RatHat: A self-targeting printable brain implant system · RatHat: A self-targeting printable brain implant system Abbreviated Title: SELF-TARGETING BRAIN IMPLANT SYSTEM Leila M

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RatHat: A self-targeting printable brain implant system 3

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Abbreviated Title: SELF-TARGETING BRAIN IMPLANT SYSTEM 6

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Leila M. Allen*1, Maanasa Jayachandran*1, Tatiana D. Viena1, Meifung Su1, Timothy A. Allen1,2 9

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*Contributed equally to this work 11

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13 1Cognitive Neuroscience Program, Department of Psychology, Florida International University, Miami, FL, 33199 14

2Department of Environmental Health Sciences, Robert Stempel College of Public Health, Florida International 15

University, Miami, FL, 33199 16

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Author Contributions: LMA, MJ, MS, BLM, and TAA designed research; LMA, MJ, MS, and TDV performed 18

research; LMA, MJ, TDV and TAA analyzed data; LMA, MJ, and TAA wrote the paper. 19

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Keywords: stereotaxic, cannula, optrodes, electrodes, tetrodes, surgery 21

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Corresponding Author: 23

Timothy A. Allen, PhD 24

Department of Psychology 25

Florida International University 26

11200 SW 8th Street 27

Miami, FL, 33199 28

email: [email protected] 29

Website: http://allenlab.fiu.edu/ 30

Twitter: @AllenNeuroLab 31

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Number of figures:5 33

Number of tables: 0 34

Number of Multimedia: 0 35

Number of words for Abstract: 244 36

Number of words for Significance Statement: 88 37

Number of words for Introduction: 500 38

Number of words for Methods: 1,419 39

Number of words for Results: 1,096 40

Number of words for Discussion: 267 41

Number of words in Abstract, Sig Statement, Intro, Methods, Results, & Discussion: 3,623 42

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Acknowledgements: We thank Jason Hays for helping with figures, and Chrystle and Catherine Cu (CocoFloss) 44

for donating UV-light curing dental cement. 45

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Conflict of Interest: The authors declare no competing financial interests. 47

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Funding Sources: This work was supported, in part, by R01 MH113626 to TAA and generous funding from the 49

Feinberg Foundation. 50

.CC-BY-ND 4.0 International licenseunder anot certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available

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SELF-TARGETING BRAIN IMPLANT SYSTEM

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Abstract 51

There has not been a major change in how neuroscientists approach stereotaxic methods in decades. 52

Here we present a new stereotaxic method that improves on traditional approaches by reducing costs, 53

training, surgical time, and aiding repeatability. The RatHat brain implantation system is a 3D printable 54

stereotaxic device for rats that is fabricated prior to surgery and fits to the shape of the skull. RatHat 55

builds are directly implanted into the brain without the need for head-leveling or coordinate-mapping 56

during surgery. The RatHat system can be used in conjunction with the traditional u-frame stereotaxic 57

device, but does not require the use of a micromanipulator for successful implantations. Each RatHat 58

system contains several primary components including the implant for mounting intracranial 59

components, the surgical stencil for targeting drill sites, and the protective cap for impacts and debris. 60

Each component serves a unique function and can be used together or separately. We demonstrate 61

the feasibility of the RatHat system in four different proof-of-principle experiments: 1) a 3-pole cannula 62

apparatus, 2) an optrode-electrode assembly, 3) a fixed-electrode array, and 4) a tetrode hyperdrive. 63

Implants were successful, durable, and long-lasting (up to 9 months). RatHat print files are easily 64

created, can be modified in CAD software for a variety of applications, and are easily shared, 65

contributing to open science goals and replications. The RatHat system has been adapted to multiple 66

experimental paradigms in our lab and should be a useful new way to conduct stereotaxic implant 67

surgeries in rodents. 68



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Impact Statement 69

We demonstrate a new approach to rodent stereotaxic surgery. Rodent neurosurgery is a complex skill 70

that requires expensive equipment for head stabilization and micromanipulators for localization. The 71

RatHat is a 3D printable brain implant system that reduces costs and time using pre-mapped and 72

printed surgical files. A surgical stencil allows for quick placement of drill holes, and a RatHat places 73

components in the brain using atlas coordinates. The RatHat system is an easily shared resource 74

facilitating open science goals for simple replications and archiving of specific experimental 75

applications. 76



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Introduction 77

Rodent neurosurgery is challenging to master, especially for implant surgeries involving multiple target 78

sites. Long surgeries cause surgeon fatigue and distress in animals, which can affect recovery time and 79

surgical outcomes (Ferry et al.,2014; Pritchett-Corning et al., 2011; Hoogstraten-Miller and Brown, 80

2008; Fox, et al., 2015). With an increased emphasis on circuit analysis that includes multiple brain 81

targets (e.g. DREADDs and optogenetics), opportunities for positioning errors are increased 82

(Jorgenson et al., 2015; Bassett & Sporns, 2017; Jayachandran et al., 2019). 83

Typically, brain implants are placed using a u-framed stereotaxic apparatus in which the rat’s 84

head is stabilized with ear bars and a tooth bar, putting the rat into a three-dimensional atlas space 85

(Paxinos and Watson, 2013). Micromanipulators attached to the u-frame allow implants to be precisely 86

moved in xyz coordinate planes. However, this setup can introduce unrecoverable user errors that go 87

unnoticed in the early stages. For example, while surgeons are trained to level the head in the 88

anterior/posterior (A/P) plane, many fail to level in the medial/lateral (M/L) plane yielding asymmetrical 89

implants/injections that produce time-consuming and expensive confounds unnecessarily increasing 90

the number of animals needed for a study (Fomari et al., 2012; JoVE, 2019). Notably, there hasn’t been 91

a major change in how neuroscientists approach stereotaxic methods in decades. 92

As a practical issue, a standard u-frame surgical apparatus can range from $5k-$50k (or more 93

with addition of specialty add-ons), costing research labs a considerable portion of their equipment 94

budgets and presenting a bar-to-entry for less well-funded laboratories. 95

Here we introduce a customizable, fully integrated 3D-printable stereotaxic brain implant system 96

called RatHat that is freely available to academic researchers (Allen et al., 2017). The RatHat system 97

can be used in conjunction with, or replace, the u-framed stereotaxic apparatus in neurosurgical 98

methods requiring atlas-based positioning. A key feature is that the system self-aligns to atlas space 99

because it fits the skull, eliminating the need for micromanipulator measurements and head-leveling. 100

The RatHat system reduces costs, training, and surgery time. It is customizable for a variety of 101

surgical applications through modifications of the Computer Aided Design (CAD) environment prior to 102

surgery (e.g., Autodesk, Blender, etc.). RatHat files are easily shared over the internet and archived for 103



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later use with versioning (aiding new experiments and replications). Printouts are considerably less 104

expensive than similar commercial products while providing a larger range of implant possibilities. 105

Furthermore, surgeons can map out coordinates in an interactive 3D environment to visualize the 106

surgery prior to implantation, reducing demands during surgery. 107

RatHat applications have been adopted in our lab for a variety of experimental needs. Here, we 108

demonstrate the use of RatHat in four experimental applications: multi-site chronic cannula, multi-site 109

optrode-electrode combination implants, a fixed microwire microarray, and a tetrode hyperdrive with a 110

microarray insertion tip. 111

RatHat is freely available to academic researchers, achieving open science goals. Academic 112

researchers interested in receiving the 3D files can contact Dr. Timothy Allen ([email protected]). We will 113

first provide you a license to be executed by your institution, and upon completion, 3D files of the 114

implant system. 115

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Materials and Methods 117

RatHat components are printed using the 3D Systems ProJet 1200, a high resolution (56-micron xy, 30-118

micron layer thickness) 3D printer that uses micro-stereolithography (laser polymerization of resin and 119

UV light-curing), but any high-resolution 3D printer can be used. With the ProJet 1200 prints, we use 120

VisiJet FTX Green resin, a UV curable and biocompatible plastic composition commercially used in 121

castings because it is a durable with a tensile strength of 30MPa (or 4,351 PSI). After devices are 122

printed, we always ensure the holes are clear of debris or resin by thoroughly cleaning prints with 123

multiple dips in a 70% isopropyl alcohol (in diH20) solution and clearing holes by using a pressurized air 124

output hose. Non-printable components such as wires or tubing are secured to the implant device prior 125

to surgery with cyanoacrylate (Zap CA+, Super Glue Corporation, California) followed by a quick-cure 126

spray (Zip Kicker, Super Glue Corporation, California). Another advantage of the RatHat system is that 127

these components are easily assembled using build-specific 3D printable assembly bases or jigs. All 128

implants are sterilized with 70% ethanol in diH20 before surgical implantation and a gas sterilizer 129

(ethylene oxide). Autoclaving is not recommended. 130



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Components: RatHat implant, Surgical Stencil, Protective Cap, and Implant Jig. Several 131

components are common to all designs. The RatHat implant is a stable and secure housing apparatus 132

for long-term neurosurgical implants. It is secured to the skull with anchor screws and dental cement. 133

The underside of the RatHat that makes contact with the skull contains horizontal channels for dental 134

cement, designed to optimize long-term adhesion of the implant to the skull and anchor screws (up to 9 135

months in our cohort). The version information and animal/experiment ID can be included on the print 136

as well for ease of identification. 137

The surgical stencil contains all alignment and drill holes for the specific target sites needed in 138

the surgery and was designed to facilitate rapid and accurate drilling of implantation and/or infusion 139

sites to match the RatHat implant base. The surgical stencil is a transformative device for any surgeon 140

to rapidly and cleanly introduce holes or craniotomies for an implant or injection. It is easy to print and 141

uses relatively small amounts of resin, so multiple copies can be used for a single surgery in case a 142

back-up is needed. This also helps with making straight and unbiased holes if free-handed drilling is 143

preferred. 144

The protective cap safeguards other RatHat components (e.g. cannula tubes, dummies, 145

electrodes, tetrode drives, etc.) from dust, debris, and impacts. It mounts on the side-walls of the 146

RatHat implant base and is secured with a screw. The walls and the protective cap are outfitted with 147

screw-holes for alignment on all sides to accommodate left- and right-handed surgeons. The protective 148

cap can be printed with lab insignia and/or animal names for quick identification purposes. Protective 149

caps can be replaced with a reprint if damaged in any way. 150

The jig (used to build cannula implants) serves to model the brain space and allows for precise 151

placement and securing of implant components such as cannula tubes in the RatHat prior to surgery. In 152

order to prepare the RatHat cannula implant base for surgery, the cured and cleaned 3D print is placed 153

inside the jig. Next, pre-measured and cut stainless-steel tubes (27 ga, Component Supply Company, 154

Sarasota, FL) are placed into the RatHat through the corresponding holes in the jig. The depths of the 155

cannula are dictated by CAD-measured ledges printed within the jig for precise D/V depths. This 156

reduces fabrication time and more importantly, measurement errors, as hand-cut cannula tubes do not 157



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need to be precision-measured to a discrete depth, since the jig dictates depth. Once the stainless-158

steel cannulae are secured to the RatHat implant base, the device can be implanted in the brain without 159

the need for coordinate mapping during surgery. In this way, the jig replaces the dorsoventral (D/V) 160

component of a stereotax micromanipulator arm, allowing for hand implants if the surgeon feels 161

comfortable doing so. 162

Animals and General Surgical Methods. Subjects used were Long Evans rats that weighed 163

250-275g on arrival (n=21, 2 female). All rats included were used for other primary experiments. Rats 164

were individually housed in clear rectangular polycarbonate cages to ensure surgical implants were 165

protected from damage by cage-mates. Rats were maintained on a 12hr light-dark cycle (lights off at 166

10:00 am). Naïve rats were briefly handled for 3 - 5 days after initial arrival. Access to food and water 167

was unrestricted before surgery. All surgical and behavioral methods were in compliance with the 168

Florida International University (FIU) Institutional Animal Care and Use Committee (IACUC) and 169

Institutional Biosafety Committee (IBC). 170

Surgically implanting the RatHat follows basic techniques for intracranial survival surgery (refer 171

to Fig. 1 for visualization). Briefly, general anesthesia was induced (5%) and maintained by isoflurane 172

(1-2.5%) mixed with oxygen (800 ml/min). Rats were placed in a stereotaxic apparatus in the sterile 173

surgical field for stabilization with ear- and tooth-bars (although RatHat surgery can be performed 174

without this apparatus). Rats were administered glycopyrrulate (0.2 mg/ml, 0.5 mg/kg, s.c.) and 5 ml 175

Ringer’s solution with 5% dextrose (s.c., over the duration of the surgery) for hydration. Temperature 176

was monitored with a rectal thermometer and maintained within ±1C° of baseline temperature with a 177

heating pad. The skull was exposed following a midline incision or fish-eye cut. The periosteum was 178

detached from the skull using cotton-tipped applicators (Puritan Medical Products, Maine) and clamped 179

with small hemostats to expose the width of the skull up to the lateral ridges (and 2 mm beyond the 180

ridges when accessing more lateral structures) and 3-4mm length (A/P) beyond bregma and lambda. 181

Score marks were made on the skull using the scalpel blade to aid dental cement adhesion. 182

The surgical stencil was aligned to bregma and lambda using the landmark holes that are 183

surrounded by crosshairs to facilitate visualization and placement. The stencil was secured to the skull 184



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using cyanoacrylate (Zap CA+, Super Glue Corporation, California) followed by a quick-cure spray (Zip 185

Kicker, Super Glue Corporation, California). Drill holes were made in the appropriate regions according 186

to the specific RatHat build using a surgical drill (OmniDrill 3S, World Precision Instruments). Dura 187

mater was ruptured at the implant sites using a 32-gauge needle. The stencil was removed using a 188

scalpel blade or a spatula and discarded. Excess cyanoacrylate residue was scraped off the skull to 189

clear any debris that could interfere with placement of the RatHat implant base. The skull was 190

thoroughly cleaned with sterile saline or hydrogen peroxide (avoiding contact with skin and muscles) to 191

ensure successful long-term adherence of the RatHat. Titanium anchor screws were secured into 192

place. The RatHat was aligned to the drill holes and carefully lowered into place using the 193

micromanipulator arm or by hand, fitting it flush with the skull. Dental cement was applied in layers to 194

secure the base to the anchor screws and skull using a wooden applicator tip, a syringe, or a paint 195

brush (saturated first in the curing liquid and then used to pick up the dry powder, which polymerized 196

into the cement and facilitated creation of a smooth and well-anchored implant, free of jagged edges). 197

The inside of the implant was filled with dental cement to further stabilize components. Once dry, the 198

protective cap was secured onto the wall of the RatHat using a small screw. The posterior incision was 199

sutured if necessary, rats were administered an analgesic (Flunixin, 50 mg/ml, 2.5 mg/kg, s.c.), and 200

topical antibiotic ointment was applied around the surgical incision. The rat was placed in a post-201

surgical recovery incubator until awake and moving, and then returned to a clean home cage. A day 202

following surgery, rats were given an analgesic (Flunixin, 50 mg/ml, 2.5 mg/kg, s.c.) and topical 203

antibiotic ointment was applied. The protective cap was removed to check that the RatHat implant 204

components were in good condition. Rats were monitored post-operatively for a week and then 205

resumed behavioral or experimental testing. 206

Upon completion of the experiments, intracranial placements were mapped using postmortem 207

brain slices. Briefly, rats were induced under general anesthesia using isoflurane (5%) and 208

transcardially perfused with 100 ml of ice cold 0.1M PBS followed by 200 ml of 4% paraformaldehyde 209

(pH 7.4; Sigma-Aldrich, St. Louis, MO). Brains were post-fixed overnight in 4% paraformaldehyde and 210

then cryoprotected in a 30% sucrose and 0.1M PBS solution prior to sectioning (Leica CM3050S, Leica 211



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Biosystems). Three sets of immediately adjacent sections (40 μm, coronal orientation) were saved. 212

One set was mounted onto microscope slides for a cell-body specific Cresyl Violet stain for placement 213

analysis. 214

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Results 216

Experiment 1: 3-pole cannula RatHat for simultaneous implantation of multiple cannula 217

(Fig. 2). Commercially available multisite cannula assemblies from vendors such as PlasticsOne and 218

WPI are custom ordered, requiring a necessary lead-time, and very expensive. Furthermore, they only 219

accommodate up to two cannulas anchored together by a thin plastic tether, and are unable to 220

incorporate poles for angled insertions (other than perpendicular to the skull). 221

The RatHat cannula system contains multiple pre-measured cannulas assembled before 222

surgery, reducing surgical time by eliminating the need to identify coordinates with micromanipulators 223

and make insertions one-at-a-time. Here, two cannulas targeted perirhinal cortex (PER) bilaterally, and 224

one cannula targeted the nucleus reuniens of the thalamus (RE). PER is a good site to demonstrate the 225

RatHat cannula approach because it is a difficult structure to access, given its depth and laterality (A/P 226

-3.0 to -7.0; M/L ±.7.2; D/V -6.5 to -7.5; Paxinos & Watson, 2013; Burwell, 2001). The third cannula 227

targeted RE, a structure that lies directly below the superior sagittal sinus (SSS; A/P -1.08 to -3.48; M/L 228

±.08; -6.8 to –7.8 D/V). The SSS can easily rupture, prolonging surgical time and causing significant 229

damage or death. Thus, we incorporated an angled cannula pole (10°) into this RatHat design to target 230

RE and avoid SSS. This angled pole is fitted with a depth-stop, eliminating the need for D/V 231

measurements, and was inserted by hand then secured to the RatHat implant base. 232

Prior to implantation, male rats (n=13) were trained in an odor sequence memory task (from 233

Jayachandran, et al., 2019). Briefly, once rats reached criterion in the task, they underwent RatHat 234

implantation surgery. Following recovery, rats were retrained on the sequence task until they reached 235

performance criteria. This task demonstrates the durability of the RatHat, which is an ideal device for 236

experiments that require extended testing periods and involve extensive task related wear-and-tear. 237

These rats completed approximately 60 sessions after surgery, with 200-300 nose-pokes/session. 238



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Additionally, rats were given 12 infusions over several weeks to either PER (bilaterally) or thalamic RE. 239

Infusions targeted the structures of interest and resulted in distinct sequence memory disruptions that 240

relate to the functioning of those regions. RatHat implants stayed on for an average of 6 months 241

(maximally 9 months), when rats were killed for histological analysis. There were no significant 242

deviations from the targets (see Fig. 2 G/H) showing great reliability. 243

Experiment 2: RatHat design for a combination of optogenetics and microwire recordings 244

(Fig. 3). We implanted rats (n = 4; 2 females) weighing approximately 275-350g at surgery. Here, the 245

optrode targeted the junction of the thalamic RE body and thalamic RE wing (perireuniens; -2.3 A/P, -246

0.5 M/L, -7.0 D/V) and was implanted vertically at an M/L slightly lateral to the midline (a different 247

approach compared to the cannula; Viena, et al., 2019). The surgical stencil for this version was 248

designed with drill hole guides for the injection site/optrode, stainless steel wire electrode, and anchor 249

screws. After the holes were drilled, an injection of AAVr-CAG-hChR2-H134R-tdTomato (experimental 250

virus to express channelrhodopsin; Addgene cat #: 28017) or pAAVr-CAG-tdTomato (control virus; 251

Addgene cat #: 59462) was made using pulled glass pipettes (P-2000 Laser-Based Micropipette Puller, 252

Sutter Instruments) with a tip diameter between 80-100 μm driven by a motorized infusion pump (0.3-253

0.5 μL at 60 nL/min; Nanoject III, Drummund Scientific). Because the optrode has a built-in depth-stop 254

for the D/V axis, a jig was not required for this version. Similar steps for implantation were used during 255

this surgery as described above. 256

We show sample data in which optogenetic stimulation of RE in experimental rats yielded a 4Hz 257

frequency rhythm in the mPFC (strong) and dHC (weak) LFP signal, but not in controls, demonstrating 258

efficacy of the optogenetic approach (Fig 3G). Implants remained in place for 4.5 months until brain 259

analysis. Proper placement of the optrode and electrode wires was verified and consistent in all rats 260

(see Fig. 3F for representative slices depicting optrode placements). 261

Experiment 3: RatHat for implanting fixed stainless-steel wire arrays targeting prelimbic 262

(PL) and infralimbic (IL) regions of the medial prefrontal cortex (mPFC; see Fig. 4) were piloted for 263

feasibility in male rats (n=2; ~350g at surgery). The electrode array was built similar to those used in 264

other experiments (Krupa et al., 2009; Narayanan et al., 2006). The surgical stencil for this version 265



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included a craniotomy window supporting the electrode arrays bilaterally targeting PL/IL (1.7-4.0 A/P, 266

±1.8 M/L, -3.0 D/V) in addition to landmark and anchor screw holes. Once the craniotomy and drill 267

holes were made, anchor screws were inserted. Next, the RatHat implant base was secured to the 268

skull. The electrode array was then inserted into the brain, docked into place on the RatHat base, and 269

secured with dental cement. The protective wall was then secured with dental cement (filling in the 270

base up to the Electrode Interface Board). Once dry, we plugged the rat into the electrophysiological 271

recording system (Plexon, Dallas, TX) to assess neural activity. After, the protective cap was placed 272

and secured to the RatHat. Neural activity was assessed over the course of the next several months. 273

RatHat electrode arrays remained in place for approximately 4 months with good signal. We 274

successfully recorded well-isolated single-unit activity (two well-isolated units are shown in Fig. 4H). 275

Marking lesions were performed using a NanoZ for localizing electrode sites in the brain (Fig. 4G). 276

Experiment 4: 8-tetrode hyperdrive RatHat (Fig. 5). Microdrive screws and shuttles for the 277

RatHat hyperdrive were assembled and implanted similar to others (Wilson and McNaughton, 1993; 278

Gray, et al., 1995; Nguyen, et al. 2009). The tetrode array was securely encased in the protective wall 279

prior to surgery. We implanted the RatHat hyperdrive in male rats (n=2; ~350g at surgery). This stencil 280

version was the same as that used in Experiment 3. After the RatHat implant base was secured to the 281

skull, the RatHat hyperdrive was carefully placed by hand and secured onto the base with dental 282

cement. Immediately after, tetrodes were driven 1 mm and the rat was plugged into the 283

electrophysiological recording system (Plexon, Dallas, TX) to assess signal on the wires. Once 284

functionality was established, the rat was unplugged and protective cap was secured into place. 285

Tetrodes were driven 250μm/day until reaching a depth of 2.8 to 3.0 mm (staggered) with a goal of 286

recording from mPFC cells (4.7 to 2.5 A/P range, ±0.2 to ±1.6 M/L range). The hyperdrive successfully 287

isolated single-units in mPFC of freely-behaving rats, demonstrating the RatHat application (Fig. 5H). 288

Four weeks after implantation, marking lesions were performed using the a NanoZ to localize tetrode 289

sites. Cresyl Violet-stained sections were analyzed for placement of the tetrode wires (Fig. 5G). 290

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Discussion 292



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RatHat is a 3D-printed stereotaxic device that can be used for a range of applications, such as cannula 293

placements, microinfusions, optogenetics, and electrophysiological recordings. The RatHat system was 294

developed to improve surgical accuracy and precision, reduce surgery time in rodent neurosurgical 295

procedures, and contribute to open science goals. The RatHat system is freely-available to academic 296

researchers. This is a major change to current stereotaxic approaches because we replaced an 297

approach that has been used for several decades that uses micromanipulators for measurements 298

during surgery. The RatHat system saves time, money, offers reliability, and provides for surgical 299

replications. The fundamental system consists of complementary components including a RatHat 300

implant base, a surgical stencil, and a protective cap. Here we demonstrated four different RatHat 301

systems for feasibility in multiple types of neuroscience experiments. We verified the durability of these 302

implants, which remained in place for up to 9 months, in spite of movement- and impact-dense 303

behavioral tasks (e.g. Jayachandran, et al., 2019). 304

We plan to develop a RatHat for other commonly used species in neuroscience, including mice. 305

We have also developed a version for chronic implants in the domestic pig, which facilitates surgery 306

without the need for a traditional large-animal stereotaxic apparatus (HogHat; US Patent 10,251,722 307

B1, 2019). In addition, RatHat versions for other common neurosurgical applications are underway 308

including an acute implant device that allows for single injections of excitotoxins, AAVs, DREADDs, etc. 309

Again, we make the RatHat freely-available for all academic researchers to aid in their 310

experiments and contribute to open science goals. We look forward to new builds and implementations 311

from other academic research groups. 312

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dynamics in medial prefrontal cortex and the hippocampus of behaving rats. Program Number: 349

242.29. 2019 Neuroscience Meeting Planner. Chicago, IL: Society for Neuroscience, 2019. 350

Online. 351

Willson, MA, McNaughton, BL (1993) Dynamics of the hippocampal ensemble code for space Science 352

261(5124): 1055-8. 353

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Figure 1: RatHat surgical procedures 360

A) Prep: i. Prepare the skin for incision. ii. Make the incision. iii. Clean skull and expose bregma and lamda. iv. 361

Mark bregma and lamda and secure clamps to periosteum as needed. B) Drill: i. Place the stencil on the skull 362

and align it to bregma and lamda. ii. Glue the stencil to the skull using cyanoacrylate and a quick-cure spray. iii. 363

Drill holes according to the stencil. iv. Holes for skull screws and cannula (RE and PER) shown. C) Implant: i. 364

Insert skull screws (holes remain for implant sites). ii. Manual placement of the preassembled cannula RatHat 365

implant base. iii. Dental cement RatHat to the skull and insert dummies (asterisks indicate location of cannula 366

poles). iv. Place the protective cap and secure with a screw. 367

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Figure 2: 3-Pole Cannula RatHat 374

A) Full RatHat cannula assembly on the skull. B) Rat skull in different orientations. C) The Protective Cap shown 375

in different orientations. D) The RatHat Implant Base with preassembled cannula. E) The Jig is used to assemble 376

the cannula tubes into the RatHat implant prior to surgery. F) The stencil contains reference marks for bregma 377

and lamda to align to the skull; once adhered, all drill marks are properly placed for cannula access points and 378

anchor screws, saving time. The stencil is removed and discarded after drill holes are made. G) i. Sample coronal 379

slice. The asterisk indicates the infusion cannula tip location in RE. ii. Microinfusion injector tip location in the RE 380

for all rats (n=13). Numbers to the right of each section indicate distance (mm) anterior to bregma. H) i. Sample 381

coronal slice. The asterisks indicate the infusion cannula tip location in PER. ii. Microinfusion injector tip location 382

in the PER for all rats (n=13). Numbers to the right of each section indicate distance (mm) anterior to bregma. I) 383

Rats were injected with AAV-hM4Di (an inhibitory DREADD) in mPFC or a control virus, and a cannula targeted 384

RE and PER (bilaterally). Well-trained rats were infused with CNO in RE and PER (the DREADD agonist) or 385

vehicle prior to testing. i. Silencing the mPFC � RE terminals (the CNO-hM4Di group) abolished sequence 386

memory. ii. Silencing the mPFC � PER terminals (the CNO-hM4Di group) abolished sequence memory. G-I 387

were reprinted from Jayachandran et al. (2019) with permission. Abbreviations: (CNO) clozapine N-oxide, (mPFC) 388

medial prefrontal cortex, (RE) nucleus reuniens of the thalamus, (PER) perirhinal cortex. 389

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Figure 3: Optrode and SS Wire Electrode RatHat Assembly 396

A) Optrode/Electrode RatHat implant on an average sized rat skull. B) Rat skull viewed across different 397

anatomical planes. C) View of the RatHat protective cap and wall in different orientations. These items protect the 398

internal components post implantation. D) RatHat implant base (in green) preassembled with optrode and 399

electrode single wires. E) RatHat surgical stencil showing prefabricated holes that correspond to brain 400

coordinates of interest, bregma and lamda, and screw locations for rapid drilling on the skull. F) Brain sections 401

showing channelrhodopsin (ii) and viral control (ii) expressed neurons in the midline thalamus and the optrode 402

placement (asterisk) just above RE in representative cases, demonstrating the effectiveness of using the RatHat 403

system. G) Perievent spectrograms of representative mPFC and dHC LFP showing the 5 minute period in which 404

the blue LED light was administered via the optrode (see asterisk for tip location) activating ChR2 ion channels in 405

infected (i; AAVr-CAG-hChR2-H134R-tdTomato) and control (ii; pAAVr-CAG-tdTomato) rats. Also shown, 60 406

seconds before and after the stimulation block. Pulsed blue light activation (4Hz, 60 ms pulse width) of RE ChR2+ 407

neurons elicited a 4Hz frequency rhythm (see arrows) in the mPFC (strong) and dHC (weak) LFP signal. We also 408

observed comparable frequency-specific activations at 1Hz, 2Hz, and 8 Hz. This change, however, was not 409

observed in control animals (on right). Abbreviations: (mPFC) medial prefrontal cortex, (dHC) dorsal 410

hippocampus, (RE) nucleus reuniens of the thalamus, (PVA) paraventricular nucleus, (MD) medial dorsal nucleus, 411

(ChR2) channelrhodopsin. 412

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Figure 4: 16 Single Wire Fixed Electrode Array RatHat 419

A) Fixed electrode array RatHat on the skull. B) Rat skull in different orientations. C) The protective cap and wall 420

that protects the electrode array after implantation. D) The RatHat electrode array with preassembled fixed 421

stainless-steel single wires docks into the RatHat implant base. E) The RatHat implant base is anchored to the 422

skull before the RatHat electrode array is docked, ensuring the wires descend to the correct DV. F) The Stencil 423

contains reference marks to align bregma, as well as a craniotomy window and drill holes for anchor and ground 424

screws. G) Sample coronal slice of the 16-Wire SS Electrode Array RatHat. The asterisks indicate single wire tip 425

locations. H) Sample cluster plot showing two isolated mPFC units on a single channel during free-roaming 426

behavior. Abbreviations: (PL) prelimbic cortex. 427



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Figure 5: 8-Wire Tetrode Hyperdrive RatHat 432

A) The fully assembled 8-Wire Tetrode Hyperdrive RatHat on the skull. B) Rat skull in different orientations. C) 433

The protective cap and wall ensure the RatHat hyperdrive is safe from impacts and debris. D) The hyperdrive with 434

preassembled drivable tetrodes targeting regions in mPFC. E) The RatHat implant base is secured to the skull 435

and has docking poles on which the RatHat hyperdrive sits, ensuring the tetrode tips are placed right above 436

cortex. F) The stencil aligns to bregma and lambda and contains guide holes for drilling craniotomies and anchor 437

screw holes. G) Sample slice with 8-wire tetrode hyperdrive RatHat. Asterisks indicate the tetrode wire tips H) 438

Implanted tetrodes in mPFC with hM4Di expression showing functional inhibition following CNO injection 439

(1mg/kg). Abbreviations: (CNO) clozapine N-oxide, (mPFC) medial prefrontal cortex, (PL) prelimbic cortex, (Veh) 440

Vehicle. 441



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