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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
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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
215
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
291
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
313
314
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References 315
Allen, TA, McNaughton BL, Su, M, and Allen, LM (2017) US Patent 9,707,049 Stereotactic device for 316
implantation of permanent implants into a rodent brain. 317
Allen, TA, Draper, AD, and Mattfeld, AT (2019) US Patent 15,251,722 Stereotactic brain implant system 318
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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
368
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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
390
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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
413
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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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