The video discusses a surgical procedure for implanting bone screws in a mouse's skull. It covers preparation steps, locating screw placement, scoring the skull, and applying liquid cyanoacrylate to secure the screws while avoiding damage to the underlying dura, ensuring the success of the micro drive implant.

surgical implantation of the micro-drive on the mouse skull

Micro-Drive Implant for Recording Daily Neural Activity in Juvenile Mice

The brain undergoes large-scale changes during the critical developmental windows of childhood and adolescence1,2,3. Many neurologic and psychiatric diseases, including autism and schizophrenia, first manifest behaviorally and biologically during this period of juvenile and adolescent brain development4,5,6. While much is known regarding the cellular, synaptic, and genetic changes that occur across early development, comparatively little is known regarding how circuit- or network-level processes change throughout this time window. Importantly, circuit-level brain function, which ultimately underlies complex behaviors, memory, and cognition, is a non-predictable, emergent property of cellular and synaptic function7,8,9,10. Thus, to fully understand network-level brain function, it is necessary to directly study neural activity at the level of an intact neural circuit. In addition, to identify how brain activity is altered throughout the progression of neuropsychiatric disorders, it is critical to examine network activity in a valid disease model during the specific temporal window when the behavioral phenotypes of the disease manifest and to track the observed changes as they persist into adulthood.
One of the most common and powerful scientific model organisms is the mouse, with large numbers of unique genetic strains that model neurodevelopmental disorders with age-dependent onset of the behavioral and/or mnemonic phenotypes11,12,13,14,15,16,17,18,19,20,21. While it is challenging to correlate precise developmental time points between the brains of humans and mice, morphological and behavioral comparisons indicate that p20-p21 mice represent the human ages of 2-3 years old, and p25-p35 mice represent the human ages of 11-14 years old, with mice likely reaching the developmental equivalent of a human 20 year old adult by p603,22. Thus, to better understand how the juvenile brain develops and to identify how the neural networks of the brain become dysfunctional in diseases like autism or schizophrenia, it would be ideal to directly monitor brain activity in vivo in mice across the ages of 20 days to 60 days old.
However, a fundamental challenge in monitoring brain activity across early development in mice is the small size and relative weakness of juvenile mice. The chronic implantation of electrodes, which is necessary for longitudinal studies of brain development, typically requires large, bulky housing to protect the fine electrode wires and interface boards23,24, and the implants must be firmly attached to the mouse skull, which is thinner and less rigid in young mice due to reduced ossification. Thus, virtually all studies of in vivo rodent physiology have been performed in adult subjects due to their relative size, strength, and skull thickness. To date, most studies exploring in vivo juvenile rodent brain physiology have been performed in wild-type juvenile rats, which necessarily limits the ability to experimentally monitor juvenile brain function in a freely behaving model of a human disorder25,26,27,28,29,30.
This manuscript describes novel implant housing, a surgical implantation procedure, and a post-surgery recovery strategy to chronically study the long-term (up to 4 or more weeks) in vivo brain function of juvenile mice across a developmentally critical time window (p20 to p60 and beyond). The implantation procedure allows for the reliable, permanent affixation of the electrodes to the skulls of juvenile mice. Furthermore, the micro-drive design is lightweight, as this micro-drive weighs ~4-6 g when fully assembled, and due to the minimal counterbalancing required to offset the weight of the implant, it does not impact the behavioral performance of juvenile mice during typical behavioral paradigms.

Author Spotlight: A Lightweight Drive Implant for Chronic Tetrode Recordings in Juvenile Mice  

A Lightweight Drive Implant for Chronic Tetrode Recordings in Juvenile Mice

We're interested in understanding how the brain operates at a network level. This method is our attempt to explore the development of brain networks to identify developmental alterations in age dependent diseases like autism, schizophrenia or bipolar disorder. The silicon probe technology provides a simpler, more consistent method to record network activity in vivo.But despite this, chronic tetra recordings do offer some advantages over silicon probes, such as simultaneously recording over a broader spatial distribution of brain regions. Chronic recordings pose a unique challenge to in vivo electrophysiology due to several factors, including gliosis at the recording sites, movement of the recording site over time or failure of the attachment method. Our recent work demonstrated that the sweeps encoded by the theta oscillation of the hippocampus during active movement of the animal every hundred milliseconds or so iteratively cycle forward, prospectively evaluating possible future states and backwards retrospectively evaluating prior actions.Performing these in vivo recordings in juvenile mice poses several engineering challenges due to the small size of the mice, their relative weakness and the lack of development in their skull. Our methodology overcomes these limitations and allows us to chronically record network level activity daily in the developing mouse brain. Besides recording chronically in juvenile mice, our method allows us to record from up to 16 distinct bilateral brain regions irrespective of the spatial relationship of those regions.These developments will allow us to identify how networks establish functional communication across developments, both in the healthy brain and in mouse models of neurodevelopmental disorders, like autism spectrum disorders.

Watch this Scientific Journal Video about Surgical Implantation of the Micro-Drive on the Mouse Skull at JoVE.com

Surgical Implantation of the Micro-Drive on the Mouse Skull  

After preparing the mouse for surgery and exposing the skull, place the bone screws in the extreme lateral, rostral or caudal portions of the skull where the bone is thickest and the bone screws are sufficiently far away from the micro drive implant. Use a scalpel blade or drill bit to score the skull near the bone screw hole locations, providing a rough surface for the liquid cyanoacrylate to bind. Then thread each bone screw into place, careful not to pierce the underlying dura.Using a sterile 30-gauge needle, apply liquid cyanoacrylate around each bone screw, thickening the skull where the bone screws have been attached. Ensure that no cyanoacrylate enters the exposed dura above the recording sites.

surgical-implantation-of-the-micro-drive-on-the-mouse-skull

Research

JoVE Journal

Neuroscience

479 Views.  UT Southwestern Medical Center. After preparing the mouse for surgery and exposing the skull, place the bone screws in the extreme lateral, rostral or caudal portions of the skull where the bone is thickest and the bone screws are sufficiently far away from the micro drive implant. Use a scalpel blade or drill bit to score the skull near the bone screw hole locations, providing a rough surface for the liquid cyanoacrylate to bind. Then thread each bone screw into place, careful not to pierce the underlying dura.Using...

Video: Surgical Implantation of the Micro-Drive on the Mouse Skull  

a lightweight drive implant for chronic tetrode recordings in juvenile mice

Here, we describe a micro-drive design, surgical implantation procedure, and post-surgery recovery strategy that allow for chronic field and single-unit recordings from multiple brain regions simultaneously in juvenile and adolescent mice across a critical developmental window from postnatal day 20 (p20) to postnatal day 60 (p60) and...

recovery after micro-drive implantation

Recovery after Micro-Drive Implantation  

implantation of a head bar on top of the mouse skull

Implantation of a Head Bar on Top of the Mouse Skull  

micro-drive array for chronic in vivo recording: drive fabrication

Micro-drive Array for Chronic in vivo Recording: Drive Fabrication

In this protocol we demonstrate how to fabricate a micro-drive array for chronic electrophysiological recordings in...

Micro-drive Array for Chronic in vivo Recording: Drive Fabrication

calvarial window generation in mouse model: a surgical procedure to create window in the top part of the mouse skull

In this video, we describe the surgical procedure to create a calvarial window for accessing the brain of a mouse...

Calvarial Window Generation in Mouse Model: A Surgical Procedure to Create Window in the Top Part of the Mouse Skull

After confirming a lack of response to pedal reflex, place the anesthetized mouse onto the frame, and use the blunt end of a cotton-tip applicator to position the teeth in the trough of the...

micro-drive system assembly and electrode array implantation for multichannel recording in mice

Micro-Drive System Assembly and Electrode Array Implantation for Multichannel Recording in Mice  

visualizing the brain and craniotomized skull for virtual surgical planning

Visualizing the Brain and Craniotomized Skull for Virtual Surgical Planning

micro-drive design and construction

Micro-Drive Design and Construction  

construction of the electrode micro-drive and polytrode plating of electrodes for neurophysiology recordings

Construction of the Electrode Micro-Drive and Polytrode Plating of Electrodes for Neurophysiology Recordings

electrophysiology of laminar cortical activity in the common marmoset

Author Spotlight: Optimizing Micro-Drive Systems for Precise Electrode Positioning in Marmoset

Custom-built micro-drives enable the sub-millimeter targeting of cortical recording sites with linear silicon...

Development of an X-Y Electrode Stage for Precise Neural Recording in Marmosets

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quantification of visual feature selectivity of the optokinetic reflex in mice

Author Spotlight: An Accurate and Quantitative Approach to Study Visual Feature Selectivity of the Optokinetic Reflex in Mice 

Here, we describe a standard protocol for quantifying the optokinetic reflex. It combines virtual drum stimulation and video-oculography, and thus allows precise evaluation of the feature selectivity of the behavior and its adaptive plasticity.

Evaluating Optokinetic Reflex Responses in Mice: A New High-Precision Protocol

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cranial ring assembly installation in mouse model: a surgical procedure to implant visualization window assembly on murine skull for high-resolution skull bone marrow imaging

Expose the skull with tweezers. Then, use scissors and tweezers to carefully remove the periosteum. Use a sterile cotton swab soaked in physiological saline to remove all the periosteum and any debris or hair which could alter the imaging. Rinse the skull with saline to remove any blood traces, and rapidly dry the bone with a cotton...

Cranial Ring Assembly Installation in Mouse Model: A Surgical Procedure to Implant Visualization Window Assembly on Murine Skull for High-resolution Skull Bone Marrow Imaging

In this video, we describe a surgical procedure to implant a cranial ring assembly on the exposed skull bone and further immobilize the mouse&#8217;s head for high-resolution brain imaging for a long...

micro-drive array for chronic in vivo recording: tetrode assembly

Micro-drive Array for Chronic in vivo Recording: Tetrode Assembly

In this protocol we demonstrate how to fabricate and condition tetrodes for use with a micro-drive array, which was designed for chronic electrophysiological recordings in rats.  In addition, we illustrate the final stages of micro-drive array construction, which includes installing ground wires and a protective...

Micro-drive Array for Chronic in vivo Recording: Tetrode Assembly

in vivo imaging of cerebrospinal fluid transport through the intact mouse skull using fluorescence macroscopy

In Vivo Imaging of Cerebrospinal Fluid Transport through the Intact Mouse Skull using Fluorescence Macroscopy

Transcranial optical imaging allows wide-field imaging of cerebrospinal fluid transport in the cortex of live mice through an intact...

a neural implant design toolbox for nonhuman primates

Author Spotlight: Streamlined Brain and Skull Modeling for Enhanced Neurosurgical Planning in NHP Research

This paper outlines automated processes for nonhuman primate neurosurgical planning based on magnetic resonance imaging (MRI) scans. These techniques use procedural steps in programming and design platforms to support customized implant design for NHPs. The validity of each component can then be confirmed using three-dimensional (3D) printed life-size anatomical...

Automated 3D Brain and Skull Modeling from MRI for NHP Neurosurgical Planning

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continuous drug infusion system in mouse model: a surgical procedure to implant micro-osmotic pump infusion system in the mouse brain for continuous drug delivery

In this video, we demonstrate the assembly of a micro-osmotic pump brain infusion system followed by its surgical implantation in the mouse skull. This system enables continuous delivery of potential drugs and facilitates the study of their effects on the brain and brain...

Continuous Drug Infusion System in Mouse Model: A Surgical Procedure to Implant Micro-osmotic Pump Infusion System in the Mouse Brain for Continuous Drug Delivery

a polished and reinforced thinned-skull window for long-term imaging of the mouse brain

A Polished and Reinforced Thinned-skull Window for Long-term Imaging of the Mouse Brain

We present a method to form an imaging window in the mouse skull that spans millimeters and is stable for months without inflammation of the brain. This method is well suited for longitudinal studies of blood flow, cellular dynamics, and cell/vascular structure using two-photon...

In vivo electrophysiology provides unparalleled insight into the sub-second-level circuit dynamics of the intact brain and represents a method of particular importance for studying mouse models of human neuropsychiatric disorders. However, such methods often require large cranial implants, which cannot be used in mice at early developmental time points. As such, virtually no studies of in vivo physiology have been performed in freely behaving infant or juvenile mice, despite the fact that a better understanding of neurological development in this critical window would likely provide unique insights into age-dependent developmental disorders such as autism or schizophrenia. Here, a micro-drive design, surgical implantation procedure, and post-surgery recovery strategy are described that allow for chronic field and single-unit recordings from multiple brain regions simultaneously in mice as they age from postnatal day 20 (p20) to postnatal day 60 (p60) and beyond, a time window roughly corresponding to the human ages of 2 years old through to adulthood. The number of recording electrodes and final recording sites can be easily modified and expanded, thus allowing flexible experimental control of the in vivo monitoring of behavior- or disease-relevant brain regions across development.

Here, we describe a micro-drive design, surgical implantation procedure, and post-surgery recovery strategy that allow for chronic field and single-unit recordings from multiple brain regions simultaneously in juvenile and adolescent mice across a critical developmental window from postnatal day 20 (p20) to postnatal day 60 (p60) and beyond.

In vivo electrophysiology provides unparalleled insight into the sub-second-level circuit dynamics of the intact brain and represents a method of ...

After digitally designing and printing the micro drive, attach two screw attachments to each tetrode advancing screw one above and one below the ridge. Hold the screw attachments together with gel cyanoacrylate. Ensure that the screw attachments do not move in the longitudinal axis of the screw, but can rotate freely with minimal resistance.Extend each polyamide section through the output holes on the micro drive cannula by a few millimeters. Secure the polyamide to the cannula using a clean 30 gauge needle in small amounts of liquid cyanoacrylate, being careful not to let it enter the inside of the polyamide tube. Retrieve the large polyamide tubes from the top of the micro drive cannula and pass them through the large polyamide holes in the micro drive body.Gently push the micro drive cannula and micro drive body together until they are adjacent and the cannula body attachment tabs interlock. Secure the micro drive body and the micro drive cannula with cyanoacrylate. Using a new sharp razor blade, severe the large polyamide tube ends extruding from the bottom of the cannula output holes, ensuring the cut is precisely at the cannula's base, making the tubes and cannula bottom flush.With sharp scissors, cut the large polyamide tubing just above the edge of the inner rim of the drive body at a 45 degree angle. Cut small polyamide tubing into four centimeter long sections. Pass the small polyamide sections through the large polyamide tubing already mounted in the micro drive and ensure that excess small polyamide tubing protrudes from the top and bottom.Secure the small polyamide tubes to the screw attachments with cyanoacrylate. Avoid letting any cyanoacrylate enter the large or small polyamide tubes. Using a new sharp razor blade, sever the small polyamide tube ends extruding from the bottom of the cannula holes.Ensure the cut is at the cannula's base and clean with nothing blocking the polyamide tube hole. Then use sharp scissors to cut the top of the small polyamide. a few millimeters above the top of the screw attachment at a 45 degree angle, ensuring the cut is clean.Using ceramic or rubber tipped forceps, carefully pass a tetrode through one of the small polyamide tubes, leaving approximately two centimeters protruding from the top of the small polyamide tube. Secure the tetrode to the top of the small polyamide tube with liquid cyanoacrylate. Retract the screw until it is near the top of the drive.Grab the tetrode wire protruding from the bottom of the drive and gently kink it at the point where it emerges from the cannula. Fully advance the screw into the drive. Using sharp scissors, cut the tetrode wire just above the kink.Then inspect it under a microscope to ensure the cut is clean and the metal of all four tetrodes is exposed. Finally, connect each electrode of each tetrode to the appropriate port on the electronic interface board. Tightly wind one bone screw with a thin, highly conductive wire that will be ground and attached to the electronic interface board.

To begin, set up the counterbalance system by connecting a 0.75-inch-diameter PVC pipe. Insert one arm of the system through the holes in the cage lid. Place the second arm on top of the cage lid, and extend the third arm above and beyond the cage.Finish by capping the top-most arm. After implanting the microdrive on the mouse goal, attach the microdrive to the counterbalance system using a counterbalance weight equal to the weight of the microdrive and bone screws. Secure a strong thread or fishing line from the electronic interface board to the counterbalance weight which hangs over the top-most arm.Finally, ensure the counterbalance is firmly connected to the microdrive electronic interface board and that the mouse has full access to the cage.