Behavioral Training Procedures for Head-fixed Virtual Reality in Mice

All procedures involving animals were conducted in strict adherence to the protocols established by the NIEHS Animal Care and Use Committee, ensuring compliance with ethical standards and welfare guidelines. C57BL/6Tac mice, approximately 8 weeks old, were utilized for the study.

1. Surgery for head-bar implantation

1. Surgery preparation
   1. Obtain the desired quantity of mice for one's cohort, ideally housing them individually to minimize interference with the head-bar implant, although this is optional¹⁹. This study used a sample size of three male and three female mice (initially balanced, but one male was excluded early in training after failing to run on the ball)
   2. Acquire the materials specified in the Table of Materials, adjusting according to the specifics of the study design.
   3. Upon acquisition of the mice, designate individual identifiers and apply either tail tattoos or ear hole punches to ensure unambiguous identification. Establish a comprehensive log to systematically record their weights as required for the water restriction procedure.
2. Anesthesia administration
   1. Ensure that all surgical instruments are readily available, including appropriate syringes, a heating pad, metalware (such as tweezers, micro scissors, and hemostats), iodine solution, eye lubricant, and beakers for saline and hydrogen peroxide. To ensure sterile conditions, sanitize all surgical equipment and sterilize all surgical tools using an autoclave.
   2. Before proceeding with surgery, conduct precise weight measurements of the mice and activate the heating pad to 34 °C. Record all necessary information in one's institution's laboratory/surgical notebook. Verify the adequacy of oxygen and isoflurane tank levels and confirm the availability of all essential materials to facilitate uninterrupted surgical procedures.
   3. Scruff the mouse and place it into a nosecone attached to a vaporizer designed for small animals receiving 4% isoflurane and an oxygen flow rate of 3 L/min to induce anesthesia. Use a scavenger to capture any potentially harmful waste gases (recommended).
   4. Administer Petroleum ophthalmic veterinary ointment to the mouse's eyes while it is under the nosecone to prevent ocular dryness. Apply a single drop to each eye initially and reapply as necessary. Ensure constant protection of the eyes by always maintaining a layer of this lubricant with periodic checking.
   5. Prepare the surgical site (Figure 1A) on the mouse's head by shaving the area where the cranium will be affixed to the head-bar.
   6. Position the mouse's incisors within the stereotaxic apparatus beneath the nosecone, adjusting the oxygen flow rate to 1 L/min and administering 1%-2% isoflurane from the vaporizer. Ensure proper anesthesia flow by adjusting the valves and switching between the induction nosecone and the stereotaxic apparatus accordingly. Extend the mouse's hind leg and apply firm pressure to the toe. If the foot does not exhibit a reflexive withdrawal response, this indicates that the anesthesia is effective. Repeat every 15 min, along with a breathing check.
   7. Secure the mouse's head in place by attaching surgical stability bars within its ear canals, minimizing any potential head movement during surgery.
   8. Prior to making incisions or injections, sterilize the shaved surgical area on the top of the head by scrubbing it with a swab dipped in an iodine antiseptic. From this stage onward, employ sterile gloves to maintain aseptic conditions.
3. Injection administration
   1. Subcutaneously inject 0.05 mL of bupivacaine (local analgesia) with a 25G needle into the surgical incision site on the scalp.
   2. Subcutaneously inject 1 mL of saline (hydration) with a 25G needle into one side of the interscapular region.
   3. Subcutaneously inject 0.05 mL of buprenorphine (whole-body analgesia) with a 25G needle into the opposite side of the interscapular region.
4. Exposing the skull
   1. Use micro scissors to create an incision of the skin above the interfrontal and internasal sutures of the cranium, starting from just above the eyebrow ridge and extending to behind the occipital notch (Figure 1A).
   2. Use hemostats to hold down the left and right flaps of the skin, exposing the cranium.
   3. Use a dry cotton swab to remove connective tissue from the scalp between the pinned-back skin folds.
   4. Use a cotton swab wetted (but not saturated) with hydrogen peroxide to scrub the scalp, ensuring visibility of the sutures and being careful not to get hydrogen peroxide on the surrounding tissue.
   5. Repeat steps 1.4.3 and 1.4.4, 2x-3x, until both bregma and lambda are distinctly visible, and the scalp is thoroughly cleaned.
5. Surgical screw implantation
   1. Attach two screws to the skull, positioning one screw posterior to bregma and the other anterior to lambda (Figure 1B) to maximize surface area for dental adhesive and increase the stability of the head bar. Position the location of the screws at targets a specified distance from bregma. Ensure that one screw is positioned on the left and the other on the right (i.e., Anterior-Posterior (AP) +1.00, Medial-Lateral (ML) -1.00 and AP -3.00, ML +3.00), ensuring there is adequate space between the screws to accommodate the placement of the head-bar and adjust the coordinates and needed.
   2. Drill target positions, ensuring that drilling is limited to the bone of the cranium and does not penetrate the brain tissue.
   3. Using a screwdriver, screw roughly half of the screw into place. Repeat for the second screw.
6. Attaching the head-bar implant
   1. Blend dental cement and administer it to the underside of the head-bar, focusing on the concave surface, and apply along the interfrontal suture of the cranium.
   2. Position the head-bar over the interfrontal suture to facilitate bonding between the dental cement on the head-bar and that on the suture. Securely hold it in place by hand at the desired angle for approximately 5 min until it sets. Apply additional dental cement as needed. (Figure 1C-E)
7. Reattaching the skin over the head-bar
   1. Release the hemostats and utilize tweezers to reunite the two skin flaps over the dried dental cement-fixed head-bar. Use topical tissue adhesive to delicately secure the skin together by slowly adhering the left and right portions of the scalp together over the head-bar, starting from the anterior incision site and ending at the posterior incision site.
   2. Allow the topical tissue adhesive to harden to confirm the surgical area has been resealed before releasing the mouse from the surgical stability bars and nosecone.
   3. Transfer the mouse to a singly housed cage and place it on a heating pad at 37.5 °C.
   4. Carefully monitor the mouse for any signs of discomfort or breathing irregularities until it has regained consciousness. Do not leave the mouse unattended until it has regained sternal recumbency, exhibits alertness, and is ambulatory.
   5. Following surgical procedures, allow mice to undergo a 1-week rest period. Monitor the mice daily to detect and address any notable fluctuations in weight. Provide mash for the mice 3 days post-surgery to aid in recovery. To prevent interference with the head-bar, house these mice individually.

2. Fluid restriction

NOTE: Water restriction induces a state of thirst in mice, heightening their motivation for liquid rewards. However, meticulous implementation is necessary to ensure the preservation of mouse well-being²⁰.

1. At 1 week from the surgery day, establish baseline weights for mice.
2. Tape one segment of a small Petri dish (60 mm x 15 mm) concave-side down onto the cage floor, taping a smaller Petri dish (35 mm x 10 mm) concave down to the center of the flat surface of the Petri dish taped to the floor, with another small Petri dish (60 mm x 15 mm) concave up taped atop the middle dish's flat surface to serve as a water reservoir (Figure 2).
3. Ensure that the height of the upper dish prevents contamination from bedding material while allowing mice easy access to water. Add the daily water allowance volume to the reservoir using a pipette.
4. On day 1, provide mice with a dose of 15 ml water per 100 g of body mass.
5. On day 2, provide mice with a dose of 10 mL water per 100 g of body mass.
6. On day 3, provide mice with a dose of 5 mL water per 100 g of body mass. Mice should receive a minimum intake of 1 mL of water per day throughout the study, regardless of body weight.
   NOTE: Researchers may opt to administer the minimal volume uniformly across all subjects, although such adjustments should be made with careful consideration.
7. Keep a consistent dosage throughout the study duration of water allocation at 5 mL per 100 g body weight (or a uniform 1 mL water allocation if preferred)
   NOTE: Mice should be provided ad libitum access to water for 1-2 days per week when mice are not performing a VR experiment (i.e., during the weekend). This will facilitate the restoration of their natural hydration levels. In instances where mice fall below 90% of their recorded baseline weights, they should be transitioned to ad libitum water access until they reach 90% of their baseline weight. Mice falling below 80% of their recorded baseline weights should be ethically euthanized.
8. Delay administering their daily water dose at least 30 min post-behavioral assessment to mitigate potential interference with their natural thirst behaviors essential for conducting the experiment accurately.
   NOTE: Providing liquid rewards immediately after a trial may inadvertently influence the mice's performance, as they may anticipate receiving an immediate reward, potentially compromising task engagement. Therefore, delaying access to water post-trial prevents habituation to immediate reward delivery and preserves the integrity of the experimental setup.

3. System setup

1. Familiarization with equipment: For the hardware components and other considerations of the VR behavioral systems see the steps below.
   See discussion section for an examination of protocol generalizability to comparable system setups.
   1. Fully immersive virtual display or dome: This virtual display provides complete immersion for an animal within a customizable virtual environment. The motion within the virtual environment is synchronized with the movement on the spherical treadmill.
   2. Liquid reward system: The liquid reward system functions through the delivery of liquid reinforcement (water or sugar water) using a peristaltic pump, which directs the reward solution through a plastic-coated metallic tube extending to the mouse when a task is successfully executed. It contains sensors that monitor the quantity of rewards obtained by a mouse during a trial.
      1. Clean the reward tube weekly using ethyl alcohol or an alternative cleaning agent. To do this, flush 2-5 mL of the cleaning agent through the tube, similar to the delivery of liquid reward, followed by a comparable flush with an equal volume of water.
      2. When starting the experiment, determine the dispensing rate of the liquid reward from the reward tube by activating it for a specified duration and measuring the volume of dispensed liquid. This procedure allows determination of the peristaltic pump's liquid delivery rate. In this investigation, a dispensing rate of approximately 0.0083 mL/s was used.
         NOTE: Most systems offer programmable settings for the duration between behavior execution and reward release, enabling precise planning of the study protocol based on the intended reward volume per trial. The quantity used was determined to be sufficient as it allowed adequate time for the mouse to consume the reward, and its volume appeared to be motivating.
      3. Some protocols may require the mouse to lick the reward spout in order to initiate reward delivery. For the type of tasks employed here this functionality was not employed, deliver rewards contingent solely on successful execution of the desired behavior (i.e. choosing the correct arm in the y-maze). This helps to avoid failures early in training where the tendency to lick has not been sufficiently established, and the initial lick is less likely to occur. It also can allow for measurement of the reward expectancy, which is dissociable from navigational performance under some conditions¹¹.
      4. While some experiments using fluid restriction opt for the use of standard water through the reward tube, here use sugar water (10% sucrose v/v) as an additional motivational stimulus within the operant paradigm. Notably, enhanced performance across multiple experimental cohorts was observed with the introduction of sugar water.
   3. Styrofoam ball: This ball acts as a spherical treadmill. When cushioned with air from below, train the mice to run or walk comfortably on the ball. Position it atop a ball holder equipped with motion tracking sensors which collect data on distance and velocity.
   4. Head holder: Position the apparatus posterior to the mouse, ensuring visual alignment with the VR display upon attachment of the head bar to the holder. This apparatus is crucial for maintaining the mouse in a head-fixed position, thereby mitigating motion artifacts, particularly when the system is utilized alongside optical imaging or electrophysiological techniques.
   5. Air flow hardware: Configure the airflow from a compressed air source to the ball for the creation of a weightless environment conducive to mice running on the ball. This setup requires a flow regulator to ensure precise control over the air pressure applied to the ball. The ball operates efficiently within the weightless environment with minimal air supply. Therefore, during system setup, ascertain the minimum quantity of air necessary to facilitate smooth and unhindered movement of the ball within the holder. A flow between 10-20 L/min is recommended.
2. Software setup: For specific details for the operation of the system, see below.
   ​NOTE: Much like the design of video games²¹, the architecture of virtual worlds integrates key elements such as an external controller, a programmable navigable environment, and a schedule file containing a state diagram delineating dynamic functions. These components synergistically converge to craft a cohesive interactive experience for subjects engaged in research studies. The operational efficacy of the software depends on precise file organization within designated folders. This explanation will outline the main steps needed to populate pre-made templates, allowing for easy adjustments to existing files and saving them as new versions. These new versions will then form the basis of the study.
   1. Use the following three files together to configure an operational virtual landscape.
      1. XML files: This file format provides users with the capability to manipulate the photo texture of various elements such as the sky, floor, and walls. Place the files used for the imagery in the Data subfolder of the VR folder. Using these, specify the dimensions of the maze and determine the initial position of the mouse within the maze. Define 3D objects (visual cues) at certain nodes within the maze using these files. Modify these files using a text editor.
      2. XLSX files: These operate as the command files that configure all three file types (XML, XLSX, and XAML) together to form a cohesive and interactive virtual presentation. Use these files to define the experimental routines that run the VR and its accessories, such as gain sensitivity, which data is extracted, and which files are grouped together for an experiment.
      3. XAML files: The software application provides a graphical interface for the creation of experimental schedules through the utilization of flowcharts. It facilitates the definition of temporal parameters for the trial, controls for teleportation after the trial is completed, and the activation timing for digital outputs within the trial framework.
   2. Use the following applications for the acquisition of data and user control of the system while it is in operation.
      1. [application] VR: Associated with the .XML file that shows the representative landscape, open the file to preview the virtual landscape on the monitors in static mode. For dynamic interaction, open the paired configuration. XLSX file in the control application.
      2. [application] control: Associated with the .XSLX file, open this application to see the accessory devices associated with the system. One can manually extend and retract the reward tube, dispense liquid reward, and view real-time data acquisition from here.
      3. [application] schedule designer: This application provides the capability to adjust XAML files to establish a schedule for event triggering within the experiment. For example, design a customizable trigger to determine the duration for dispensing rewards and define the duration of breaks between trials for mice.
3. Example Startup: Begin by deciding what the study protocol will look like based on the adjustable components from steps 3.2.1.1-3.2.1.3. After an operant protocol is clearly defined, open one of the template experiments that came preset with the VR system by following the steps below.
   1. Open the VR application, which opens to the Data subfolder. Save the virtual landscape created as an XML file. Open this file, and the virtual landscape should appear on the VR monitors.
   2. Open the Control application and navigate to the Open folder icon in the top right of the screen. Click the Icon, which should bring up the Configs folder, where the corresponding .XLSX experimental configuration is located. Open the .XLSX file with the same name as the .XML file opened in the VR application. The defined system accessories, like the pump and motor for the extendable reward device, are now visible under the control tab within the application.
   3. Initiate the experimental trial, as the coordination between these two applications enables the creation of an interactive virtual landscape. Ultimately, this integration facilitates the monitoring of essential data, including distance in the XY plane and the collection of rewards with timestamps.
4. Data acquisition: Extract the most valuable behavioral data from the system, which are timestamped positional data and rewards. These data are saved separately as log files.
   1. Position data: To acquire this, follow the steps described below.
      1. To acquire timestamped XY positional data of the mice, first open the spreadsheet file of the maze of desired data acquisition. In table 1, put the command WriteVRAndCamInfoToFile in one of the cells below the others in column A. Now, positional data will be saved as a dated CSV file (named Log files-MM.DD.YYYY_VRandPathPos.csv) automatically into the configs folder after a trial.
      2. To export the position data after a trial, close the control application, and the data will be saved to a dated CSV file. This file will contain all the specific data for a particular day, so be careful to manually note when each subject was placed on and taken off the ball. Open the file and import it using the Unicode UTF-8 character set. Column A is labeled DateTime, right click on the A tab and click Format Cells. Go to time and click the MM/DD/YYYY HH:MM: SS option. Now, each system event will be cataloged chronologically for further data analysis.
   2. Reward data: To acquire this, follow the steps described below.
      1. Data on pump activation (reward dispensing) are automatically saved as dated log files in the system, so there is no need to put in a command like one would for the position data. To access these, go to the Log Files subfolder of the configs folder.
      2. Repeat step 3.4.1 for position data for the reward data to export the data as a spreadsheet file. Open the configs folder and select the dated reward file (named Corridor- MM.DD.YYYY or Corridor_Linear_Run- MM.DD.YYYY) when viewed in the folder. This will provide the date and time for when the mice acquired the rewards, and one can use this in further data analysis depending on the paradigm they employed.

4. Behavioral Tasks

NOTE: In accordance with established methodologies in behavioral neuroscience, the formulated tasks employ a reward-based associative learning technique. By employing immediate rewards to reinforce specific behaviors, animals are trained effectively to execute repetitive tasks, facilitated by the teleportation capability of VR. Within a virtual behavioral framework, teleportation functionality affords mice the ability to engage in tasks without the stress associated with physical manipulation, concurrently reducing the setup duration necessary for analogous real-world tasks. During the training sessions, use dim red overhead lighting within the experimental setting. This precaution is recommended due to the diminished visual perceptive sensitivity in mice to red light, which mitigates potential interference with their perception of the virtual reality (VR) screens, as opposed to the use of white light²².

1. Habituation
   1. Start the habituation to the spherical treadmill at the same time as their habituation to fluid regulation to associate the lick tube with the reward using the correctly timed physiological motivation. A three-day habituation period is recommended prior to starting the linear track training.
   2. On the 1^st day, handle the mice for 5 min following weighing. During this interaction, it is advised to gently grasp the head-bar implant while the mice are in their cage, creating familiarity with such manipulation. Introduce them to the area where the VR is housed on this day to allow them to anticipate the spatial environment in which experimental trials will occur. This initial day of habituation coincides with the initiation of fluid regulation of 15 mL per 100 mg of body mass.
   3. On the 2^nd day, which aligns with the transition to the 10 mL per 100 mg of body mass fluid regulation step, again handle the mice for 5 min. Continue repeated gentle grasping of the head-bar while in the cage. Affix the head bar to the holder while allowing mice to familiarize themselves with the spherical treadmill for 5-20 min, either on an infinitely repeating track or without the software program activated. This facilitates their adaptation to the head-fixed condition. It should be anticipated that mice may excrete waste during this period, which typically diminishes over successive sessions.
   4. On the 3^rd day, which corresponds to the final day of the fluid regulation paradigm (5 mL per 100 mg of body mass), handle the mice for 5 min. Then, securely attach them to the air-cushioned spherical treadmill and introduce them to liquid rewards through the reward tube.
      1. Introducing the lick spout to naïve mice will confuse them at first, so make sure the mouse is aware that they are supposed to drink from the tube.
   5. Without being too forceful, apply the positioning of the mouse guidelines below and individualize the positioning of the mouse on the ball with respect to the tube in a way that will deliver the reward to them in a comfortable manner. When starting, make sure the mice are drinking from the tube; this will occur in most mice naturally under water-restricted conditions when presented with liquid to drink.
2. Positioning the mice
   1. Pre-positioning: Prior to placing the mouse on the ball, extend the centered reward tube with a small droplet of reward at the tip. Extend the reward tube before positioning the mouse on the ball to prevent any potential injury resulting from inadvertently extending the tube too far forward once the mouse is head fixed. Elevate the reward tube 5-15 mm above the spherical treadmill such that licking the spout requires a natural forward-facing posture of the head.
   2. Head fixing: To head-fix the mouse, place the mouse on the handler's dominant side of the spherical treadmill. Then, using the handler's dominant hand, pull the mouse by its headbar towards the head-fixing platform. Place the head-bar in the slot meant for fixing, after which using the handler's non-dominant hand, click the head-bar into place.
   3. Location on ball: Individualize placement on the spherical treadmill for each mouse but ensure they meet the following requirements to ensure motivation for tasting the reward and minimize overall levels of stress.
      1. Line the midsagittal plane of the mouse with the center of the spherical treadmill. In cases where the head-bar is not straight, ensure that the midsagittal plane of the mouse, rather than the head-bar, is in line with the center of the placement. For visual clarity, see Figure 3C.
      2. Ensure that the mouse's hind paws are no more than 11 cm from the apex of the spherical treadmill and that the head is behind the apex. Ensure that all four paws are touching the treadmill and that the abdomen can touch the treadmill when the mouse is at rest; this will support proper gait and stability on the ball for running.
      3. When mice do not run, this is called ball refusal. If the mice continue to freeze and do not attempt to run, they are likely experiencing excessive anxiety, and as the investigator choose to exclude them from the experiment. In this study, a quantitative threshold of 5 days of ball refusal was used to determine exclusion from the data.
   4. Side bias: When mice first start habituating to the training routine, they will favor one side over the other. This can interfere with task performance, so take care to ensure that any side preference is not due to asymmetry in how the animal is mounted on the ball. The y-maze task employed here specifically requires the animal to make both right and left choices to optimize reward delivery, which facilitates overcoming side preferences.
   5. Reward spout: This approach involves a gentle maneuver referred to as the kiss it method, where the mouse is guided towards the extended lick tube until its mouth almost touches the tip of the spout, thus ensuring accurate delivery of the reward. Set the duration of the extended reward tube to 1 s when mice receive rewards, permitting the mouse adequate time to consume the droplet fully. Individualize lick spout positioning for each mouse, as the size and preferred positioning for each individual mouse may differ. Ensure the reward tube stays centered throughout all trials for licking standardization; the mouse should always expect to receive the reward in the same physical location regardless of the virtual maze design.
      ​NOTE: While the determination of this duration is at the discretion of the investigator, these findings indicate that this timeframe was effective in facilitating complete reward intake by the mouse before the retraction of the tube. Figure 3B displays an example of a preferable positioning for placement.
   6. Linear track: Consistent with prior studies employing similar methodologies, use a linear track task to investigate two key inquiries: the time required to train mice to traverse a straight corridor and the anticipated success rate of reward acquisition by mice.
      1. Ensure mice have acclimated to both the fluid restriction paradigm and the experimental hardware.
      2. Perform a daily 30 min session to move along a linear virtual corridor starting with a 1 m length. Upon reaching the end of the corridor and receiving the sugar droplet reward, teleport the mice back to the starting point.
      3. Determine a criteria-based advancement to longer mazes (e.g., 1 m, 2 m, 3 m). Advance the mice to the next maze length after achieving 2 consecutive days of receiving an average of 2 rewards per min (Figure 4A).
      4. Document daily recordings of timestamped data concerning reward retrieval, and the distance covered by mice on the spherical treadmill for further analysis (Figure 4B-D).
      5. For mice receiving an average of 2 rewards per min on the 3 m linear tracks, mark them as proficient on the linear track paradigm. It is recommended that mice reach this phase before progressing to more complex behavioral tasks requiring decision-making.
   7. Complex Behavioral Tasks Requiring Decision-Making (Y-Maze): This phase explores the feasibility of progressing from a simple to a more complex behavioral task that requires decision-making. To accomplish this, create a devised signal discrimination Y-Maze task.
      1. In this Y-maze paradigm^23,24, ensure the mice navigate towards a choice point where two arms extend out 45° in either direction like the shape of a Y. Deactivate rotation from the starting point of the maze until reaching the choice point, two arms of varying color, and then activate it within the decision zone to allow the mouse to pivot towards its desired direction.
      2. Upon entry into the arm leading to the reward zone, deactivate the rotation once again. A black arm represents the correct path, while a white arm represents the incorrect path. Use the black arm and white arm as cues to accommodate for potential limitations in mouse visual acuity as they are easily distinguishable, facilitating an examination of their usage of visual information in its simplest form.
      3. Train the mice to navigate towards the black arm to obtain a sugar reward, with each trial concluding with the mice being teleported back to the starting location. Incorporate in the experimental design a random shuffling of the reward location between the left and right sides, ensuring that mice associate the reward with the visual cues rather than the specific side.
      4. Use the same steps for setting up the Y-maze as the linear corridor. Mirror the criteria for progression in the Y-maze paradigm to those of the linear corridor: each trial lasts 30 min, and mice need to reach a predetermined reward threshold for 2 consecutive days. A threshold of 70% rewards correctly acquired is recommended based on the average performance of previous pilot cohorts in the Y-maze; it is above the chance threshold (50%), and it represents a reasonably attainable percentage indicative of mice comprehending the task (Figure 5A).
      5. Upon reaching the choice point, ensure the mouse selects one of the correct or incorrect arms. At the end of the arm, teleport it back to the starting point to repeat the maze within a 30 min timeframe.
      6. This approach employed a visual psychophysics-inspired approach where the mazes got progressively more challenging to distinguish. Follow the description below for the progression in the Y-maze paradigm.
         1. In the initial Y-maze, present solid black and white arms at the maze's choice point. If the mouse correctly chose the black arm for 70% of the trials for 2 consecutive days, progress it to a subsequent level with increasingly challenging discrimination tasks. To achieve this, gradually introduce an additional 10% of the contrasting color to each arm at every level of progression. For example, transition the white arm to being composed of 90% white and 10% black, and vice versa, rendering discrimination more demanding with each advancement.
            NOTE: The idea for increasing is if a 50% white/black can be reached, it would be an effective control as the arms would be indistinguishable. However, the furthest the mice were able to visually discriminate against was 80%:20% (Figure 5B).