, walls with male header pins and copper mesh: 0.98 g each, and 000–120 screws: 0.05 g; Figure 2C). A chronically implanted mouse can carry this cap with one (or more) implanted silicon probe and with a custom connector for electrical stimulation (Figure 2D). High-quality electrophysiological signals can be collected from freely moving mice for weeks (Figure 2E and F). The system can be customized as needed, using our CAD files (see Methods section). We recommend printing the cap system on the Formlabs Form 2/3 resin printer or a comparable 3D printer (requires 25–50 µm resolution).

Rat cap

The typical Long-Evans rat is approximately ten times heavier than the mouse (~400 g), and requires a sturdier cap system, capable of withstanding forceful impacts and provide increased protection of the electronics and hardware. The rat cap is composed of four parts: a base, a left-side wall, a right-side wall, and a top cover (Figure 3A). The octagon-shaped base aligns with the outer rim of the rat’s dorsal skull surface and is attached with Metabond cement, with no need for skull support screws, making it minimally invasive (Figure 3—figure supplement 1A). The two side walls are attached to the base with a single rotation-axis located in the front of the base, attached with a long screw (Figure 3B, top part). The walls are held in place on the base, using a rail and two screws in the back. The sidewalls have two sets of male header pins for soldering standard Omnetics probe connectors (see Surgical Instructions). The lid can be locked with a thumb screw and has holes for air ventilation (Figure 3B bottom part, Figure 3—video 1). High-quality electrophysiological signals can be collected from freely moving rats for weeks either using silicon probes (Figure 3C and D) or Neuropixels probes (Figure 3—figure supplement 2).

Figure 3 with 5 supplements see all
Rat cap.

(A) The rat cap consists of four main 3D printed plastic parts: a base, two side walls, and a lid. To assemble the components, an M2 nut, M2 thumb screw, a 00–80, 1’ screw, a 00–80 insert, and two 00–80, 5/32’ screws are also needed. (B) The assembled rat cap is shown with sidewalls in an open position (top image), closed configuration without (bottom left) and with the lid in place (bottom right). (C) A Long-Evans rat in its home cage with the rat cap, connected to preamplifier and cable. (D) Extracellular traces recorded on post-op day 18 from the same animal.

For more complex experiments, the cap system can be modified to increase the available skull surface (Figure 3—figure supplement 1A and B). This modified base is held by bone screws implanted in the temporal bone and covered with dental cement (Figure 3—figure supplement 1B right part). Increasing the inner volume of the cap system and using metal recoverable microdrives enable multiprobe implantations (Figure 3—figure supplement 1C).

The entire design weighs 11.03 g (base: 1.04 g, right wall with male header pins and copper tape: 3.48 g, left wall with male header pins and copper tape: 3.68 g, top with thumb screw: 2.35 g and 00–80 screws: 0.48 g; Figure 3B bottom, right).

Surgical advantages using the head cap systems

The modular system decreases the duration of the surgery and allows for faster post-operative recovery of the animal, due to four important modifications. 1. The head cap is prepared before surgery and can be reused easily. 2. The cap does not need support screws, reducing the invasiveness of the surgery and accelerating the animal’s recovery. 3. The 3D printed cap-base is secured with a single step, by attaching it to the dorsal surface of the skull with Metabond cement. This ensures alignment precision relative to the brain surface, easier probe recovery, and reusability. 4. The electric shielding and structural support is implemented in the reusable head cap, decreasing extra manual steps for the construction of the protective cap from copper mesh, male header pins and grip cement during surgery (Vandecasteele et al., 2012).

These steps offer a time savings from 40 to 90 min (Figure 3—videos 2 and 3), compared to a manually constructed cap during surgery (Vandecasteele et al., 2012).

Further, the modular cap system substantially increases flexibility during an implantation procedure. Because the sides can easily be disassembled and reassembled, complex surgical procedures can be split into multiple sessions when needed. In the first session the skull is prepared, and the base of the cap is attached to the skull. After recovery, the craniotomy and implantation are performed in a second surgery. This double-step procedure results in a speedy recovery of the animal and reduces the likelihood of human error during extended procedures. Additionally, subsequent troubleshooting can be performed through the course of long chronic experiments with minimal disruption to the animal and the implanted components.

Probe recovery

To recover the probe at the end of a chronic experiment, the drive holder is aligned with the drive using the stereotactic frame. Once the position is aligned in the x-y plane, the drive holder is moved downwards (Figure 4A, step 1). Next, the top of the drive is secured with the screw located on the side of the drive holder (Figure 4A, step 2). The 000–120 screw is removed from the base (Figure 4B, step 1) and the drive is moved upwards carefully (Figure 4B step two and C). We recommend to carefully monitor the shanks of the probe under a microscope during the entire recovery procedure and, if any unexpected movement of the probe is observed, return to the previous step to make sure that everything is secured properly (Figure 4—videos 1 and 2).

Figure 4 with 2 supplements see all
Probe recovery procedure.

(A) The stereotaxic probe holder is attached to the microdrive (step 1) and is fixed with the black screw (step 2). Precise alignment is critical to avoid tissue damage and to prevent breaking the shanks when retracting the probe. (B) The microdrive is detached from the drive-base by removing the 000–120 steel screw (step 1) and moved upwards (step 2). Camera angle rotated 90o. (C) The drive with the attached probe after retracting it from the brain. The drive-base can be reused by cleaning it in chloroform or acetone.

The removed silicon probe (NeuroNexus, Cambridge Neurotech, Diagnostic Biochips products; Neuropixels) is cleaned by initially rinsing it in distilled water, then contact lens solution (containing protease) and distilled water again; each washing step should last for at least 12 hr. Soak the silicon shanks only (avoid soaking the backend area). If extra tissue or debris is detected between the shanks, it can be carefully removed by a fine needle (26 gauge or smaller) under a microscope. After recovering Neuropixels 1.0 probes, the probe shank should be soaked in 1% tergazyme (Alconox) for 24–48 hr, then rinse in distilled water and isopropyl alcohol (Luo et al., 2020).

Quantification of single unit quality measures

Microdrives allow experimenters to record from novel sets of neurons in successive sessions, surveying thousands of neurons from the same structure in a single animal (Girardeau et al., 2017). With the recoverable metal microdrive, we recorded across several days from the same animal while adjusting the implantation depth (500 µm to 780 µm) across days (Figure 5A–F). Across days of recordings, while the probe was moved to record from different depths, the unit yield increased (Figure 5H), the waveform amplitude increased (Figure 5E) while the relative noise level decreased (Figure 5F), suggesting either that the distance between the electrode sites and neuron bodies decreased or that large size neurons were recorded.

Single unit quantification.

(A) Recordings from the prefrontal cortex at multiple depths across 12 days with a four-shank silicon probe in a mouse (128 channel probe from Diagnostic biochip; P128-5). Individual shanks are displayed as rows across days to better visualize the cells across days. The probe was moved in 70 µm steps to record from a new population of cells across days. Colored dots: position of single cells determined by spike amplitude trilateration. Grey dots: electrode sites. (B) Left: Histogram and boxplots of the distribution of recorded neurons as a function of cortical depth (µm) for each session shown in A. Each colored histogram and corresponding box-plot correspond to the same days shown in A. Right: Probe layout (shanks now shown in a horizontal layout) with the shanks shown with the depth for day 8–10; shanks are spaced by 150 µm. (C) Number of isolated single units across days after the first implantation (black) and after reimplantation of the probe (blue). (D–F) Firing rate (D), waveform amplitude (E; trough-to-peak) and relative noise level (F; waveform std divided by the waveform amplitude). (G–I) Comparison with other control mice and rats (n = 10 subjects), implanted with custom built drives (Vandecasteele et al., 2012), comparing waveform amplitude (G), number of cells/recording site (H) and relative noise level (I; same definition as in panel F). Lines refer to different animal subjects. Thick black line: rat with the metal drive; thick blue line: rat with reimplanted silicon probe mounted on metal drive (panel A-F). Days relative to the first recording session from each animal. (J–L) Neuropixels probe recording, where the same putative interneuron was tracked across four days. (J) Average waveforms (bandpass filtered 300–10000 Hz) of a putative interneuron recorded on16 channels across 4 days (left). The average waveforms recorded at the site with the largest amplitude waveform is highlighted on the right (waveforms are color-coded by recording days). Autocorrelation histograms (K) and spike amplitudes (L; from Kilosort) for the same single unit, color-coded by recording day.

Effect of head gear type on behavior

Finally, we compared the behavioral effect of the 3D printed head cap system with manually built headmounts. To this end, we compared the running speed of rats and mice between subject implanted with the 3D printed and manually built headgears. Rats and mice were water deprived and had to collect water reward on a linear track or figure-eight circular maze. We observed no significant difference between the median running speed of the two rat groups (Kolmogorov-Smirnov test (KS-test); p=0.35) or the 95 percentiles (KS-test; p=0.95). We also performed the same test on mice and found a significant difference between the median running speed of the two groups (KS-test; p=0.045G) but no significant difference between the 95 percentile speeds (KS-test; p=0.24; Figure 6H).

Effect of head gear type on running speed.

Rats and mice implanted with the 3D printed head cap system and subjects with manually built headgear. (A–B) The distribution of running speeds within individual sessions (A) and within individual subjects (B). Three rats implanted with the 3D printed head gear (13 sessions), compared to four subjects with manually built headgear (22 sessions). (C–D) Median (C) and 95 percentiles (D) of the running speed, per sessions. (E–F) The distribution of running speeds within session (E) and average across sessions per subjects (F). Three mice implanted with the 3D printed head gear (nine sessions), compared to five control subjects with manually built headgear (54 sessions). (G–H) Median (G) and 95 percentiles (H) of the running speed, per sessions.

Discussion

We have developed a recoverable microdrive printed in stainless steel and a head cap system for chronic electrophysiological recordings in freely behaving rats and mice. The cap system allows for considerably faster and more standardized surgeries to be performed and faster post-surgical recovery of the animals. Importantly, recovery of the probe and head cap becomes an easy and routine procedure, allowing the same silicon probes to be used in multiple animals, offering substantial savings.

Our head caps are minimally invasive and do not require supportive bone screws, shortening surgery time and postoperative recovery. Except for the base, the entire headgear is reusable, making experiments performed on multiple animals less variable. For multiple surgeries (e.g., virus injection for optogenetic or pharmacogenetic experiments), implantation of the base during the first surgery provides fixed coordinates for a subsequent surgery. The head cap system is flexible, due to the large internal volume, and allows for multiple probe implants, optical fiber implants, and other optional components. In contrast, manually constructed cap systems are time-consuming to build, require extensive experience, and its construction may vary from animal to animal and across investigators even in the same laboratory. The main disadvantage of existing headgears is the limited success for probe recovery. Even after successful recovery of the recording probe, a new protective cap must be built from scratch in subsequent surgeries. In contrast, our modular cap system is prepared before surgery, decreasing the time the animal spends under anesthesia, reducing potential complications during and after surgery. Using this strategy, we were able to explant and implant the same silicon probe in >10 mice (Senzai et al., 2019).

The metal microdrive weighs 0.87 g with a footprint area of 15.5 mm2, allowing the implantation of multiple probes in rats and even in mice. Because the entire headgear can be removed from the base with a screwdriver, recovery of the silicon probes is simple and highly successful. The drives are printed in stainless steel, with a stiffness prints approximately 100 times higher than that of plastic (Young’s modulus of stainless steel: ~180 GPa vs. plastic: ~2 GPa). Steel drives provide higher stability, potentially better recording quality, and prevent potential wobbling while turning the screw to adjust the probe’s position in the brain. Commercially available drives are typically built from plastic, are non-recoverable, and more expensive. Hand-made drives introduce variability across drives and experiments. In contrast, 3D steel printing provides high consistency across drives, reducing interexperimental variability.

Our system allows electrophysiologists to record the neuronal activity from multiple brain regions simultaneously in freely moving rodents. The large internal volume of the head cap and the small footprint of the metal microdrive enable researchers to perform more than one silicon probe implantation in freely moving mice and rats. Despite the highly successful recovery of silicon probes, the probe itself can deteriorate over time, limiting the number of reimplants (decreased signal-to-noise over time, reduced number of high-quality single unit clusters). Appropriate cleaning procedures can extend the lifetime of silicon probes.

To facilitate wide use of the 3D printed designs, we share all necessary details of parts, fabrication process, and vendor source for easy replication by other laboratories (see Materials and methods). We offer several video tutorials, which describe the construction of the microdrive, the cap systems, the probe implantation, and the probe recovery. The CAD system allows different laboratories to customize both the drive and headgear according to their specific goals and needs.

Materials and methods

Key resources table
Reagent type
(species) or resource
DesignationSource or referenceIdentifiersAdditional information
OtherRecoverable drive (base)‘This paper’ – Github repositoryBase_v7.step
OtherRecoverable drive (drive)‘This paper’ – Github repositorydrive_v7.step
OtherRecoverable drive (arm)‘This paper’ – Github repositoryarm_v7_50 um.step
Other00–90 nutMcMaster92736A112
Other00–90 screw 1/2’McMaster92482A235
Other00–120 screw 1/8’McMaster96710A009
OtherMale header pinDigiKeySAM1067-40-ND
OtherT1 and T2 screwdriverMcMaster52995A31
Other00–90 tapMcMaster2504A14
Other000–120 tapMcMaster2504A15
Otherstereotax_attachment_metal‘This paper’ – Github repositorystereotax_attachment_metal_v7.stl
Other00–90 nutMcMaster92736A112
Other00–90 screw 1/4’McMaster93701A005
OtherMale header pinDigiKeySAM1067-40-ND
Otherdrive_holder_metal‘This paper’ – Github repositorydrive_holder_metal_v7.stl
Other3 × 00–90 nutMcMaster92736A112
Other2 × 00–90 screw 1/4’McMaster93701A005
Other00–90 screw 3/16’McMaster93701A003
Other3D printed mouse cap (left wall)‘This paper’ – Github repositoryleft_wall_v12_L11.5
mm_W10.00mm.stl
Other3D printed mouse cap (right wall)‘This paper’ – Github repositoryright_wall_v12_L11.5
mm_W10.00mm.stl
Other3D printed mouse cap (base)‘This paper’ – Github repositorymouse_base_v12_L11.5
mm_W10.00mm.stl
Other3D printed mouse cap (cut out)‘This paper’ – Github repositorymouse_hat_copper
Mesh_cutOut_v02.stl
Other3 × 000–120 screw 1/8’McMaster96710A001
OtherMale header pinDigiKeySAM1067-40-ND
OtherCopper meshDexmet3CU6-050FA
OtherT1 screwdriverMcMaster52995A31
Other000–120 tapMcMaster2504A15
Other3D printed rat cap (left wall)‘This paper’ – Github repositoryrat_cap_left_wall_v8.stl
Other3D printed rat cap (right wall)‘This paper’ – Github repositoryrat_cap_right
_wall_v8.stl
Other3D printed rat cap (base)‘This paper’ – Github repositoryrat_cap_base_v8.stl
Other3D printed rat cap (top)‘This paper’ – Github repositoryrat_cap_top_v8.stl
Other00–80 screw 1’McMaster92196A060
Other00–80 brass insertMcMaster92395A109
Other2 × 00–80 screw 5/32’McMaster92196A053
OtherMale header pinDigiKeySAM1067-40-ND
OtherCopper tapeMcMaster76555A724
OtherM2 × 0.4 thumb screwMcMaster99607A256
OtherM2 × 0.4 thin nutMcMaster93935A305
Other00–80 tapMcMaster2523A461
Other0.05’ hex keyMcMaster5497A22
Other3D printerFormlabsForm2
OtherClear resinFormlabsRS-F2-GPCL-04
OtherCotton swabsFisher Scientific19-062-616
OtherKimwipesKimtech34120
OtherGelfoamFisher ScientificNC1861013
OtherScrewdriverAmazonB0058ECJIE
Other000–120 screw 1/16’Antrin Miniature SpecialtiesAMS120/1B-25
OtherBurrs for micro drill 0.7 mmFine Science Tools19008–07
Chemical compound, drugC and B Metabond Base 10 mlParkellP16-0116
Chemical compound, drugC and B Gold CatalystParkellP16-0052
Chemical compound, drugC and B Metabond Clear PowderParkellP16-0121
Chemical compound, drugUnifast Trad Powder IvoryPearson DentalG05-1224
Chemical compound, drugUnifast Trad LiquidPearson DentalG05-1226
Chemical compound, drugUnifast 1:2 Package A2Pearson DentalG05-0037
OtherDental LED LightAphroditeAP-016B
Chemical compound, drugCyanoacrylateLoctite45208
OtherGround/reference wireSurplus Sales(WHS) LW-12/36
OtherGround/reference wirePhoenix Wire Inc36744MHW - PTFE
Chemical compound, drugUltrazyme Enzymatic Cleaner TabletsUltrazymeB000LM0ZYS
OtherDieffenbach Vessel Clips
Straight (rats)
Harvard ApparatusST2 72–8815
OtherIntan USB Eval boardIntan Technologies LLCC3100
OtherIntan headstageIntan Technologies LLCC3324 and C3325
OtherIntan cableIntan Technologies LLCC3216

Microdrive assembly instructions

Request a detailed protocol

The base of the microdrive anchors the body of the microdrive via a tapped hole in the back (000–120 tap) and four rectangular holes inside the base (0.5 × 0.5 mm2). Thin walls around the base prevent cement flowing between the base and the body during surgery (Figure 1A). Glue a nut inside the arm (referred to as ‘arm nut’; 00–90 brass nut) before attaching it to the body. The body has an opening in the top part of the back where a nut can fit inside (‘top nut’; 00–90 brass nut). Insert the ‘top nut’ from the back, then insert the arm from the front and introduce a screw (00–90, 1/2’, brass screw) through the ‘top nut’ and the ‘arm nut’. Tighten the screw completely and release it a quarter-turn (or less). Fix the ‘top nut’ and the screw together using solder so the arm can be moved linearly relative to the body by turning this screw. Attach the body-arm complex to the base using a screw in the back (000–120, 1/8’, stainless steel screw). Finally, insert a male header pin into the body and secure it using dental acrylic cement (Unifast Trad). This can be used as a soldering joint during surgery. Finally, attach the backend of the silicon probe to the arm using cyanoacrylate glue and solder the Omnetics connector (Omnetics Connector Corporation) of the probe to the male header pin of the drive holder. The fully assembled microdrive weighs 0.87 g (base: 0.225 g, arm with nut: 0.159 g, body with screw and metal bar: 0.486 g).

  • Assembly_instructions_microdrive_metal_v7.pdf contains instructions with photographic documentation.

  • Figure 1—video 1 shows the assembly of the metal microdrive.

  • Figure 1—video 2 shows the attachment of a Neuropixels probe to metal microdrive.

Implantation/recovery tool assembly instructions

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Insert and glue one nut (00–90, brass nut) and a male header pin into the stereotactic attachment using cyanoacrylate glue. Insert a 00–90, 1/4’ stainless steel screw into the nut. Tightening this screw will secure this piece to the electrode holder of the stereotactic arm (Model 1770, Kopf Instruments). The male header pin should be used as a temporary soldering joint for the Omnetics connector of the silicon probe. Insert and glue two nuts (00–90, brass nut) into the bottom of the drive holder and one nut (00–90, brass nut) into the body of the drive holder. Insert a 00–90, 3/16’ stainless steel screw through this latter nut. This screw should be used to secure the top part of the body of the drive to the drive holder. Attach the stereotaxic attachment to the drive holder using 00–90 screws (00–90, 1/4’, T2 screw).

Assembly_instructions_implantation_tool_metal_v7.pdf contains instructions with pictures.

Mouse cap assembly instructions

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The base has a rectangular hole for a male header pin (0.8 × 0.8 mm2) for fixing the left and right walls temporarily during surgery (Figure 2B). This can help to open the cap using a fine pair of tweezers. The tip of the tweezer is squeezed between the rectangle and the walls. Pushing the tweezer against this rectangle readily opens the walls. The right wall has one tapped hole in the front and one in the lower part of the back (000–120 thread, 1.9 mm length). In addition, it has a hole in the upper part of the back (1 mm in diameter, 1.4 mm length). The left wall has one hole in the front and one in the lower side of the back (1 mm in diameter, 1.4 mm length) and a tapped hole in the upper side of the back (000–120 thread, 1.9 mm length). In addition, there are two rectangular holes in each wall (0.8 × 0.8 mm2) in which male header pins are glued with cyanoacrylate glue to serve as soldering points for the Omnetics connector and for the shielding copper mesh. To reduce weight, walls are perforated and covered with light copper mesh by gluing it with dental acrylic (Unifast Trad). The walls are closed using two screws in the back and one screw in the front (000–120, 1/8’ stainless steel pan head torx screws).

  • Assembly_instructions_mouse_hat_10_39 mm_v11.pdf file contains instructions with pictures.

  • Figure 2—video 1 shows the assembly of the mouse cap.

Rat cap assembly instructions

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The base has a hole for a brass screw-to-expand insert (00–80 thread size, 1/8’ installed length) and serves to hold together the left and right walls. It has a rectangular protrusion in the back (3 × 1.5 × 1.67 mm3) to help opening of the cap using a fine tweezer. The right and left walls have a front hole (diameter 1.8 mm) in which a screw can be passed (00–80, 1’ 18–8 stainless steel socket head screw) for fixing the walls to the metal insert of the base. In addition, there is a rail on each wall at the bottom part that grabs onto the base piece (1.2 mm height and 1 mm deep).

During surgery, the walls are kept open with the screw loosely tightened (Figure 3B, top part). After all the connectors are attached to the male header pins, the walls are closed, and the front screw is tightened. The right wall has a hole in the upper side of the back (1.8 mm, 2 mm length) and a tapped hole in the lower side of the back (00-80 thread, 2 mm length). The left wall has a hole in the lower side of the back (diameter 1.8 mm, 2 mm length) and a tapped hole in the upper side of the back (00-80 thread, 2 mm length). The walls are closed in the back using two screws (18-8 stainless steel socket head screw, diameter 0-80, 5/32” length). The left wall also has an insert in the upper part of the back side for a nut (18-8 stainless steel thin hex nut, M2.5 × 0.45 mm thread). This latter nut serves as a locking mechanism for the top cover. There are four rectangular through-holes in each wall (0.8 × 0.8 mm) in which male header pins are glued with epoxy (Araldite Standard Epoxy) and serve as soldering points. The locations of the holes can be modified according to user specifications to adapt different connector placements. To protect the implanted electrodes, the rat cap is covered by either self-adherent wrap (3M Coban) or the plastic top cover. The edge is extruded on the outer surface on top of the walls to provide extra surface for better adhesion. The plastic cover is attached to the walls using the front slide-in slot and the back screw (stainless steel flared-collar knurled-head thumb screw, M2 × 0.40 mm thread size, 4 mm long). To protect the neuronal signal from environmental electromagnetic interference noise, conductive copper coil electrical tape is glued to the walls by cyanoacrylate glue (copper tape: 1" wide, McMaster product number: 76555A724) and connected to the ground.

  • Assembly_instructions_rat_cap_v8.pdf file contains instructions with photographs.

  • Figure 3—video 1 shows the assembly of the rat cap.

3D designing and printing parts

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All parts were designed in Autodesk Fusion 360 (https://www.autodesk.com/products/fusion-360). We tested and printed cap designs on a Form two printer from Formlabs with 50 µm resolution using their standard resins. The metal microdrive prints were produced by Proto Labs (https://www.protolabs.com/services/3d-printing/direct-metal-laser-sintering). All designs are available from our GitHub repository https://github.com/buzsakilab/3d_print_designs (copy archived at swh:1:rev:a073716d89c32f13eb76a5ac5e7fa6f7fa11e18a; Vöröslakos, 2021).

3D metal print submission procedure with Proto Labs.

  1. Download the. step files from our GitHub.

  2. Create a user account at Proto Labs (https://www.protolabs.com/) and upload the files.

    1. Add the following note to the drive body:’ Use orientation as in quote 9301–742.’

    2. Choose Stainless Steel 316L material, high resolution and standard finish.

Proto Labs (Proto Labs, Inc, MN, USA) currently charges 250$ for a single print but substantial savings are available for larger print orders. Coordination of orders across laboratories therefore can reduce the price.

Subjects

Rats (adult male Long-Evans, 250–450 g, 3–6 months old, n = 11) and mice (adult male C57BL/6JxFVB mice, 32–40 g, n = 6) were kept in a vivarium on a 12 hr light/dark cycle and were housed two per cage before surgery and individually after it. All experiments were approved by the Institutional Animal Care and Use Committee at New York University Medical Center. Animals were handled daily and accommodated to the experimenter before the surgery and behavioral recording.

Surgery instructions

The following instructions cover surgeries in both rats and mice, with differences highlighted. Prior to surgery, prepare the 3D printed cap, the microdrive(s), the implantation tool and attach a silicon probe to the microdrive (as described above).

We recommend measuring the impedance of the silicon probe before implantation using the RHD USB interface board from Intan (Intan Technologies LLC, CA, USA). Lower the probe into 0.9% saline and ground the saline to the recording preamplifier ground. Connect the probe to an Intan preamplifier headstage (RHD 32- or 64-channel recording headstages) and to the main Intan board to perform the impedance test measurement at 1 kHz frequency. This measurement provides a quick and rough estimate about the quality of the recording sites. If higher precision is required, users can perform impedance measurement with a nanoZ device following the manufacturers recommendation (nanoZ Impedance Tester, Plexon Inc, TX, USA).

Prepare the stereotaxic apparatus and tools

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  1. Place the heating pad under the position of the ear bars.

  2. Sterilize surgical instruments.

  3. Weigh the animal subject.

  4. Place all components in alcohol for disinfection.

  5. Mice: prepare bupivacaine in an insulin syringe (0.4–0.8 ml/kg of a 0.25% solution).

Surgery

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Anesthesia and pre-incision preparations
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  1. The animal is anesthetized for 3 min (until after it loses its righting reflex) in an anesthesia-bucket with 2.5:1.5 (Anesthetic % to Airflow ratio).

  2. Apply a local anesthetic to the tips of the ear bars before insertion (LMX-4 Lidocaine 4% topical cream). Fix the head with ear bars and attach the closed ventilation nosepiece. Once the animal is in the stereotactic apparatus, the level of anesthesia is lowered (1.2–2%).

  3. Remove the hair above the planned surgery site using either Nair-hair remover or a hair trimmer.

  4. Clean the hairless skin with the antiseptic solution and repeat the process two more times (Povidone-Iodine – 10% topical solution). Apply the antiseptic solution with Kimtech wipes using anterior to posterior swipes. The last swipe must be done in one stroke to minimize infections. Between each swipe with the antiseptic solution, the skin is cleaned by 70% alcohol applied with the same technique.

Incision and skull cleaning
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  1. Inject bupivacaine (0.4–0.8 ml/kg of a 0.25% solution) subcutaneously along the scalp midline for local anesthesia. Make one injection site and distribute the anesthetics along the midline.

  2. Make a median incision from the level of the eyes to the back of the skull (neck).

  3. Separate the skin from the skull, pull the skin sidewise and attach four bulldog clips to create a rectangular shape opening. The bulldog clips should be attached to the subcutaneous soft tissue, not the skin.

  4. Scrape the skull with a scalpel and remove the periosteum from the top flat surface of the skull. This is necessary to achieve a strong bond with the 3D printed base.

  5. Clean the skull surface with saline and vacuum suction.

  6. Clean the skull with hydrogen peroxide and rinse it with saline. The hydrogen peroxide is applied with cotton swabs (about 5 s) and rinsed quickly thereafter thoroughly with saline. Avoid touching the skin and muscle with the solution.

  7. Cauterize any bleedings along the skull and exposed skin.

Attaching the base to the skull
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  1. Prepare the Metabond on ice. Mix four drops of base with 1 drop of catalyzer.

  2. Paint, using a brush, the whole surface of the cleaned and dried skull and let it dry.

  3. Mix a new solution of Metabond with powder: four drops of base, one drop of catalyzer and 2 scoops of powder and apply a second layer of Metabond paint to the skull surface. Paint also along the edge of the skull surface.

  4. Paint the bottom surface of the 3D printed base with Metabond and align it above the skull and attach it to the skull before it solidifies.

  5. Paint with Metabond along the inner contact line between the hat base and the skull and create a sealed area inside the hat.

  6. Gently hold the hat base in place (for about for 60 s) until it stays attached to the skull using your fingers. Let the Metabond cure before proceeding to the next steps.

Craniotomy marking and screw placement
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  1. Align Bregma and Lambda in the same horizontal plane. Determine the position of Bregma using stereotactic coordinates with a fine needle attached to the stereotactic arm.

  2. Calculate the relative positions of the probe incision points.

  3. Mark the positions of the planned craniotomies with a scalpel (gently make two orthogonal lines crossing at the planned incision points with the scalpel) and a pen (fill the scalpel-drawn lines with the pen).

  4. Mark the position of the reference and ground screws with the scalpel/pen.

  5. Remove the stereotactic arm.

  6. Drill holes for ground and reference screws in the skull above the cerebellum with a high-speed drill. If bleeding occurs, rinse it with saline and vacuum suction until the bleeding stops.

  7. Insert the ground and reference screws in. Begin with a slight counterclockwise turn. For mice, allow a margin of about 0.5 mm. In rats, drive the screws tight. Alternatively, 125 µm stainless steel wires can be used for reference and ground, instead of screws.

Craniotomy
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  1. Perform the craniotomy with a high-speed drill (drill bit size depends on the goal). Rinse it with saline and vacuum suction to ensure visibility while drilling.

  2. Clean around the craniotomy with the drill or a scraping/sharp scooping tool.

  3. Remove the dura with a hook-shaped needle at the planned incision site for probe insertion: bend the tip of the 30G needle to form a small hook (gently tap the tip of the needle into a hard surface to form the hook). Lift the dura with the hook and cut with a pointed scalpel (size 11). Avoid damaging blood vessels.

  4. Apply saline and Gelfoam to the craniotomy to maintain a wet brain surface.

Probe implantation
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  1. Place the silicon probe in the implantation tool on the stereotactic arm and position it according to the specified surface coordinates.

  2. Lower the silicon probe to the brain surface at the marked coordinates.

  3. Insert the probe to the desired target depth in the brain.

  4. Fix the base of the microdrive to the skull and hat-base with regular grip cement.

  5. Apply silicone to the craniotomy, let the silicone run along the shanks and seal the craniotomy completely. This protects the brain and limits bleedings and blood coagulation. Alternatively, apply a mixture of paraffin oil/wax to the craniotomy with a needle and heat it using the tip of a soldering iron.

  6. Solder the reference and ground wires to the corresponding sites on the Omnetics connector.

  7. Attach the cap sidewalls to the base.

  8. Cover the top with the lid or Coban tape.

  9. Turn off the anesthesia and release the animal from the stereotactic setup.

Post-operative care
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  1. Weigh the animal after surgery to determine the weight of the added headgear.

  2. Place the animal back in a home cage. The cage should be placed on a heating pad during the first night.

  3. Inject Buprenex subcutaneously after 20 min (0.05–0.1 mg/kg).

General notes
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  • Apply mineral oil to the eyes of the animal at regular intervals.

  • To keep the animal properly hydrated during the postoperative days, provide an aqua-gel and a small container with water. Provide regular rodent pills.

Additional implantation information

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Rats and mice were implanted with silicon probes to record local field potential and spikes from the CA1 pyramidal layer in rats and from the prelimbic cortex from mice. Silicon probes (NeuroNexus, Ann-Arbor, MI and Cambridge Neurotech, Cambridge, UK) were implanted in the dorsal hippocampus (rats: antero-posterior (AP) −3.5 mm from Bregma and 2.5 mm from the midline along the medial-lateral axis (ML); mice: antero-posterior (AP) +1.75 mm from Bregma and 0.75 mm from the midline, 10 degree relative to the sagittal axis). The probes were mounted on the recoverable metal microdrive and previous design iterations made in plastic (unpublished work; stl files and instructions are available at our GitHub repository https://github.com/buzsakilab/3d_print_designs/tree/master/Microdrives/Plastic_recoverable), allowing for precise vertical movement after implantation and implanted by attaching the base of the micro-drives to the skull with dental cement (Supplementary file 1). The small footprint of the metal microdrive enables researchers to perform more than one silicon probe implantation in freely moving mice. For this purpose, larger mice (>35 g) were selected.

After the post-surgical recovery, we moved the probes gradually in 50 µm to 150 µm steps until the tips reached the pyramidal layer of the CA1 region of the hippocampus. The pyramidal layer of the CA1 region was identified by physiological markers: increased unit activity and the presence of ripple oscillations (Mizuseki et al., 2011). In mice, the probe was implanted 500 µm below the surface of the brain and recordings were performed each day. The probe was moved 70 µm after each recording day. Data was collected daily. The implanted animals were single housed, and they do not carry the headstage while in the vivarium. During recordings, the headstage is attached and a counterbalanced pulley system ensures that the animal is not carrying the extra weight of the headstage and cable.

Electrophysiology data

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Electrophysiological recordings were amplified using an Intan recording system: RHD2000 interface board with Intan 32 and 64 channel preamplifiers sampled at 20 kHz (Intan Technologies). All data is available from https://buzsakilab.com/wp/database/ (Petersen et al., 2020a).

Behavioral data

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Rats were trained to run on a 2.3 m linear track, or a 120 cm diameter circular track with a diagonal path allowing the animals to run in a figure-eight pattern (n = 3 rats implanted with the 3D printed head gear, n = 13 sessions, and n = 4 control subjects with manually built headgear, n = 22 sessions). In both behavioral paradigms, rats were water deprived and had to collect water reward (~0.02 ml).

Mice were either trained to run on a 1.1 m linear track (n = 3 mice implanted with the 3D printed head gear, n = 9 sessions), or to run on a 80 cm diameter circular maze with a diagonal path allowing the animals to run in a figure-eight pattern (same layout as the rats but a smaller maze, n = 5 control subjects with manually built headgear, n = 54 sessions). In all behavioral paradigms, mice were water deprived and had to collect water reward (~0.02 ml).

The linear tracks had ‘reward areas’ on each end where water reward was delivered via a custom-built infrared-beam triggered system. Animals only received water reward for trials in which they travelled from one reward site to the other. On the circular maze the animals ran along the central arm after which they ran along the outer circle in a alternation fashion. Water reward was delivered in the reward area on correct trials.

The behavior of the animals was recorded using the Optitrack camera system (NaturalPoint, Inc, OR, USA).

Spike sorting and data processing

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Spike sorting was performed semi-automatically with KiloSort (Pachitariu, 2016) https://github.com/cortex-lab/KiloSort, using our pipeline KilosortWrapper (a wrapper for KiloSort, https://github.com/petersenpeter/KilosortWrapper) (Petersen et al., 2020b), followed by manual curation using the software Phy (https://github.com/kwikteam/phy) and our own designed plugins for phy (https://github.com/petersenpeter/phy-plugins). Finally, we processed the manually curated spike sorted data with CellExplorer (Petersen and Buzsáki, 2020) and performed further analysis with custom Matlab code.

Data availability

All documentations for parts and device fabrication are included in the manuscript and supporting files, including video recordings. The same information is made public via GitHub (https://github.com/buzsakilab/3d_print_designs/tree/master/Microdrives/Metal_recoverable; copy archived at https://archive.softwareheritage.org/swh:1:rev:a073716d89c32f13eb76a5ac5e7fa6f7fa11e18a) and the associated website: https://buzsakilab.github.io/3d_print_designs/. Data from example electrophysiological recordings are available here (https://buzsakilab.com/wp/projects/entry/65723/).

The following data sets were generated
    1. Vöröslakos Ml
    2. Petersen PC
    3. Buzsáki Gr
    (2021) Buzsaki lab databank
    ID 65723. Metal microdrive and head cap system for silicon probe recovery in freely moving rodent.
The following previously published data sets were used
    1. Petersen PC
    2. Buzsáki Gr
    (2020) Buzsaki lab databank
    ID 4919. Theta rhythm perturbation by focal cooling of the septal pacemaker in awake rats.

References

  1. Conference
    1. Scholvin J
    (2016) Heterogeneous neural amplifier integration for scalable extracellular microelectrodes
    2016 38th Annual International Conference of the IEEE Engineering in Medicine and Biology Society (EMBC). pp. 2789–2793.
    https://doi.org/10.1109/EMBC.2016.7591309

Decision letter

  1. Laura L Colgin
    Senior and Reviewing Editor; University of Texas at Austin, United States
  2. Liset M de la Prida
    Reviewer; Instituto Cajal, Spain
  3. Ashley L Juavinett
    Reviewer; University of California, San Diego, United States

Our editorial process produces two outputs: i) public reviews designed to be posted alongside the preprint for the benefit of readers; ii) feedback on the manuscript for the authors, including requests for revisions, shown below. We also include an acceptance summary that explains what the editors found interesting or important about the work.

Acceptance summary:

The authors describe a customizable and versatile microdrive and head cap system for silicon probe recordings in freely moving rodents. The added value in this work beyond previous designs is: a) a carefully designed solution to facilitate probe recovery, thus reducing experimental costs and favoring reproducibility; b) flexibility to accommodate several microdrives and additional instrumentation; c) open access design and documentation to favor customization and dissemination. The design is quite simple and versatile, and the authors provide detailed instructions. This will be a very useful resource for many readers who are interested in performing multi-site silicon probe recordings of large ensembles of neurons in freely behaving rodents. The method will appeal to a broad range of systems neuroscientists who seek to understand neurophysiological mechanisms underlying cognition and behavior.

Decision letter after peer review:

Thank you for submitting your article "Metal microdrive and head cap system for silicon probe recovery in freely moving rodent" for consideration by eLife. Your article has been reviewed by 3 peer reviewers, and the evaluation has been overseen by Laura Colgin as the Senior Editor. The following individuals involved in review of your submission have agreed to reveal their identity: Liset M de la Prida (Reviewer #1); Michael Okun (Reviewer #2), and Ashley L Juavinett (Reviewer #3).

The reviewers have discussed their reviews with one another, and the Reviewing Editor has drafted this to help you prepare a revised submission. We have also prepared public reviews of your work, which are designed to transform your unrefereed author manuscript into a publicly accessible refereed preprint (read more about this in the "Posting public reviews" section below).

Essential revisions:

Reviewers were enthusiastic about the potential impact of this work. However, some concerns were raised regarding transparency, specific quantitative support of usability measurements, and clearer descriptions regarding the novelty of this resource compared to other previously published resources. Showing long-term recordings is recommended, but it is expected that the authors already have these recordings on hand. That is, reviewers believe that no new data are required, simply an inclusion and analysis of existing data. Authors are also requested to be more specific and transparent about which probes work with this resource. Specific details can be found in the three independent reviews, which were remarkably consistent and are included in their entirety below. Reviewers would like to see authors addressing all of their comments.

Reviewer #1 (Recommendations for the authors):

Comments and suggestions (the order is not informative)

1. The report reads very descriptive. While authors declare having used n = 7 rats and n=5 mice in methods, this is not exploited quantitatively in the paper. The ms will benefit from including some assessment to support ease of use (e.g. behavior in implanted versus not implanted animals), stability of recordings over days, weeks or months; or any other relevant supportive example.

2. Page 4, line 100 and table 1: What travel distance this refers to? The effective travel distance should be better explained. Once the probe is mounted in the shuttle, the travel distance is limited by the probe length and tip location from the microdrive base. Unexperienced users may misunderstood this important issue.

3. Table 1: What are inclusion criteria? Are these microdrives designed for tetrodes or for silicon probes; mice only? Korshunov 2006 is for single wires in a wide range of species and I fail to see how it could be repurposed for silicon probes. If the goal of the table is to summarize existing solutions, then it may fall short for including other microdrive designs. I would recommend focusing. Is Voroslakos et al., referring to this study ? Please, clarify.

4. Figure 1D: the stereotaxic microdrive holder is very similar to that in Chung et al., (Figure 4). Is that the case? What are the slots in the attachment for?

5. Figure 2F-H and Figure 3D: In general, data is very descriptive and poorly relate with the potential added value of the system. I would consider providing some quantitative assessment to prove stability, or reusability, or consistency of some LFP feature between animals. What metrics is shown in Figure 2H and why is this useful to illustrate the added value of the microdrive? Same for Figure 2F.

6. Introduction, line 49: authors make a good case for the emergence of novel integrated solutions such as uLED and microfluidic probes which benefit from the use of versatile headgear designs. However, both Wu et al. and Kim et al. seem to report on uLEDs only. Consider Altuna et al., Lab on Chip 2013 https://doi.org/10.1039/C3LC41364K of any other appropriate reference for integrated fluidic multi-site probes, or just avoid the mention to microfluidity.

7. Introduction, lines 52-54: flexDrive, shuttledrive, etc. all of them are mostly dedicated to carry tetrodes. There are fewer designs for silicon probes, which may support the need for this paper, but authors avoid discussing the potential overlapping and added value between existing solutions and their own design. I feel the introduction will benefit from addressing this more sharply.

8. Videos: Only two videos are provided to illustrate the head-cap systems of rats and mice. While documentation provided is useful, many relevant parts of the paper will strongly benefit from providing video support (e.g. microdrive assembly, implantation, recover).

9. Methods, line 352: stl files are provided via a Github. It would be useful to upload all of them together as supplementary material of the paper itself. Also, please, consider adding specific links to the different files repository in the Methods section.

10. Methods, line 369: authors recommend measuring probe impedance. This is not particularly easy with silicon probes. Please, add equipment information.

Reviewer #2 (Recommendations for the authors):

This manuscript provides an updated guide on the procedures for performing chronic recordings with silicon probes in mice and rats in the lab of the senior author, who is one of the leaders in the use of this experimental method. The new set of procedures relies on metal and plastic 3D printed parts, and represents a major improvement over the older methodology (i.e. Vandecasteele et al. 2012).

The manuscript is clearly written and the technical instructions (in the Methods section) seem rather detailed. The main concerns I had are as follows.

1. The present design is an improvement over Chung et al., (the most similar previously published explantable microdrive design, as far as I am aware) in terms of the footprint and travel distance. However, a main disadvantage of the system in its present form is that (apparently) it does not support Neuropixels probes. While such probes might not be suitable for some uses (e.g. to record from large populations in dorsal hippocampus), Neuropixels probes are of considerable interest to many labs.

2. The total weight of the mouse implant seems quite high (together with the headstage, I estimate it is >= 4gr). Could the authors provide the exact value, and describe whether this has any impact on the way the animal moves? Also, the authors should describe how the animals are housed (e.g. do they carry the headstage even when not being recorded). The authors say that a mouse can be implanted with more than one microdrive. The authors should clarify whether they actually have an experience with such implants, or is this just a suggestion based on their educated estimate?

3. There is no information in the Results section on the number of implants performed, the duration the animals were implanted, the quality of the recordings obtained, number of successes or failures. The figures merely provide examples of one successful recording in a mouse and in a rat. All these details should be provided, along with details of how many probes were reused and how many times (a brief mention of one case, lines 252-253 and 359-360, is not sufficient).

4. In Figure 2, spike waveforms are classified as pyramidal, wide or narrow interneurons. I did not find any description of how this classification was performed.

5. Also in Figure 2, refractory period violations are reported in percent (permille in fact). First, it is not clear how refractory period was defined. Second, such quantification is incorrect in principle: we use refractory period violations to infer the rate of false positives. Yet the relationship between fraction of ISI violations and false positive rate depends on the firing rate of the neuron. For example, 0.1% of ISI violations is quite good for a unit spiking at 10 spikes/s, is so so for a unit spiking at 1 spike/s, and is very bad if the firing rate is 0.1 spike/s (see Hill et al. JNeurosci. 2011 for derivation). Alternatively, the authors can follow an approach described in an old paper by the same lab (Harris et al., JNeuropsysiol. 2000), quantifying the violations in spike autocorrelogram relative to its asymptotic height.

6. Line 477: the authors write that the probes were mounted on a plastic microdrive. This seems to contradict the key claim of the manuscript (namely that the microdrives were from stainless steel).

7. I believe that the work of Luo and Bondy et al., (eLife 2020) and should be references and compared to.

Reviewer #3 (Recommendations for the authors):

First, I'd like to applaud the authors for the development of a very clever device and for their clear description of how to use it. With the inclusion of more details around the compatibility of this device with specific probes, more support for specific claims in the manuscript, and more transparency about the success of probe recovery, this manuscript will inevitably serve as an important resource for many researchers.

I'll leave one thought for consideration here before diving into specifics: the organization of this manuscript was a bit unclear to me. I think it makes sense to have a "Results section" which gives a high level description of the procedure and your design choices, as well as a "methods section" which outlines the protocol, but this needs to be clear. This could be solved by a line at the end of the introduction that says something like "Here we'll describe the results we obtained and a high level summary …. Readers can find detailed instructions in the methods as well as in the attached files…". I would also defer to the eLife editors for how they would like to handle this organization.

The introduction of the paper is very well written, but towards the end there are several unsubstantiated claims. Specifically, the idea that this design reduces surgery time substantially. Can you be more specific, or back this up with timelines from other designs? On line 203 there is a reference to another paper after this claim, but this could be made more clear in the introduction. Which aspect of this procedure is quicker than other procedures? As I'll come to later in this review, there is also a claim about the recovery being "reliable" – knowing exactly how reliable, given your experience, would be extremely useful.

Overall, I have two suggestions that would greatly improve this manuscript.

First, more neural data should be shown. In Figure 2 E-H and Figure 3, some neural data recorded using this device is shown, but this is not nearly enough for users to assess the usability of these probes. The mouse data is particularly sparse, and it is very difficult to make much out of Figure 3D without seeing isolated units. If 7 rats and 5 mice were recorded, more of this data should be shown. Specifically, users may be interested in seeing the stability of units over time, as well as the SNR levels on various days of recording. The methods there was daily recording – showing some of this data would be useful. How long has one of these devices been successfully used to record activity? These types of details are essential to ensuring that your device enables quality data collection.

Secondly, there should be a table detailing each procedure done with these devices, including the type of animal (mouse/rat), age (if available), sex, success of recording (and for how many days/weeks), and success of the probe recovery (beyond saying it is "highly successful", line 257). This comprehensive overview of exactly how reliable your device is will be very useful to readers.

Several aspects of the manuscript could be clarified.

1. The "footprint" of the device is given multiple times, but a height and width would also be useful.

2. There is a clear trade-off between the minimal wear and increased reusability of metal drives with the weight of these drives, and that should be acknowledged. Would it be possible to create such a drive with a sturdy but lighter plastic, and if not, why?

3. Line 105 says the microdrive weighs 0.87 g – is this without any materials to attach it to the skull? Similarly, the overall weight of the entire assembly should be given.

4. It is unclear what material the stereotaxic attachment is made of.

5. There doesn't seem to be any mention of how to actually attach the probe to the arm/shuttle. Is it glued? Which probes were used should be clear in the Results section (I see it is eventually mentioned in the methods). Relatedly, it should be clear which types of probes were tested with this microdrive, and which probes you would recommend using with it. Specifically, will this microdrive and assembly work with Neuropixels 1.0 probes? Being clear about which probe was used is especially important for the multi-probe implantation – presumably this will not work with probes with large PCB boards and/or headstages. Also, are multi-probe implantations possible in mice using your microdrives and assembly?

6. It is unclear how the headstage is affixed in either the rat or mouse assembly.

7. For probe recovery, it's important to note that distilled water will not be recommended for all probes. For example, neuropixels have very clear restrictions on what you should use with them. I would advise the reader accordingly. Tergazyme may be useful here as well.

8. How is grounding handled in these devices? There are multiple mentions of a skull screw used to affix the protective assembly, but in most designs, a skull screw is there to serve as a reference and/or grounding. (Sidenote: It is not clear to me why a ground screw is so bad for the animal, as is emphasized multiple times in the manuscript. We are also putting a large open whole in the skull…) Is there a ground wire in these devices? Similarly, is the copper mesh inside electrically connected to the probe, or is it kept isolated?

9. Line 476 in methods is very unclear and mentions a plastic microdrive: "The probes ere mounted on a plastic recoverable microdrive to allow precise vertical movement after implantation (github.com/YoonGroupUmich/Microdrive) and implanted by attaching the base of the micro-drives to the skull with dental cement." Is this the same microdrive mentioned in the main manuscript? In general, it is unclear how the "Additional implantation information" relates to the main methods and seems that this information should be integrated into the other methods sections.

10. There is no mention of where to download the design files.

11. Data is provided for only 2 animals (one mouse, one rat) and 2 sessions – could more data be made available?

12. Code is provided for sorting (Kilosort Wrapper and phy plugin).

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "Metal microdrive and head cap system for silicon probe recovery in freely moving rodent" for further consideration by eLife. Your revised article has been evaluated by Laura Colgin (Senior Editor) and a Reviewing Editor.

The manuscript has been improved but there are some remaining issues that need to be addressed, as outlined in the individual reviews below:

Reviewer #1 (Recommendations for the authors):

Authors have addressed all comments. The revised version is substantially improved. Videos are superb useful as well as the accompanying information.

Reviewer #2 (Recommendations for the authors):

I would like to thank the authors for carefully and thoroughly addressing the concerns and comments that I have raised. I believe that the revised version (including the addition of the videos illustrating the key procedures) is significantly improved.

I would like to encourage the authors to incorporate into the final version of the paper all the relevant technical details from the rebuttal. For example, in their response the authors mention using a pulley to counterbalance the headstage (yet this seems not to be mentioned in the manuscript), and similarly with preferentially using large (>35gr) mice.

Reviewer #3 (Recommendations for the authors):

This manuscript has improved significantly since the initial submission with numerous additional figures, tables, and methodological details that address the initial concerns. The reviewers have added multiple useful insights to the manuscript, including the footprint of the device, more clear justification for the use of stainless steel, the material of the implantation tool. The additional videos explaining how to assemble the headgear are very well done and clear. These details will undoubtedly help readers who wish to implement this headgear system in their own labs. The addition of more quantification of the neural data is also very appreciated. The authors have added additional analysis for Figure 2, showing single units for the mouse recording (though it would be useful to have a similar analysis in Figure 3 for the rat data).

There are several points that should be addressed within the revised manuscript:

The new Figure 5 illustrates the use of two different recording devices in mice. Figure 5A demonstrates the units recorded over time on each shank as the probe is lowered each day. However, it is unclear to me how Figure 5B relates to 5A – in 5B it looks as if all four shanks are at the same depth, whereas in 5A they are at different depths. Ultimately, it seems like some sort of integration representation of these two panels where viewers can appreciate the location of single units on each shank over days would be the most useful. On this same figure, the axis labels for 5F and 5I are a bit misleading, because this is not about the noise level of the recording, but about the waveform. I'd suggest changing the figure and the wording in the text to "Waveform relative noise level" so that readers do not confuse this with overall signal to noise in the recording. The axis on 5G could be readjusted so that readers can appreciate the data. In the figure caption, I'd suggest spelling out "ACGs" for readers unaccustomed to that shorthand. Presumably these ACGs are from the unit in the box? If so, I'd clarify in the caption.

Related to Figure 5, the corresponding text says, "suggesting either that the distance between the electrode sites and neuron bodies decreased or that large size neurons were recorded," but it is important to note that the probe was being moved into different brain areas over these days of recording. The text should be modified to clarify this – the clear difference from day 5 to the other days is almost definitely explained by the movement to deeper brain structures. This paragraph should also note the clear decrease in the # of units with the reimplanted probe, as well as the clear increase in the noise level in the reimplanted probe.

The authors have also added behavioral data, shedding light on the ability of animals to move with this headgear, however some clarity around these behavioral findings is needed. On line 219 it reads, "The 3D printed head cap system is comparable in weight to manually built headmounts" which is an abrupt and unclear transition to the paragraph about the impact of the headgear on behavior – reading between the lines, I think the authors are saying, "… however, we wanted to verify still that our headgear would not impede the animal's behavior." It is also unclear from this paragraph how this behavior was measured. The methods section describes mice and rats running on a track – were they headfixed? It is also unclear how the water reward is relevant here – did animals need to collect a water reward in order for the track to continue moving? Also, the mice with 3D printed headgear ran on a different track than the mice with manually printed headgear – some mention of this in the main text is warranted for transparency in interpreting the behavioral results shown. The wording in the methods is also unclear – is the track circular or a figure eight? Finally, contextualizing the results found here in terms of typical speeds on such a treadmill (e.g., in the discussion, see the point below) would be very useful to readers.

In this same paragraph, the authors state they found a "small significant difference," but this wording is misleading. A statistical test result is or is not significant, and the authors should remove the word small. Readers can determine whether or not the absolute value of the p-value is informative. There is a typo in this section also: "We also performed the same test on mice subjects and found a small significant difference between the median running speed of the two *rat* groups (KS-test; p = 0.045) but no significant different between the 95 percentile speeds (KS-test; p = 0.24)." I would also recommend that the authors write out the full name of the KS test, at least on the first mention.

Line 139 says, "High-quality electrophysiological signals can be collected from freely moving mice for weeks and months (Figure 2E and F)" however it is unclear from this figure whether this data was indeed recorded over weeks and months. If this is a claim the authors cannot substantiate with data, it should be removed. Figure 5 does indeed show "weeks" with the headgear (showing data out to day 16 for the 4-shank probe and 17 for neuropixels), so it seems like an applicable figure to reference here, without the mention of "months." Similar changes should be made for the reference to the rat recordings and Figure 3 – it's unclear based on the figure caption how long after implant these extracellular traces were obtained.

Finally, although these are significant additions to the main portion of the text, there is no mention of either the single-unit figure or the behavioral results in the discussion. A contextualization of these data in terms of the headgear's usability as well as a comparison of these findings to other recoverable drives would be useful and warrant discussion.

https://doi.org/10.7554/eLife.65859.sa1

Author response

Reviewer #1 (Recommendations for the authors):

Comments and suggestions (the order is not informative)

1. The report reads very descriptive. While authors declare having used n = 7 rats and n=5 mice in methods, this is not exploited quantitatively in the paper. The ms will benefit from including some assessment to support ease of use (e.g. behavior in implanted versus not implanted animals), stability of recordings over days, weeks or months; or any other relevant supportive example.

We added a table to the manuscript to better describe what kind of cap and microdrive system was used in each animal (Supplementary File 1). Our developmental process was the following: first, we tested the ease of use and longevity of the cap systems in a chronic rat (R_01, n = 90 days) and a chronic mouse (M_01, n = 60 days). After this initial test, we implanted flexible probes with the cap system to test longevity of electrophysiology recordings using our cap system both in a mouse and a rat (M_02, 32-channel flexible probe, n=27 days and R_02, 64-channel flexible probe, n = 76 days). Finally, we tested the cap systems with our recoverable, plastic microdrive system which we have been using for the last two years in our laboratory. We used 32 and 64-channel silicon probes, Neuropixels probes as well as tungsten wire electrodes in mice and rats (for details see Supplementary File 1). After the initial tests, we changed from plastic recoverable to metal recoverable microdrives. In the first submission, we tested our full system (cap with metal, recoverable microdrive) in a mouse and a rat and we are continuously implanting animals using these newly developed methods. Since our submission, we have performed two chronic and two acute experiments in rats and two chronic experiments in mice.

Since our original submission, our lab has performed the following experiments.

R_08Long Evans330RPlastic recovNeuropixels from R07HippocampusChronicOngoingNAHomecage
R_09Long Evans345RMetal recovNeuropixelsHippocampusChronicOngoingNAHomecage linear maze
R_01ALong Evans430NAMetal recov128-5 (DB) from M05HippocampusAcuteNANANA
R_02ALong Evans430NAMetal recov128-5 (DB) from R_01AHippocampusAcuteNA2 shank brokeNA
M_06C57BL/631MPlastic recovMicro-LEDHippocampusChronicOngoingNAHomecage linear maze
M_07C57BL/627MPlastic recovMicro-LEDHippocampusChronicOngoingNAHomecage linear maze
M_08DBA/2J34MMetal recov128-5 (DB) from R02_AHippocampusChronicOngoingNAHomecage
M_09C57BL/628BMetal recovAssy-E1 (CN)Hippocampus, entorhinal cortexChronicOngoingNAHomecage radial arm maze

2. Page 4, line 100 and table 1: What travel distance this refers to? The effective travel distance should be better explained. Once the probe is mounted in the shuttle, the travel distance is limited by the probe length and tip location from the microdrive base. Unexperienced users may misunderstood this important issue.

The travel distance refers to the overall distance that the arm can move from the top to the bottom of the drive body. The reviewer is right that travel distance depends on many factors, including length of shank(s), initial distance between arm and bottom of the drive body. We added a sentence to the revised manuscript for clarification.

3. Table 1: What are inclusion criteria? Are these microdrives designed for tetrodes or for silicon probes; mice only? Korshunov, 2006 is for single wires in a wide range of species and I fail to see how it could be repurposed for silicon probes. If the goal of the table is to summarize existing solutions, then it may fall short for including other microdrive designs. I would recommend focusing. Is Voroslakos et al., referring to this study ? Please, clarify.

Our inclusion criteria was the overall weight of the microdrive system, regardless of whether it uses silicon probes or wires. In the revised manuscript, we included microdrive systems that used silicon probes in mice. We also added an extra line in the legend explaining our inclusion criteria.

4. Figure 1D: the stereotaxic microdrive holder is very similar to that in Chung et al., (Figure 4). Is that the case? What are the slots in the attachment for?

We have been working with Sebastian Royer’s and Kamran Diba’s groups since 2017. They shared their design files which served as a basis of our microdrive and head cap developments. Our design is fundamentally the same as in Chung et al., and we make this clear in the text and describe the steps of improvements below.

Chung et al., 2016 design was designed for mice only. Hiroyuki Miyawaki has adapted Chung’s design for rats. We adapted Miyawaki’s design for mice (shorter travel distance) and made the following improvements over the years:

1. v01

a. Added nut to arm (reduced metal-to-plastic erosion).

2. v14

a. Updated shell component

i. Improved stability.

ii. Reduced footprint from 5.2x7.5 mm to 3.2x7.5mm. This enabled bilateral silicon probe implantation in the CA1 region of the hippocampus of rats (Rogers et al., 2021).

b. Reduced weight allowed us to use the same drive in mice (Valero et al., 2021) and rats. (We do not use the shorter mouse microdrive anymore).

3. v19

a. Added second male header pin to drive body (further improved stability).

4. v20

b. Increased thickness of the bottom part of the drive. Drive body became sturdier, less prone to bending.

5. v21

a. Improved arm design to overcome limitations of resin 3D printing. Most of the arms were not at 90-degree angles.

6. Metal prototype printed in plastic

a. Enables double silicon probe implantation in the mouse.

b. Nut is moved from the bottom of the drive to the top. This creates a flat surface at the bottom of the drive body making the shell-drive connection more stable.

7. 3D printed, metal recoverable microdrive.

5. Figure 2F-H and Figure 3D: In general, data is very descriptive and poorly relate with the potential added value of the system. I would consider providing some quantitative assessment to prove stability, or reusability, or consistency of some LFP feature between animals. What metrics is shown in Figure 2H and why is this useful to illustrate the added value of the microdrive? Same for Figure 2F.

We agree that Figure 2E and F did not illustrate the quality of the single cells well. It merely served to illustrate recordings of the single cells through the raw traces, approximated position and z-scored waveforms. Figure 2G and 2H show the quantitative measures of the single unit data. 2H illustrates the distribution of the refractory period violations in the recorded population (a common quality measure) and their spike amplitude. In the revised manuscript, we dedicate a full new figure (Figure 5) to the quantification of single cell features. We want to emphasize though that our head gear development is not primarily about long-term stability but for reusing silicon probes and head gear components with ease, decreasing the effective cost, experimental effort and complexity. The metal drive has the potential to allow for better long-term recordings, but we do not have firm quantitative data to support this, since this was not the goal of our experiments. For long-term recordings of multiple single units different technologies are needed (flexible probe recordings from the Frank group (Chung et al., 2019); Lieber group’s injectable electrode (Schoonover et al., 2020; Zhou et al., 2017)).

6. Introduction, line 49: authors make a good case for the emergence of novel integrated solutions such as uLED and microfluidic probes which benefit from the use of versatile headgear designs. However, both Wu et al. and Kim et al. seem to report on uLEDs only. Consider Altuna et al., Lab on Chip 2013 https://doi.org/10.1039/C3LC41364K of any other appropriate reference for integrated fluidic multi-site probes, or just avoid the mention to microfluidity.

In response to the Reviewer’s comment, we have removed reference to microfluidity from the revised manuscript because microfluidic probes require different solutions.

7. Introduction, lines 52-54: flexDrive, shuttledrive, etc. all of them are mostly dedicated to carry tetrodes. There are fewer designs for silicon probes, which may support the need for this paper, but authors avoid discussing the potential overlapping and added value between existing solutions and their own design. I feel the introduction will benefit from addressing this more sharply.

In our revised introduction, we address the different needs of tetrode and silicon drives. A disadvantage of tetrode recordings is that each tetrode needs to be moved separately.

8. Videos: Only two videos are provided to illustrate the head-cap systems of rats and mice. While documentation provided is useful, many relevant parts of the paper will strongly benefit from providing video support (e.g. microdrive assembly, implantation, recover).

In response to the Reviewer’s comment, we have created the following additional videos:

– Figure 1-video 1, showing how to assemble the recoverable metal microdrive.

– Figure 1-video 2, showing how to attach a Neuropixels probe to a recoverable, metal microdrive.

– Figure 2-video 1, showing how to assemble the mouse cap.

– Figure 3-video 1, showing how to assemble the rat cap.

We have also added the following two videos.

– Figure 4-video 1, showing how to recover a silicon probe from a mouse cap.

– Figure 4-video 2, showing how to recover a silicon probe from a rat cap.

9. Methods, line 352: stl files are provided via a Github. It would be useful to upload all of them together as supplementary material of the paper itself. Also, please, consider adding specific links to the different files repository in the Methods section.

Pending on acceptance of our revised manuscript, we will work with the editor to add our stl files as Supplementary material. We added the appropriate links in the Methods section.

10. Methods, line 369: authors recommend measuring probe impedance. This is not particularly easy with silicon probes. Please, add equipment information.

To collect electrophysiology data, we recommend using the RHD USB interface board from Intan (Intan Technologies LLC, CA, USA). This system enables users to measure the impedance of the attached device. For more details see:

https://intantech.com/files/Intan_RHD2000_eval_system.pdf page 14 Electrode Impedance Measurement section. This measurement provides a quick and rough estimate about the quality of the recording sites, whether they are below or above 2 MOhm (we can collect good wide-band signal if the impedance of the recording sites is below 2 MOhm). If higher precision is required, users can perform impedance measurement with NanoZ device (nanoZ Impedance Tester, Plexon Inc, TX, USA).

We added this information to the revised manuscript.

Reviewer #2 (Recommendations for the authors):

This manuscript provides an updated guide on the procedures for performing chronic recordings with silicon probes in mice and rats in the lab of the senior author, who is one of the leaders in the use of this experimental method. The new set of procedures relies on metal and plastic 3D printed parts, and represents a major improvement over the older methodology (i.e. Vandecasteele et al. 2012).

The manuscript is clearly written and the technical instructions (in the Methods section) seem rather detailed. The main concerns I had are as follows.

We thank the reviewer for carefully reading our manuscript and providing useful and constructive comments.

1. The present design is an improvement over Chung et al. (the most similar previously published explantable microdrive design, as far as I am aware) in terms of the footprint and travel distance. However, a main disadvantage of the system in its present form is that (apparently) it does not support Neuropixels probes. While such probes might not be suitable for some uses (e.g. to record from large populations in dorsal hippocampus), Neuropixels probes are of considerable interest to many labs.

Our microdrive and head cap system can also support Neuropixels probes. Since our initial submission, we have implanted a Neuropixels probe in the intermediate hippocampus of a rat using our recoverable, plastic microdrive. At the end of the experiment, the Neuropixels probe was successfully recovered, cleaned, and implanted again in a new rat. In addition, we designed a new arm for our metal microdrive which can support Neuropixels probes (Author response image 1) and implanted another rat (Author response image 2 and Figure 3—figure supplement 2).

We have also created a video showing how to attach Neuropixels probe to a metal microdrive (Figure 1-video 2).

Author response image 1
Metal microdrive adapter for Neuropixels probe.

A. Arm design for 64-channel silicon probes. 45o, front, side and top views are shown (from left to right). All dimensions are in mm. B. Changing the overall length (from 7.35 mm to 10 mm) and width (from 4 mm to 5.4 mm) of the 64-channel arm makes our metal microdrive compatible with Neuropixels probe. Note, that only three dimensions of the 64-channel arm were modified (red numbers). 45-degree, front, side and top views are shown (from left to right). All dimensions are in mm. C. Photograph of the different arm designs of the metal, recoverable microdrive (top shows an arm designed for a 64-ch silicon probe, bottom shows an arm designed for Neuropixels probe).

Author response image 2
Recording of unit firing with Neuropixels probe attached to a metal microdrive in freely moving rat.

A. Metal microdrive for Neuropixels probe (a – stereotax attachment, b – drive holder, c – metal microdrive, d – Neuropixels probe and e – Neuropixels headstage). B. Photo of Neuropixels probe attached to a metal microdrive (a-e same as in A). C. Location of probe implantation (Bregma – 4.8 mm, mediolateral + 4.6 mm, 11-degree angle). D. High pass filtered traces (1s) from a freely moving rat implanted with Neuropixels probe. Note the single unit activity in the cellular layer of cortex (top) and hippocampus (bottom).

2. The total weight of the mouse implant seems quite high (together with the headstage, I estimate it is >= 4gr). Could the authors provide the exact value, and describe whether this has any impact on the way the animal moves? Also, the authors should describe how the animals are housed (e.g. do they carry the headstage even when not being recorded). The authors say that a mouse can be implanted with more than one microdrive. The authors should clarify whether they actually have an experience with such implants, or is this just a suggestion based on their educated estimate?

The total weight of the metal microdrive, including the base, body and arm is 0.87 gram. Additional weight is the metabond and dental acrylic cement. The amount of cement that is used during surgery can vary between researchers and the type of surgery. The overall weight of the assembly also depends on the silicon probe with Omnetics connector(s) that is used for the surgery, e.g.: 32-channel micro-LED probe is 1.11g (NeuroLight Technologies LTD.), 64-channel 4-shank probe is 0.96g (ASSY E-1, Cambridge NeuroTech), 64-channel 5-shank probe is 1.05g (A5x12-16-Buz-Lin-5mm, NeuroNexus Ltd.) and a 128-channel 4-shank probe with integrated Intan chips is 0.94g (P128-5, Diagnostic Biochips). In addition, the overall weight of the entire assembly can change if optic fibers are used in optogenetic studies or if any custom connectors are implanted (e.g., connector and wires for brain stimulation). That is the reason why we reported the overall weight of each system (metal microdrive, mouse cap and rat cap) individually.

The implanted mice are single housed, and they do not carry the headstage while in the vivarium. During recordings, the headstage is attached and a counterbalanced pulley system ensures that the animal is not carrying the extra weight of the headstage. We have quantitatively compared running speed with traditional and the new head caps in both rats and mice (Figure 6).

The small footprint of the metal microdrive enables researchers to perform more than one silicon probe implantation in freely moving mice. For this purpose, larger mice (>35 g) are selected (Author response image 3).

Author response image 3
Metal microdrive enables double silicon probe recordings in freely moving mice.

A. Intraoperative photograph of double silicon probe implantation. Note that the metal microdrive on the left had been secured to the skull and the second drive is being implanted using the stereotaxic attachment and drive holder. The probe PCBs are placed on the copper mesh. B. Photograph focused on the metal microdrives.

3. There is no information in the Results section on the number of implants performed, the duration the animals were implanted, the quality of the recordings obtained, number of successes or failures. The figures merely provide examples of one successful recording in a mouse and in a rat. All these details should be provided, along with details of how many probes were reused and how many times (a brief mention of one case, lines 252-253 and 359-360, is not sufficient).

We have added Supplementary File 1 explaining all the details of our implants. We would like to refer the Reviewer to response #1 to Reviewer 1.

Adapting new technology is challenging. To date, we have extensive experience with the rat cap system only (n=3 users in the lab, n = 25 rats implanted). Two lab members have started to adapt our mouse cap and implanted 3 mice since our submission. We included their maze running behavioral data for comparison between the copper mesh and cap system.

Prior to the development of the metal microdrive, we have conducted an internal lab survey comparing the hand-made microdrive (Vandecasteele et al., 2012) and our recoverable, plastic microdrive. Six lab members who had extensive experience with both types participated (Figure 1—figure supplement 2). Our questions were:

1. On a scale 1-10, how would you compare the plastic, recoverable drive to the Vandecasteele et al., 2012 one in terms of: (a) ease of building a drive, (b) size and (c) ease of recovery.

Overall, the success rate of recovery is much higher using a recoverable microdrive system, but the size of the plastic, recoverable microdrive is limits certain experiments. This was one of the main motivations to develop the metal, recoverable microdrive.

4. In Figure 2, spike waveforms are classified as pyramidal, wide or narrow interneurons. I did not find any description of how this classification was performed.

We have removed the single cell putative cell types from the manuscript as this issue is not relevant to the current manuscript. Figure 2 has been simplified and a new figure 5 is dedicated to the single cell quantification.

5. Also in Figure 2, refractory period violations are reported in percent (permille in fact). First, it is not clear how refractory period was defined. Second, such quantification is incorrect in principle: we use refractory period violations to infer the rate of false positives. Yet the relationship between fraction of ISI violations and false positive rate depends on the firing rate of the neuron. For example, 0.1% of ISI violations is quite good for a unit spiking at 10 spikes/s, is so so for a unit spiking at 1 spike/s, and is very bad if the firing rate is 0.1 spike/s (see Hill et al. JNeurosci. 2011 for derivation). Alternatively,

the authors can follow an approach described in an old paper by the same lab (Harris et al., JNeuropsysiol. 2000), quantifying the violations in spike autocorrelogram relative to its asymptotic height.

We have removed this panel from Figure 2 and dedicated a new figure (Figure 5) to the single cell quantification. Refractory violations can be used as an alarm for poor cluster quality. Absence of refractory violations alone does not guarantee good separation for the reasons the Reviewer mentioned.

6. Line 477: the authors write that the probes were mounted on a plastic microdrive. This seems to contradict the key claim of the manuscript (namely that the microdrives were from stainless steel).

We apologize if this description was not clear in the original manuscript. In the revised version, we have added a table (Supplementary File 1) explaining all details of each animal subject (species, strain, weight, cap type), type of silicon probe and microdrive used. As we explained in Response 3, our main goal was to test each system individually and once all components have been verified, we combined everything into one surgery.

The plastic and metal microdrives are based on the same principles. The implantation/recovery tools are also identical in design concepts. Based on our own experience, users dol not recognize any changes in terms of ease of use, ease of implantation and ease of recovery when changing from plastic recoverable microdrives to metal ones. The advantage of metal drives is size reduction, their multiple reusability and stability.

7. I believe that the work of Luo and Bondy et al., (eLife 2020) and should be references and compared to.

We reference Luo et al., (2020) in our revised manuscript. One of the main advantages of using a microdrive system is the ability to move the recording probe inside the brain tissue and sample new sets of neurons. This is not the case in Luo and Bondy et al., (eLife 2020).

Reviewer #3 (Recommendations for the authors):

First, I'd like to applaud the authors for the development of a very clever device and for their clear description of how to use it. With the inclusion of more details around the compatibility of this device with specific probes, more support for specific claims in the manuscript, and more transparency about the success of probe recovery, this manuscript will inevitably serve as an important resource for many researchers.

I'll leave one thought for consideration here before diving into specifics: the organization of this manuscript was a bit unclear to me. I think it makes sense to have a "Results section" which gives a high level description of the procedure and your design choices, as well as a "methods section" which outlines the protocol, but this needs to be clear. This could be solved by a line at the end of the introduction that says something like "Here we'll describe the results we obtained and a high level summary …. Readers can find detailed instructions in the methods as well as in the attached files…". I would also defer to the eLife editors for how they would like to handle this organization.

The introduction of the paper is very well written, but towards the end there are several unsubstantiated claims. Specifically, the idea that this design reduces surgery time substantially. Can you be more specific, or back this up with timelines from other designs? On line 203 there is a reference to another paper after this claim, but this could be made more clear in the introduction. Which aspect of this procedure is quicker than other procedures? As I'll come to later in this review, there is also a claim about the recovery being "reliable" – knowing exactly how reliable, given your experience, would be extremely useful.

We thank the reviewer for carefully reading our manuscript and providing useful and constructive comments.

Overall, I have two suggestions that would greatly improve this manuscript.

First, more neural data should be shown. In Figure 2 E-H and Figure 3, some neural data recorded using this device is shown, but this is not nearly enough for users to assess the usability of these probes. The mouse data is particularly sparse, and it is very difficult to make much out of Figure 3D without seeing isolated units. If 7 rats and 5 mice were recorded, more of this data should be shown. Specifically, users may be interested in seeing the stability of units over time, as well as the SNR levels on various days of recording. The methods there was daily recording – showing some of this data would be useful. How long has one of these devices been successfully used to record activity? These types of details are essential to ensuring that your device enables quality data collection.

We thank the Reviewer for these comments, which prompted us to add more quantifications and details. We added 3 new figures, 6 new videos and a Supplementary Table to address the Reviewer’s comment. Figure 5 is dedicated to the quantification of unit parameters.

Secondly, there should be a table detailing each procedure done with these devices, including the type of animal (mouse/rat), age (if available), sex, success of recording (and for how many days/weeks), and success of the probe recovery (beyond saying it is "highly successful", line 257). This comprehensive overview of exactly how reliable your device is will be very useful to readers.

As suggested, we have added Supplementary File 1, which list all critical technical details of all experiments included in the manuscript.

Several aspects of the manuscript could be clarified.

1. The "footprint" of the device is given multiple times, but a height and width would also be useful.

The outer dimensions of the microdrive are specified in figure 1B. All dimensions of the drive are available in Table 1. We also added the width, height, and length to the main text.

2. There is a clear trade-off between the minimal wear and increased reusability of metal drives with the weight of these drives, and that should be acknowledged. Would it be possible to create such a drive with a sturdy but lighter plastic, and if not, why?

This is an important point. Stainless steel is about 100 times less flexible than plastic (Young's modulus of stainless steel: ~180 GPa vs plastic: ~2 GPa), yet the density is only 8 times higher (steel: ~8 kg/m3, plastic ~1 kg/m3). This is why the device can be made sturdier at a similar size. Manufacturing a plastic drive with the same strength would make the plastic drive substantially larger. While plastic drives are cheaper, their reusability is more limited. Both materials have their own advantages and weaknesses. However, reusability is an important factor since preparing a new head gear requires many hours of dedicated time.

3. Line 105 says the microdrive weighs 0.87 g – is this without any materials to attach it to the skull? Similarly, the overall weight of the entire assembly should be given.

The Reviewer is correct that the total weight of the metal microdrive, including the base, body and arm is 0.87 gram. Additional weight is the metabond and dental acrylic cement. The amount of cement that is used during surgery can vary between researchers and the type of surgery. The overall weight of the assembly also depends on the silicon probe with Omnetics connector(s) that is used for the surgery, e.g.: 32-channel micro-LED probe is 1.11g (NeuroLight Technologies LTD.), 64-channel 4-shank probe is 0.96g (ASSY E-1, Cambridge NeuroTech), 64-channel 5-shank probe is 1.05g (A5x12-16-Buz-Lin-5mm, NeuroNexus Ltd.) and a 128-channel 4-shank probe with integrated Intan chips is 0.94g (P128-5, Diagnostic Biochips). In addition, the overall weight of the entire assembly can change if optic fibers are used in optogenetic studies or if any custom connectors are implanted (e.g., connector and wires for brain stimulation). That is the reason why we reported the overall weight of each system (metal microdrive, mouse cap and rat cap) individually.

4. It is unclear what material the stereotaxic attachment is made of.

The stereotaxic attachment is made of plastic (clear v4 resin from FormLabs). We have added this information in the revised manuscript.

5. There doesn't seem to be any mention of how to actually attach the probe to the arm/shuttle. Is it glued? Which probes were used should be clear in the Results section (I see it is eventually mentioned in the methods). Relatedly, it should be clear which types of probes were tested with this microdrive, and which probes you would recommend using with it. Specifically, will this microdrive and assembly work with Neuropixels 1.0 probes? Being clear about which probe was used is especially important for the multi-probe implantation – presumably this will not work with probes with large PCB boards and/or headstages. Also, are multi-probe implantations possible in mice using your microdrives and assembly?

We attach the backend of the silicon probes to the arm using cyanoacrylate glue (Loctite, #45208). We have included this step in our Methods section, and we have also added this information to the figure legend of Figure 1 in the revised manuscript.

We added a new table summarizing which probes were used with our microdrive system. We have also performed the necessary modification of the arm design to support Neuropixels 1.0 probes (Figure 2) and performed a chronic surgery in a rat. Our microdrive can support any type of silicon probes provided that the arm is customized for the right size. We do not fully understand the Reviewer comment about “large PCB boards and/or headstages”. If the PCB boards and/or headstages are small/light enough for a rodent to carry them, our system will work with that device. Using our system with regular silicon probes, the headstages are attached to the animals during the recording session only. A counterbalanced pulley system makes sure that the animal is not carrying the extra weight of the headstage during recordings. In the revised manuscript, we provide quantitative data that compares the running speed of animals with various drives.

Dual probe implantations are possible with our system, as illustrated in Figure 5.

6. It is unclear how the headstage is affixed in either the rat or mouse assembly.

The headstage is attached to the Omnetics connector sitting on the PCB of the probe. A male header pin is attached to the probe PCB using cyanoacrylate glue and dental cement. During surgery, this male header pin is soldered to the cap system. There is no further need to support the headstage.

7. For probe recovery, it's important to note that distilled water will not be recommended for all probes. For example, neuropixels have very clear restrictions on what you should use with them. I would advise the reader accordingly. Tergazyme may be useful here as well.

The Reviewer is right that some probes might not be suitable for our cleaning procedure. In the revised manuscript, we clearly specified which probe types can be cleaned in the described way. We have also added a separate recommendation for cleaning Neuropixels 1.0 probes based on Luo et al. (2020) work.

8. How is grounding handled in these devices? There are multiple mentions of a skull screw used to affix the protective assembly, but in most designs, a skull screw is there to serve as a reference and/or grounding. (Sidenote: It is not clear to me why a ground screw is so bad for the animal, as is emphasized multiple times in the manuscript. We are also putting a large open whole in the skull…) Is there a ground wire in these devices? Similarly, is the copper mesh inside electrically connected to the probe, or is it kept isolated?

We either implant a 100-µm stainless-steel wire or a skull screw with an attached insulated wire (California Fine Wire, CA, USA) above the cerebellum to serve as ground for our recordings. After the silicon probe is implanted, this ground wire is soldered to the copper mesh or copper tape (mouse and rat cap, respectively). The probe’s ground and reference wires are also soldered to the copper mesh or copper tape. For the mesh to function as a Faraday cage, it must be grounded. Traditionally, the reference and ground electrodes were isolated from each other, but since we have been using Intan-based recording devices, we have not observed any difference in the quality of our recordings after shorting the reference and ground wires. The Intan headstages are shipped with shorted reference and ground (there is a zero-ohm jumper at R0 location on the printed circuit board of all headstages).

In the manuscript we refer to support skull screws as additional screws and not to the ground or reference screws. It has been common practice to use multiple support screws in rats and mice to attach the headgear to the skull. For instance, Vandecasteele et al., (2012) recommends using at least 4 support screws in rats. Recently, we eliminated this step and replaced the support skull screws with a surface bond material (Metabond). In our current protocol, we apply one or two layers of Metabond instead of multiple skull support screws, making the surgical procedure less invasive. We have also found that the animals recover faster. The bond is stable across the animal’s lifetime, and we have not observed any headgear loss with properly implanted animals.

9. Line 476 in methods is very unclear and mentions a plastic microdrive: "The probes ere mounted on a plastic recoverable microdrive to allow precise vertical movement after implantation (github.com/YoonGroupUmich/Microdrive) and implanted by attaching the base of the micro-drives to the skull with dental cement." Is this the same microdrive mentioned in the main manuscript? In general, it is unclear how the "Additional implantation information" relates to the main methods and seems that this information should be integrated into the other methods sections.

Please see our detailed answer to Comment 4 from Reviewer #1. Throughout the development of the metal microdrive and cap systems, we have gone through several iterations of the 3D printed plastic microdrive. This section in the manuscript was revised to better explain this history.

10. There is no mention of where to download the design files.

We have added links in the method section to our GitHub repository containing instructions and stl files: https://github.com/buzsakilab/3d_print_designs

We will create a separate zip file with all design files with the publication.

11. Data is provided for only 2 animals (one mouse, one rat) and 2 sessions – could more data be made available?

All our data will be made available after our manuscript is accepted for publication.

Pending on acceptance, we will work with the editors of ELife to link our data to this manuscript as well. Our lab has a long-lasting history of sharing our neuroscience data immediately after acceptance.

12. Code is provided for sorting (Kilosort Wrapper and phy plugin).

We used KiloSort together with KilosortWrapper:

https://github.com/petersenpeter/KilosortWrapper

Phy1 and phy2 plugins: https://github.com/petersenpeter/phy1-plugins

https://github.com/petersenpeter/phy2-plugins

[Editors' note: further revisions were suggested prior to acceptance, as described below.]

Reviewer #2 (Recommendations for the authors):

I would like to encourage the authors to incorporate into the final version of the paper all the relevant technical details from the rebuttal. For example, in their response the authors mention using a pulley to counterbalance the headstage (yet this seems not to be mentioned in the manuscript), and similarly with preferentially using large (>35gr) mice.

We have added this information to the manuscript.

Reviewer #3 (Recommendations for the authors):

There are several points that should be addressed within the revised manuscript:

The new Figure 5 illustrates the use of two different recording devices in mice. Figure 5A demonstrates the units recorded over time on each shank as the probe is lowered each day. However, it is unclear to me how Figure 5B relates to 5A – in 5B it looks as if all four shanks are at the same depth, whereas in 5A they are at different depths. Ultimately, it seems like some sort of integration representation of these two panels where viewers can appreciate the location of single units on each shank over days would be the most useful. On this same figure, the axis labels for 5F and 5I are a bit misleading, because this is not about the noise level of the recording, but about the waveform. I'd suggest changing the figure and the wording in the text to "Waveform relative noise level" so that readers do not confuse this with overall signal to noise in the recording. The axis on 5G could be readjusted so that readers can appreciate the data. In the figure caption, I'd suggest spelling out "ACGs" for readers unaccustomed to that shorthand. Presumably these ACGs are from the unit in the box? If so, I'd clarify in the caption.

We apologize for this confusion. Indeed, Figure 5A captures the single unit stability visually with individual shanks plotted above each other and panel B shows the correct horizontal layout and captures the distribution across days for units across all shanks. We have altered the figure legend to better describe this and switched the subpanels in B. We have added “Waveform relative noise level” to the appropriate labels. We also updated the figure caption for J-L. It reads now: “J-L Neuropixels probe recording, where the same putative interneuron was tracked across four days. J Average waveforms (bandpass filtered 300-10000 Hz) of a putative interneuron recorded on 16 channels across 4 days (left). The average waveforms recorded at the site with the largest amplitude waveform is highlighted on the right (waveforms are color-coded by recording day). Autocorrelation histograms (K) and spike amplitudes (L; from Kilosort) for the same single unit, color-coded by recording day.”

Related to Figure 5, the corresponding text says, "suggesting either that the distance between the electrode sites and neuron bodies decreased or that large size neurons were recorded," but it is important to note that the probe was being moved into different brain areas over these days of recording. The text should be modified to clarify this – the clear difference from day 5 to the other days is almost definitely explained by the movement to deeper brain structures. This paragraph should also note the clear decrease in the # of units with the reimplanted probe, as well as the clear increase in the noise level in the reimplanted probe.

In response to the Reviewer’s suggestion, we added this information to the revised manuscript. We would also like to point out that despite a successful probe recovery, the recording quality can deteriorate over time reducing the number of high-quality single unit clusters (e.g., increased impedance of recording sites and decreased signal-to-noise over time).

The authors have also added behavioral data, shedding light on the ability of animals to move with this headgear, however some clarity around these behavioral findings is needed. On line 219 it reads, "The 3D printed head cap system is comparable in weight to manually built headmounts" which is an abrupt and unclear transition to the paragraph about the impact of the headgear on behavior – reading between the lines, I think the authors are saying, "… however, we wanted to verify still that our headgear would not impede the animal's behavior." It is also unclear from this paragraph how this behavior was measured. The methods section describes mice and rats running on a track – were they headfixed? It is also unclear how the water reward is relevant here – did animals need to collect a water reward in order for the track to continue moving? Also, the mice with 3D printed headgear ran on a different track than the mice with manually printed headgear – some mention of this in the main text is warranted for transparency in interpreting the behavioral results shown. The wording in the methods is also unclear – is the track circular or a figure eight? Finally, contextualizing the results found here in terms of typical speeds on such a treadmill (e.g., in the discussion, see the point below) would be very useful to readers.

The Reviewer is correct that we wanted to test whether our new headgear system would interfere with the behavior of freely moving mice and rats. We added the suggested sentence to the revised manuscript. All animals were freely moving, not head fixed. Rats and mice were water deprived and had to collect water as reward. The linear mazes had ‘reward areas’ on each end where reward was delivered via an automatic infrared-beam triggered system. Animals only received water reward for trials in which they travelled from one reward site to the other. We clarified the maze design in the manuscript: ”120-cm diameter circular track with a diagonal path allowing the animals to run in a figure-eight pattern”, its similar to a classical figure-eight maze, yet return arms are altered to a continuous circle, so that mice make fewer sharp turns. The animals are water deprived and earn reward when performing the alternation task correctly. The water reward was given at the start of the central path. We added this information to the Methods section.

In this same paragraph, the authors state they found a "small significant difference," but this wording is misleading. A statistical test result is or is not significant, and the authors should remove the word small. Readers can determine whether or not the absolute value of the p-value is informative. There is a typo in this section also: "We also performed the same test on mice subjects and found a small significant difference between the median running speed of the two *rat* groups (KS-test; p = 0.045) but no significant different between the 95 percentile speeds (KS-test; p = 0.24)." I would also recommend that the authors write out the full name of the KS test, at least on the first mention.

We agree with the Reviewer that a statistical test result is either significant or not. We have removed “small” from the revised manuscript. We also spell out Kolmogorov-Smirnov test (KS-test) when it is first mentioned. The typo was also corrected.

Line 139 says, "High-quality electrophysiological signals can be collected from freely moving mice for weeks and months (Figure 2E and F)" however it is unclear from this figure whether this data was indeed recorded over weeks and months. If this is a claim the authors cannot substantiate with data, it should be removed. Figure 5 does indeed show "weeks" with the headgear (showing data out to day 16 for the 4-shank probe and 17 for neuropixels), so it seems like an applicable figure to reference here, without the mention of "months." Similar changes should be made for the reference to the rat recordings and Figure 3 – it's unclear based on the figure caption how long after implant these extracellular traces were obtained.

We have removed the “months” from the revised manuscript and added the requested information (post-op day 18) to the caption of Figure 3.

Finally, although these are significant additions to the main portion of the text, there is no mention of either the single-unit figure or the behavioral results in the discussion. A contextualization of these data in terms of the headgear's usability as well as a comparison of these findings to other recoverable drives would be useful and warrant discussion.

In response to the Reviewer’s comment, we included a new paragraph in the Discussion.

https://doi.org/10.7554/eLife.65859.sa2

Article and author information

Author details

  1. Mihály Vöröslakos

    Neuroscience Institute, New York University, New York, United States
    Contribution
    Conceptualization, Software, Investigation, Methodology, Writing - original draft
    Contributed equally with
    Peter C Petersen and Balázs Vöröslakos
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-1022-1355
  2. Peter C Petersen

    Neuroscience Institute, New York University, New York, United States
    Contribution
    Conceptualization, Software, Funding acquisition, Investigation, Methodology, Writing - original draft
    Contributed equally with
    Mihály Vöröslakos and Balázs Vöröslakos
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-2092-4791
  3. Balázs Vöröslakos

    Budapest University of Technology and Economics, Faculty of Mechanical Engineering, Budapest, Hungary
    Contribution
    Conceptualization, Methodology
    Contributed equally with
    Mihály Vöröslakos and Peter C Petersen
    Competing interests
    No competing interests declared
  4. György Buzsáki

    1. Neuroscience Institute, New York University, New York, United States
    2. Department of Neurology, Langone Medical Center, New York University, New York, United States
    Contribution
    Conceptualization, Supervision, Funding acquisition, Writing - original draft
    For correspondence
    gyorgy.buzsaki@nyulangone.org
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-3100-4800

Funding

National Institutes of Health (U19 NS107616)

  • György Buzsáki

National Institutes of Health (U19 NS104590)

  • György Buzsáki

National Institutes of Health (R01 MH122391)

  • György Buzsáki

Lundbeckfonden (R271-2017-1687)

  • Peter C Petersen

Danish Council for Independent Research, Medical Sciences (DFF-5053-00279)

  • Peter C Petersen

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank Manuel Valero, Antonio Fernandez Ruiz, and Kathryn McClain for useful comments on the manuscript. We also thank Viktor Varga, and Kathryn McClain for behavioral data and Thomas Hainmueller for intraoperative photographs. Supported by U19 NS107616, U19 NS104590, R01 MH122391, and The Lundbeck Foundation.

Ethics

Animal experimentation: All experiments were approved by the Institutional Animal Care and Use Committee at New York University Medical Center (protocol number: IA15-01466).

Senior and Reviewing Editor

  1. Laura L Colgin, University of Texas at Austin, United States

Reviewers

  1. Liset M de la Prida, Instituto Cajal, Spain
  2. Ashley L Juavinett, University of California, San Diego, United States

Publication history

  1. Received: December 16, 2020
  2. Accepted: May 18, 2021
  3. Accepted Manuscript published: May 19, 2021 (version 1)
  4. Version of Record published: June 4, 2021 (version 2)

Copyright

© 2021, Vöröslakos et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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