OpenScope Predictive Processing Community Project - Data Release

Experimental animals¶

All animal procedures were approved by the Institutional Animal Care and Use Committee (IACUC) at the Allen Institute under protocol 2427 and conducted in accordance with NIH guidelines. Following surgery (see below), all mice were single-housed and maintained on a reverse 12-hour light cycle in a shared facility with room temperatures between 68º and 72ºF and humidity between 30 and 70%. All experiments were performed during the dark cycle. All mice in these experiments were given ad libitum access to food (regular or doxycycline diets) and water.

To systematically collect physiological data, we used standardized data collection and processing pipelines that were previously introduced de Vries et al., 2020Groblewski et al., 2020Durand et al., 2023Bennett et al., 2024Siegle et al., 2021. The data collection workflow progressed from surgical headpost implantation and craniotomy to retinotopic mapping of cortical areas using intrinsic signal imaging, in vivo recording of neuronal activity using various modalities (Neuropixels, Mesoscope two photon imaging, SLAP2 dendritic two-photon imaging), brain fixation and brain histology (see Figure 1). We describe each one of those steps in the dedicated sections below. As part of this workflow, all mice were trained on one of two possible cohorts: A motor cohort and a sequence cohort. Details of each cohort is described in the behavioral training section below. Both behavioral cohorts were recorded with the Neuropixels and Mesoscope recording modality. Only the motor cohort was used for SLAP2 experiments. Each modality (Neuropixels, Mesoscope, SLAP2) was recorded with separate mice as they had different, modality-specific brain implants.

Mesoscopic two-photon calcium imaging experimental animals¶

For experiments involving calcium imaging of GCaMP8s positive cells, male and female transgenic mice (n=14) with pan-neuronal cortical expression of GCaMP8s were used. Snap25-IRES-Cre Harris et al., 2014 was bred in-house and crossed with a GCaMP8s reporter line (Oi4), both of which are maintained on a C57BL/6J background. Snap25-IRES-Cre;Oi4 breeding sets consisted of heterozygous Snap25-IRES-Cre mice (JAX stock #023525) crossed with heterozygous or homozygous TIGRE2-RiboL1-jGCaMP8s-IRES-tTA2 (aka Oi4) mice (JAX stock #039267) Daigle et al., 2018Zhang et al., 2023. Experimental animals were heterozygous for both transgenes (full genotype Snap25-IRES2-Cre/wt;Oi4(TIT2L-jGCaMP8s-RiboL1-WPRE-ICL-IRES-tTA2-WPRE)/wt). Experimental mice show pan-neuronal expression patterns of GCaMP8s in Snap25-expressing cell populations. All mice (including breeders) were maintained on a 200 mg/kg rodent doxycycline diet (Bio-Serv; Flemington, NJ) to suppress tTA2 activity (and subsequent TRE2 promoter-driven expression) during development. Breeding cages were continuously maintained on a doxycycline diet. At weaning (~p20) mice with correct genotype were transferred to cages with standard chow for the remainder of the experiment (PicoLab 5L0D; Lab Diet, Richmod, IN). Despite limiting GCaMP8s expression during development with doxycycline the mice exhibited a sex-dependent reduction in survival rates at older age; at p200 males had a 72% survival rate while females had a 61% rate, n=123). We were unable to identify a clear, single cause for this reduced survival rate.

Neuropixels electrophysiology experimental animals¶

For experiments involving opto-tagging of inhibitory cells, male and female transgenic mice (n=17) expressing ChR2 in Cre-defined cell populations were used. Sst-IRES-Cre mice were bred in-house and crossed with an Ai32 channel rhodopsin reporter line, both maintained on a C57BL/6J background. Sst-IRES-Cre;Ai32 breeding sets consisted of heterozygous Sst-IRES-Cre mice (JAX stock #028864) crossed with homozygous Ai32(RCL-ChR2(H134R)_EYFP) mice (JAX stock #024109) Madisen et al., 2012Taniguchi et al., 2011. Experimental mice were heterozygous for both transgenes (full genotype Sst-IRES-Cre/wt;Ai32(RCL-ChR2(H134R)_EYFP)/wt). Cre+ cells from Ai32 lines are highly photosensitive, due to expression of Channelrhodopsin-2 Zhang et al., 2006.

SLAP2 experimental animals¶

For experiments involving simultaneous glutamate and calcium imaging, male and female wild-type C57BL/6J mice from JAX laboratory injected with Cre-dependent hSyn.FLEX.iGluSnFR4f.NGR and CAG.FLEX.RCaMP3 (or jRGECO1a) AAVs (serotype PHP.eB) were used. Sparse labeling was achieved via low titers (6E+8 vg/mL) of CaMKII-Cre virus. For experiments involving dendritic voltage imaging, male and female wild-type C57BL/6J mice from JAX laboratory injected with Cre dependent ASAP7 virus were used.

Surgery & cranial window procedure: two-photon calcium imaging experiments¶

A subset of mice received a headpost and cranial window surgery as previously described Groblewski et al., 2020de Vries et al., 2020. Headpost and cranial window surgery was performed on healthy male and female transgenic mice (p54-p80) weighing no less than 14 grams at time of surgery. Pre-operative injections of dexamethasone (3-4.2 mg/kg, S.C.) and ceftriaxone (100-125 mg/kg, S.C.) were administered at 1h before surgery. Additionally, carprofen was administered for pain management (5-10 mg/kg, S.C.), and atropine was administered to suppress bronchial secretions and regulate heart rhythms (0.02-0.05 mg/kg, S.C.). Mice were initially anesthetized with 2-4% isoflurane and placed in a stereotaxic frame (Model# 1900, KOPF; Tujunga, CA), and isoflurane levels were maintained at 1.0-2.0% for surgery. An incision was made to remove skin, and the exposed skull was levelled with respect to pitch (bregma-lambda level), roll and yaw. The stereotax was zeroed at lambda using a custom headframe holder equipped with a stylus affixed to a clamp-plate. The stylus was then replaced with the headframe to center the headframe well at 2.8 mm lateral and 1.3 mm anterior to lambda. The headframe was affixed to the skull with white dental cement (C&B Metabond; Parkell; Edgewood, NY) and once dried, the mouse was placed in a custom clamp to position the skull at a rotated angle of 23° such that the visual cortex was horizontal to facilitate creation of the craniotomy. A circular piece of skull 5 mm in diameter was removed, and a durotomy was performed. A glass coverslip (cut from a single piece of glass to obtain a “stacked” appearance that consisted of a 5 mm diameter “core” and 7 mm diameter “flange”), was cemented in place with Vetbond (3M; St. Paul, MN). Dental cement was then applied around the cranial window inside the well to secure the glass window, and subsequently covered with black tempura paint to reduce glare during imaging. The mouse was given 1.0-1.5mL Lactated Ringers Solution (LRS) to help recover from the surgery and replace lost fluids. Post-surgical brain health was documented using a custom photo-documentation system and animals were assessed one, two, and seven days following surgery for overall health (bright, alert and responsive), cranial window clarity and brain health.

Surgery & cranial window procedure: Neuropixels experiments¶

A subset of the mice received the SHIELD surgical procedure, which has previously been described Bennett et al., 2024. The SHIELD procedure was performed on healthy male and female transgenic mice (p53-p139) weighing no less than 14 grams at time of surgery. Pre-operative injections of dexamethasone (3-4.2 mg/kg, S.C.) and ceftriaxone (100-125 mg/kg, S.C.) were administered 1 h before surgery to reduce swelling and postoperative pain. Additionally, carprofen (5-10 mg/kg, S.C.) was administered for pain management, and atropine (0.02-0.05 mg/kg, S.C.) was administered to suppress bronchial secretions and regulate heart rhythms. Mice were initially anesthetized with 2-4% isoflurane and placed in a stereotaxic frame (Model# 1900, KOPF; Tujunga, CA). Isoflurane levels were maintained at 1.0-2.0%, and body temperature was maintained at 37.5°C for the duration of the surgery. An incision was made on the dorsal surface of the skull, and skin was removed in a teardrop shape, exposing the rostral rhinal vein between the eyes, the dorsal surface of the parietal and occipital skull plates, and stopping where the neck muscle begins to attach to the back of the skull. Next, the periosteum was removed from the skull surface to improve adhesion of the cement to the skull and prevent future scabbing and infection. Starting posterior of the left eye, angled forceps were used to separate cheek muscle from the skull, as well as connective tissue and muscle above the left ear. The cheek muscle was then pulled away from the skull and is stretched out so that it makes a seal with the left lateral portion of the well. The exposed skin (and any exposed soft tissue such as the cheek muscle) was then sealed with Vetbond, and the exposed skull was leveled with respect to pitch, roll, and yaw. Once the skull was leveled, bregma was identified using the custom bregma stylus. Without moving the stereotaxic arm in X or Y, the stylus was replaced with a custom “tracer” that provides a guide for marking the craniotomy with respect to bregma. A #11 scalpel blade (or forceps) was used to etch a faint line in the skull around the tracer, which was then replaced with a shallow trench by lightly drilling without breaking through the skull (NeoBurr EF4). After etching was complete, the tracer was replaced with the headframe, which was then lowered in Z to make contact with the skull. Next, dental cement (C&B Metabond, Parkell) was used to attach the headframe to the skull. Once the cement hardened, the headframe was clamped into a custom frame, and the craniotomy and durotomy were performed. After sterilization, a custom implant was then lowered into the craniotomy. To provide a surface that can be glued to the skull, a flange on the implant extends beyond the cranial window and sits directly on the bone, where it was sealed with Vetbond and attached to the skull Once dry, the vetbond was covered with metabond to further secure the implant to the skull. The inner part of the implant sits on the brain surface. Any areas of exposed skull were then covered with cement. Any white cement on the inside of the well was coated with a layer of black tempura paint to reduce glare during ISI imaging. The mouse was given 1.0-1.5mL Lactated Ringers Solution (LRS) to help recover from the surgery and replace lost fluids. After removing the mouse from anesthesia, but prior to it waking up, a photo-documentation image was acquired. Finally, a removable plastic cap was placed over the well to protect the coated implant from cage debris, and the mouse was returned to its home cage for recovery. Over the following 7-14 days, mice were monitored regularly for overall health, cranial window clarity, and brain health.

Surgery & cranial window procedure: SLAP2 experiments¶

A subset of mice received the SLAP2 Visual Cortex laser level procedure. The SLAP2 Visual Cortex laser level procedure was performed on healthy male and female transgenic mice (p53-p139) weighing no less than 14 grams at time of surgery. This is a variation on the Visual Cortex surgery used for the mesoscope experiments. To achieve single cell resolution the optimal angle for the SLAP2 microscope is as close to perpendicular to the craniotomy coverslip as possible. To facilitate this the headframe was attached after the coverslip was secured. This allowed the headframe to be installed on a consistent plane relative to the coverslip, ensuring proper interfacing between the craniotomy and SLAP2 microscope objective. Pre-operative injections of dexamethasone (3-4.2 mg/kg, S.C.) and ceftriaxone (100-125 mg/kg, S.C.) were administered 1 hour before surgery to reduce swelling and postoperative pain and protect against infection. Additionally, carprofen (5-10 mg/kg, S.C.) was administered for pain management, and atropine (0.02-0.05 mg/kg, S.C.) was administered to suppress bronchial secretions and regulate heart rhythms. Mice were initially anesthetized with 4% isoflurane and placed in a stereotaxic frame (Model# 1900, KOPF; Tujunga, CA). Isoflurane levels were maintained at 1.0-2.0%, and body temperature was maintained at 37.5°C for the duration of the surgery. An incision was made on the dorsal surface of the skull. Skin was removed in a teardrop shape; exposing the rostral rhinal vein between the eyes, the dorsal surface of the parietal and occipital skull plates and stopping where the neck muscle begins to attach to the back of the skull. Next, the periosteum was removed from the skull surface to improve adhesion of the cement to the skull and prevent future scabbing and infection. Starting posterior to the left eye, angled forceps were used to separate cheek muscle from the skull, as well as connective tissue and muscle above the left ear. The cheek muscle was then pulled away from the skull and stretched out so that it makes a seal with the left lateral portion of the well. The exposed skin, and any exposed soft tissue such as the cheek muscle, was sealed with Vetbond (3M; St. Paul, MN). The 5mm craniotomy “tracer” provided a guide for marking the craniotomy and was centered at 2.8 mm lateral and 1.3 mm anterior to lambda. Forceps were used to etch a faint line in the skull around the tracer. Using the drill (NeoBurr EF4), without breaking through the skull, the etch was then replaced by a shallow trench. The mouse was rotated clockwise approximately 23 degrees to create a level surface to drill on and help angle the coverslip to accurately interface perpendicularly with the SLAP2 microscope. A well was created with a silicone polymer (Body Double Fast; Smooth-On; East Texas, PA) large enough to encompass the craniotomy and bregma. Craniotomy and durotomy were then performed. The mouse was rotated back 23 degrees counterclockwise for virus injection. A Nanoject III (Drummond; Broomall, PA) was then zeroed at bregma, and virus was injected with a beveled pipette at 3.1 mm lateral, 3.1 mm posterior to and at 3.1 mm lateral, 3.9 mm posterior to bregma, at depths of 0.3 mm and 0.6 mm below surface of the cortex for each point. The mouse was rotated back clockwise approximately 23 degrees to return the craniotomy to an approximately leveled plane relative to pitch, roll, and yaw. A glass coverslip (cut from a single piece of glass to obtain a “stacked” appearance that consisted of a 5 mm diameter “core” and 7 mm diameter “flange”), was lowered into the craniotomy. The 2mm flange was cemented in place with Vetbond. A custom in-house laser leveling tool was used to level the coverslip so that it was perpendicular to the stereotaxic arm. The mouse skull was turned back 23 degrees counterclockwise, and pitch was raised 6 degrees to keep the coverslip in a parallel plane relative to the headframe. This allowed for proper interfacing of the glass coverslip and the SLAP2 microscope when the mouse was head fixed in the headframe. Using the stereotaxic arm, a headframe was lowered onto the skull and secured in place with clear dental cement (C&B Metabond; Parkell; Edgewood, NY), covering all remaining exposed skull. Black tempura paint was applied on top of the clear cement outside the headframe to reduce light leaks. The mouse was given 1.0-1.5mL Lactated Ringers Solution (LRS) to help recover from the surgery and replace lost fluids. After removing the mouse from anesthesia, prior to it waking up, post-surgical brain health was documented using a custom photo-documentation system. Animals were assessed on days one, two, and seven following surgeries for overall health (bright, alert and responsive), cranial window clarity and brain health.

Intrinsic signal imaging / retinotopic mapping¶

Intrinsic signal imaging (ISI) measures the hemodynamic response of the cortex to visual stimulation across the full field of view. This retinotopic mapping represents the spatial relationship between the visual field and cortical locations within each visual area. For both Neuropixels and Mesoscope two-photon imaging experiments, retinotopic maps were used to delineate functionally defined visual area boundaries and to guide targeting of in vivo two-photon calcium imaging to retinotopically defined locations in primary and secondary visual areas.

Animal preparation¶

For every ISI imaging session, mice were lightly anesthetized with 1–1.4% isoflurane delivered via a SomnoSuite system (model 715; Kent Scientific, CT, USA) at a flow rate of 100 mL/min, supplemented with ~95% O₂ (Pureline OC4000; Scivena Scientific, OR, USA). Lubricating eye ointment (Lacri-Lube; Refresh) was applied to maintain corneal hydration and clarity during anesthesia. Mice were positioned on a lab jack and head-fixed such that the cranial window was normal to the imaging axis. The head frame and clamping mechanism ensured consistent positioning of the eye relative to the stimulus monitor across experiments.

Image acquisition system¶

To map retinotopic organization and standardize data acquisition, a custom ISI system coupled to visual stimulation was used. The cortical surface was illuminated with a ring of independently controlled LEDs, including green (peak λ = 527 nm, FWHM = 50 nm; Cree Inc., C503B-GCN-CY0C0791) and red (peak λ = 635 nm, FWHM = 20 nm; Avago Technologies, HLMP-EG08-Y2000) wavelengths mounted on the objective. A tandem lens configuration (Nikon Nikkor 105 mm f/2.8 rear lens and Nikon Nikkor 35 mm f/1.4 front lens) provided 3.0× magnification (M = 105/35). The back focal plane of the front lens was positioned adjacent and coplanar to the cranial window (working distance: 46.5 mm). A bandpass filter (Semrock FF01-630/92 nm) was used to preferentially transmit longer-wavelength reflectance signals while minimizing contamination from the stimulus monitor and ambient light.

Image acquisition and illumination were controlled via custom Python software. Images were acquired using an Andor Zyla 5.5 10-tap sCMOS camera at 40 Hz, with frame timing governed by the camera’s 40 MHz hardware clock. Image acquisition and stimulus presentation were synchronized via hardware triggering from a National Instruments digital I/O board. Raw images (2560 × 2160 pixels, 16-bit) were spatially (4×4) and temporally (4×) binned to yield 640 × 640 pixel frames at 10 Hz with 32-bit dynamic range and an effective pixel size of 10 µm.

Intrinsic imaging visual stimulus¶

The lambda–bregma axis of the skull was oriented at a 30° pitch relative to horizontal, corresponding to an eye position approximately 60° lateral to the midline and 20° above the horizon Oommen & Stahl, 2008. A 24″ monitor was positioned 10 cm from the right eye to maximize visual field coverage. The monitor was rotated 30° relative to the dorsoventral axis and tilted 70° relative to the horizon to maintain perpendicularity to the optic axis.

The visual stimulus consisted of a drifting bar containing a contrast-reversing checkerboard pattern on a gray background. The bar swept across the four cardinal directions at 0.1 Hz, with 10 repetitions per direction Kalatsky & Stryker, 2003. The bar measured 20° × 155°, with individual checker squares of 25°. Stimuli were spatially warped to approximate a spherical visual field projection on a flat display Marshel et al., 2011.

Image acquisition and processing¶

A high-resolution image of the cortical vasculature was first acquired under green illumination to serve as a fiducial reference. The imaging plane was then defocused 500–1500 µm below the surface to capture intrinsic signals. Up to 10 ISI time series were collected per experiment.

Time series were preprocessed by removing the time-averaged pixel intensity (DC component) to improve signal-to-noise ratio. A discrete Fourier transform (DFT) was computed at the stimulus frequency. Phase maps were generated from the phase angle of the DFT and used to map visual field position onto cortical coordinates. Sign maps were derived by computing the sine of the angle between the gradients of the altitude and azimuth phase maps. Final sign maps were averaged across at least three time series (minimum of 30 sweeps per direction).

Automated sign map segmentation and annotation¶

For each experiment, visual field sign maps were segmented into distinct visual areas using an automated algorithm (adapted from Garrett et al., 2014). Segmentation was based on three criteria: (1) each area contains a uniform visual field sign, (2) each area represents a unique (non-redundant) portion of visual space, and (3) adjacent areas with the same sign exhibit redundant visual field representations.

Eccentricity and target map generation¶

Eccentricity maps were computed relative to the center of visual space (0° azimuth, 0° altitude). When the eye is properly aligned, this point corresponds approximately to the anatomical center of V1. The eccentricity at the V1 centroid was used as a quality control metric to identify experiments with significant eye misalignment (>15°).

To define target regions for two-photon imaging, eccentricity maps were thresholded to include locations within 10° of the V1 center. These targeting maps were overlaid onto vasculature images to provide fiducial landmarks for aligning imaging fields across sessions and ensuring retinotopic consistency.

ISI quality control¶

Quality control of ISI-derived maps consisted of four steps:

1. Brain surface and vasculature images were inspected before and after acquisition for clarity, focus, and cranial window positioning.

2. Individual trials were evaluated for sufficient visual coverage, continuity of phase maps, localization of amplitude maps, and expected sign map organization. Only trials meeting these criteria were included (minimum of three trials).

3. Automated segmentation was required to identify at least six visual areas (VISp, VISlm, VISrl, VISal, VISam, VISpm).

4. Final maps were assessed for visual field coverage (35–60° altitude, 60–100° azimuth), minimal bias (<10° range imbalance), alignment of the retinotopic center with the V1 centroid, and minimum V1 area (>2.8 mm²).

Behavioral training¶

Mice were trained in individual sound-attenuating enclosures arranged in clusters. Each enclosure was arranged similarly to that of the mesoscope two-photon microscope, SLAP2 two-photon imaging microscope and the Neuropixels rigs. A 24” LCD monitor was positioned 15 cm from the mouse’s right eye, with the sagittal axis of the head parallel to the screen. A registered headframe clamp was attached to a behavior stage equipped with kinematic mounts to ensure repeatable placement of the stage in the enclosure. Each stage consisted of a fixed-position headframe clamp and adjustable running wheel. Enclosures were equipped with a camera coupled with IR illumination to monitor mouse activity. Those videos were recorded but kept temporarily on disk.

Animals received a 2-week training procedure following a previously published head-fixation habituation protocol de Vries et al., 2020.

Habituation to handling and head-fixation was performed for five days: On days 1-2 mice were removed from the home cage and gently handled for 1-2 min. On days 3-5 mice were removed from the home cage and handled for 1-2 min, then head-fixed by securing the headframe in the behavior stage clamping mechanism. The stage was then placed in the lit behavior enclosure for 5-10 min.

Passive behavior training was then performed to habituate mice to extended periods of head-fixation and expose the mice to visual stimuli. On days 6-10 mice were removed from the home cage and handled for 1-2 min. Mice were then head-fixed to the behavior stage and the stage was placed in the behavior enclosure. A set of full-screen stimuli was displayed for increasing periods. Visual stimuli were displayed for increasing durations of 8, 18, 28, 38, and 48 min on days 6 through 10, respectively.

The habituation protocol included two cohorts of mice. (1) The sequence cohort passively viewed a repeating sequence of four drifting gratings (90°–45°–0°–45°) followed by a grey inter-sequence interval, without any mismatch events. Each element was presented for 250 ms, and gratings were full-field (0.04 cpd, 2 Hz temporal frequency, 100% contrast). Animals were free to walk and rotate the disc beneath their body; however, disc rotation had no effect on the stimuli presented on the screen. This established a learned expectation for the sequential structure. (2) The motor cohort was habituated in a closed-loop paradigm in which locomotion on the running disc controlled the phase of a full-field sinusoidal grating (0° orientation, 0.04 cpd) via a rotary encoder. This configuration approximated natural visual flow, such that forward locomotion produced corresponding backward grating motion consistent with that experienced by a freely moving mouse. No mismatch events were presented during any habituation sessions in either cohort. The full-length habituation sessions additionally included control blocks (standard control, sequential control, jitter control, open-loop prerecorded, natural movie, and receptive field mapping) identical to those used in the experimental sessions, further habituating the animals to the full session structure.

Visual stimulation¶

All visual stimuli were generated using BonVision Lopes et al., 2021, an open-source visual environment package running within the Bonsai reactive programming framework Lopes et al., 2015. For behavior training, Neuropixels recordings and mesoscope imaging, stimuli were rendered at 60 Hz and displayed on a gamma-calibrated ASUS PA248Q LCD monitor (1920 × 1200 pixels, 55.7 cm wide, 60 Hz refresh rate) positioned 15 cm from the animal’s right eye (see Figure 1). A spherical warping correction (BonVision SphereMapping) was applied to all stimuli to compensate for the close viewing distance and flat display geometry, ensuring that apparent size, speed, and spatial frequency were constant across the visual field as seen from the mouse’s perspective. The monitor subtended 120° × 95° of visual space. Mean luminance was 50 cd/m². Stimulus timing was synchronized to neural recordings via a photodiode placed on a sync square region of the monitor that alternated between black and white every 60 frames, and via digital synchronization pulses sent to a National Instruments digital board. The full stimulus code, Bonsai workflow, and parameter files are available on the project’s GitHub repository (https://github.com/AllenNeuralDynamics/openscope-community-predictive-processing). For SLAP2 recordings, the stimulation screen was smaller to account for physical constraints of the SLAP2 rig. Work in progress: verify and add the SLAP2 display model, pixel dimensions, physical size, and visual-angle coverage.

For each session, a stimulus table (CSV file) was generated programmatically by a Python script (generate_experiment_csv.py) using a session-specific random seed derived from the session UUID and timestamp, ensuring unique trial sequences across sessions while maintaining reproducibility. This CSV table specified all trial parameters (orientation, spatial frequency, temporal frequency, contrast, duration, delay, position, phase, trial type, and block membership) and was read by the Bonsai workflow (generic_oddball.bonsai) to drive stimulus presentation in sequence.

In the sensorimotor mismatch context, the phase of the drifting grating was coupled to the angular position of the running disc via a rotary encoder. The wheel-to-visual coupling was computed as: phase (radians) = 2π × R × θ / tan(1/f × π/180), where R is the wheel radius-to-screen ratio (0.36), θ is the wheel angle in degrees, and f is the spatial frequency (0.04 cpd). This coupling was calibrated so that the resulting visual flow approximated the optic flow a freely moving mouse would experience during forward locomotion.

Stimuli parameters¶

All drifting grating stimuli shared the following base parameters unless otherwise specified: spatial frequency 0.04 cpd, temporal frequency 2 Hz, 100% contrast, sinusoidal luminance profile, full-field extent (360° diameter with spherical correction). For the standard oddball, sequence mismatch, and duration mismatch sessions, gratings drifted at a fixed temporal frequency of 2 Hz. For the sensorimotor mismatch session, the grating phase was updated at 30 Hz (every other video frame) based on wheel rotation, with temporal frequency set to 0 in the stimulus table (wheel-controlled mode). Oddball/mismatch events occurred at a rate of 1.35 per minute per deviant type (5.4/min total across four deviant types) in all session types. The stimulus parameters for each session type and shared control blocks are detailed in the Experimental design section above.

Neuronal recording modalities¶

Neuropixels extracellular electrophysiology.¶

Habituation to the Neuropixels rigs.¶

Prior to the first recording, mice were habituated to the Neuropixels rigs every day for a week. These sessions were similar to the recording session in length but did not include oddballs. There were 2 cohorts of mice, one exposed to motor stimuli and the other to the sequence stimuli (see stimuli description).

Implant design for recordings¶

The implant was designed as stated in the surgery section and was described previously Bennett et al., 2024. Briefly, we created a CAD file with holes strategically placed above the areas of interest for our study (see Supplementary Figure 1). Some hole coordinates have been previously validated to allow targeting of areas such as the center of VISp, and other hole coordinates were derived from publications, such as VISa Lyamzin & Benucci, 2019. Hole positions were adjusted using Pinpoint Birman et al., 2023. Note that the placement of the implant on the brain can vary slightly due to mouse-to-mouse variability and surgical precision. The coordinates of holes were referenced from bregma; their diameters and intended target areas are represented in Supplementary Figure 1.

SORTA-Clear plug removal and agarose application¶

To prepare the brain for recording, the SORTA-Clear coating over the implant is removed and replaced with a temporary layer of Kwik-Cast (World Precision Instruments). The mouse is anesthetized with isoflurane (5% induction, 1-2% maintenance, 100% O2) and eyes protected with ocular lubricant (I-DROP, VetPLUS). Body temperature is maintained at 37.5°C (TC-1000 temperature controller, CWE, Incorporated). The well is cleaned of any debris using ethanol swabs. Then, the inside of the well surrounding the SORTA-Clear plug is painted with white Metabond to improve visibility during probe insertion. Once the Metabond is dry, the well is flooded with enough ACSF to completely submerge the SORTA-Clear sheet, which is then removed with small forceps, starting at the anterior or posterior end of the sheet and peeling gently to remove it in one piece. Once the SORTA-Clear sheet is completely detached from the implant, the edges of implant holes are tested with small forceps to ensure all holes are free of SORTA-clear or debris. ACSF is removed from the well using Sugi spears (Kettenbach) and the well is filled with Kwik-Cast. Once the Kwik-Cast is fully dry, a plastic protective cap is secured on the well to protect against debris and the mice are returned to their home cage. This preparation is generally performed Fridays for recordings on Mondays to avoid using isoflurane on the day of experiment.

Head fixation.¶

On the day of recording, the mouse is removed from its home cage and clamped to the running wheel on the experimental rig. Wheel height is adjusted as needed for each mouse. Once head-fixed, the protective well cap and Kwik-Cast layer are removed. The ground wire is tucked into the side of the well and any excess debris cleaned using a Sugi spear or cotton tipped applicator. Approximately 0.2ml of agarose (4%: 0.4 g of BioRad Low Melt and 0.4 g of Sigma agarose high electroendosmosis in 20.2 ml ACSF Durand et al., 2023 is applied in a smooth layer over the entire implant surface and ground wire. After popping any large bubbles, the agarose is allowed to set for ∼10 seconds. To prevent the agarose from drying out during the experiment, a layer of silicon oil is applied over exposed agarose with a toothpick. At the end of the experiment, the agar is removed and replaced with Kwik-Cast.

The 3D-printed protective cone was then lowered to prevent the mouse’s tail from striking the probes. An infrared dichroic mirror was placed in front of the right eye to allow the eye-tracking camera to operate without interference from the visual stimulus.

Grounding.¶

A 32 AWG silver wire (A-M Systems) is epoxied to the headframe during the implant surgery and served as the ground connection. The wire is pre-soldered to a gold pin embedded in the headframe well, which mates with a second gold pin on the protective cone. This second gold pin is connected to both the behavior stage and the probe ground. Prior to the experiment, the brain-to-probe ground path is checked using a multimeter. The reference connection on the Neuropixels probes is permanently soldered to ground using a silver wire, and all recordings are made using the tip reference configuration. The headstage grounds (which are contiguous with the Neuropixels probe grounds) are connected with 36 AWG copper wire (Phoenix Wire). All probes are connected in parallel to animal ground.

Neuropixels probes¶

All neural recordings were carried out with Neuropixels 1.0 probes Jun et al., 2017, as previously described Siegle et al., 2021Durand et al., 2023. The 384 electrodes closest to the tip are used, providing a maximum of 3.84 mm of tissue coverage. The signals from each recording site are split in hardware into a spike band (30 kHz sampling rate, 500 Hz highpass filter) and a Local Field Potential (LFP) band (2.5 kHz sampling rate, 1000 Hz lowpass filter). Our goal was to insert six probes into the same mouse’s brain on each of 4 consecutive days. To distinguish the paths of these different penetrations, we use two dyes, CM-DiI (1 mM in ethanol; ThermoFisher Product #V22888) on day 1 and 2 and CM-DiD (1mM in ethanol; AAT Bioquest catalogue number 22060) on day 3 and 4. The probes are coated with dye before recordings by immersing them at least 3mm into a well filled with dye. Each probe are dipped five times to ensure adequate coating.

Neuropixels probe insertion¶

Our custom experimental rig can insert up to six Neuropixels probes simultaneously (see Figure 3). Each probe is mounted on a separate 3-axis micromanipulator with a 15 mm travel range (New Scale Technologies, Victor, NY). Probes are driven to their target holes and lowered to the surface of the brain while the experimentalist monitores a camera feed to avoid vasculature and watches real-time signals on the OpenEphys GUI to identify activity indicative of the brain surface. If the probe need adjustment when attempting to insert (e.g. to avoid vessels), the probe are completely retracted out of the silicon oil to prevent probe bending. Once all probes reach the brain surface, each probe is zeroed and set to insert 3100 μm deep at 200 μm/min and then retracted 100 μm to their final depths to reduce tissue compression and subsequent electrode zdrift relative to the brain. Once all probes reach their final depth, the probes are allowed to settle for ∼30 minutes, and a photo documentation of the inserted probes is captured. Sometimes a probe can not be inserted into its assigned hole, failures are generally due to dura regrowth. Overall, we achieved a penetration success of 5.8 probes per session.

Data acquisition and synchronization.¶

Neuropixels data is acquired at 30 kHz (spike band) and 2.5 kHz (LFP band) using the Open Ephys GUI Siegle et al., 2017. Gain settings of 500× and 250× are used for the spike band and LFP band, respectively. Probes are connected to a PXIe card inside a National Instruments chassis.

Videos of the eye, body and face are acquired at 60 Hz. The angular velocity of the running wheel is recorded at the time of each stimulus frame, at approximately 60 Hz. Synchronization signals for each frame are acquired by a dedicated computer with a National Instruments card acquiring digital inputs at 100 kHz, which is considered the master clock. A 32-bit digital ‘barcode’ is sent with an Arduino Uno (SparkFun DEV-11021) every 30 s to synchronize all devices with the neural data. Each Neuropixels probe has an independent sample rate between 29,999.90 Hz and 30,000.31 Hz, making it necessary to align the samples offline to achieve precise synchronization. The synchronization procedure uses the first matching barcode between each probe and the master clock to determine the clock offset, and the last matching barcode to determine the clock scaling factor.

To synchronize the visual stimulus to the master clock, a silicon photodiode (PDA36A, Thorlabs) was placed on the stimulus monitor above a “sync square” that alternated between black and white every 60 frames.

Stimulus Monitor¶

We lower a black curtain over the front of the rig, placing the mouse in complete darkness except for the visual stimulus monitor. Visual stimuli are generated using custom scripts based on PsychoPy⁹ and are displayed using an ASUS PA248Q LCD monitor, with 1,920 × 1,200 pixels (55.7 cm wide, 60 Hz refresh rate). Stimuli are presented monocularly, and the monitor is positioned 15 cm from the right eye of the mouse and spans 120° × 95° of visual space before stimulus warping. Each monitor is gamma corrected and has a mean luminance of 50 cd m⁻². To account for the close viewing angle of the mouse, a spherical warping is applied to all stimuli to ensure that the apparent size, speed and spatial frequency were constant across the monitor as seen from the mouse’s perspective.

Probe removal and cleaning¶

After each experiment, the probes are retracted from the brain at a rate of ~3000μm/min and the mouse is removed from head fixation and returned to its home cage. If another recording session is to occur the same day, the probes are then immersed in a well of freshly made 1% Tergazyme mixed with agarose for 5 minutes to remove excess residue, followed by immersion in 1% Tergazyme for 30 minutes, immersion in Milli-Q water for 25 minutes, and dipped in 100% isopropyl alcohol for 1 minute. After the last recording session of the day, the probes are immersed in a well of freshly made 1% Tergazyme for ~12 hours.

Quality Control for Neuropixels recording sessions¶

Possible QC failure can come from these cases: white foam buildup on the edge of the eye covering the pupil, software failures compromising critical data streams, visual stimulus synchronizing failure, cortical bleeding or compromised brain health resulting in low unit activity and/or atypical visual responses, gap in data acquisition and discovery of purulent material over right hemisphere during ex-vivo imaging. Out of a total of 56 sessions, 12 are excluded for eye foam.

Optotagging protocol¶

At the end of every experiment, an optotagging protocol is run during which the cortical surface is stimulated with blue light. In Sst-IRES-Cre/wt;Ai32(RCL-ChR2(H134R)_EYFP)/wt mice, this protocol allowes us to identify putative Sst+ cortical interneurons by an increase in spiking activity time-locked to laser stimulation (and consequent ChR2 activation). Blue light is delivered by a 473 nm laser (Laser Quantum, model Ciel or Cobolt model 06-MLD). The light source is coupled to a 400 μm diameter fiber optic cable (Thorlabs) or bifurcated fiber bundle (Thorlabs, BFYL4LF01), with the tip(s) positioned such that blue light illuminates the entire cranial window. Two types of stimuli at 3 different light levels are randomly interleaved: a 10 ms pulse, and a 1s raised cosine ramp. For the pulse stimulus, a 0.5 ms ramp is applied at the beginning and end of the pulse. Stimuli are presented at intervals of 1.5 s plus a uniformly distributed delay between 0 and 0.5 s.

Clearing with life canvas¶

We use published protocols to perform the tissue sample preparation steps for clearing a whole mouse brain. Briefly, the brain is perfused and fixed in 4% paraformaldehyde in order to prepare it for light sheet microscopy. In a timeline of two weeks, the brain will be stripped of lipids Myers & Toglia, 2023 and rendered transparent in an index matching solution Myers & Toglia, 2023, allowing for viewing the morphology of anatomical brain structures. Then the brain is embedded in agarose for imaging Myers & Toglia, 2023. This protocol collection is ideal for experiments where high quality clearing is desired for imaging finer cell structures that are located deep in the brain, or when it is necessary to preserve endogenous fluorescence.

Imaging of cleared brains for Neuropixels probe trajectory¶

Agarose blocks containing cleared mouse brains were fixed to the sample arm of a light sheet microscope (LifeCanvas Technologies) and submerged into an immersion oil bath matching the refractive index for the clearing technique described above (Cargille Laboratories). Specimens were oriented such that the light sheet and focal plane aligned to the transverse anatomical plane with the superior surface closest to the imaging objective and the excitation light entering from either the left or right hemisphere, determined relative to the sagittal mid-plane. Data was collected using a 4X 0.20 NA objective, modified to 3.6X for oil immersion, stepped axially to produce voxels 2.0 x 1.8 x 1.8 𝞵m^3 in (z, y, x).

Raw data was then packaged with relevant metadata, uploaded to cloud storage, and used to create derived data, including contiguous image volumes for each channel, transform fields mapping to/from the Allen CCFv3, and neuroglancer viewer links for visualizing results.

Two-photon mesoscope calcium imaging¶

Multi-plane calcium imaging was performed using a dual-beam mesoscope (Multiscope), enabling simultaneous imaging of two planes and effectively doubling imaging throughput (Orlova, Tsyboulski, Najafi et al., 2020). The system builds on the 2P-RAM platform (Sofroniew et al., 2016) with a compact optomechanical add-on that introduces a second excitation beam and simplifies alignment.

The dual-beam configuration consists of: (1) a delay line to split the excitation beam and temporally offset one beam by half the laser pulse period; (2) a secondary z-scanner to independently position each beam along the axial (z) dimension; and (3) a custom demultiplexing unit. Temporal encoding of the excitation beams enables separation of fluorescence signals based on photon arrival time at the detector. Laser excitation was provided by a Coherent Axon laser operating at 910 nm.

The system was controlled using customized ScanImage software (Vidrio Technologies) in conjunction with an in-house workflow sequencing engine (WSE; see below). Emitted fluorescence was detected with a single photomultiplier tube (PMT), and signals from the two imaging planes were separated using a custom analog demultiplexing circuit (Orlova et al., 2020). Demultiplexing was achieved by multiplying the PMT signal with two complementary square waveforms corresponding to the temporal windows of each excitation beam.

The integration window (6.25 ns; half the laser pulse period) does not fully capture the fluorescence decay, resulting in partial signal bleed-through between channels (~10% inter-plane crosstalk). This residual crosstalk was reduced using an independent component analysis (ICA)-based demixing algorithm (see below). National Instruments data acquisition hardware (PXI chassis, PXIe-6363 boards) was used for system control and data acquisition.

To coordinate hardware and software components, a workflow sequencing engine (WSE) was developed in Python. The WSE uses a distributed messaging interface to communicate with ScanImage, the stimulus presentation computer, synchronization hardware, and behavioral monitoring systems (body and eye tracking). The WSE also integrates user-guided steps: when manual intervention is required (e.g., hardware adjustments), the system prompts the operator and records task completion. Upon completion of each experiment, the WSE aggregates all data streams and automatically initiates transfer to a centralized data repository.

Habituation to Mesoscope rig¶

Prior to the first imaging session, mice were habituated to the imaging rig under head fixation for 30 minutes. Habituation sessions were conducted under the same ambient conditions as experimental recordings (dim red light, imaging environment) but without visual stimulus presentation. These sessions allowed animals to acclimate to head fixation, the rotating disk, and the experimental setup prior to data collection.

During habituation, the mesoscope objective was aligned to be as close as possible to perpendicular to the cranial window to optimize imaging quality. Alignment was verified using an infrared (IR) viewer by directing the excitation beam onto the cranial window and confirming that the reflected signal returned to the objective. Stage coordinates (X, Y, R1 & R2 - objective rotation) were recorded to facilitate consistent field-of-view positioning and cell matching across subsequent imaging sessions.

Mesoscope imaging data collection¶

All experimental setup was performed under dim red illumination to preserve the reversed light–dark cycle; imaging was conducted in darkness. Mice were head-fixed on a freely rotating disk, allowing voluntary locomotion. During imaging, the mouse eye was positioned 15 cm from the display. The screen center was located 118.6 mm lateral, 86.2 mm anterior, and 31.6 mm dorsal relative to the right eye, aligning the display normal to the average gaze axis.

The disk surface was covered with removable foam (Super-Resilient Foam, McMaster) to reduce motion-related artifacts. Water-based ultrasonic gel was used as the immersion medium to minimize evaporation and leakage during imaging.

On the first imaging day, ISI-derived targeting maps generated for each animal were used to identify regions of interest (ROIs) in VISp and VISl. Target locations were verified by registering the ISI-derived targeting map, overlaid on a reference image of the cortical surface vasculature, to live epifluorescence images acquired under blue light illumination. Alignment was performed using superficial vascular landmarks (e.g., vessel branching patterns and intersections) across the 5 mm cranial window, enabling accurate localization of regions of interest and consistent targeting across imaging sessions.

Following this alignment, the cranial window was visualized under two-photon (2P) imaging. The live 2P surface image was compared to the ISI-targeting map and vasculature reference to define ROIs for each recording site (400 × 400 µm field of view; 512 × 512 pixels). For each ROI, a 2P reference image of the cortical surface was acquired.

Imaging planes were then positioned relative to the cortical surface at depths corresponding to cortical layers I (~0–100 µm), II/III (~100–300 µm), IV (~300–400 µm), and V (~400–500 µm), with exact depths adjusted based on cortical landmarks and image features. A reference image was acquired at each imaging plane to support subsequent field-of-view alignment and subsequent session cell matching.

After acquisition of reference images at the cortical surface and imaging depths, a z-stack centered on the imaging plane (±30 µm, 0.75 µm step size) was collected to assess cortical structure and estimate axial motion (see Quality Control, Z-axis stability). At the end of the first recording session, a widefield epifluorescence image of the cranial window is also acquired. This image is used for aligning and targeting the same field of views for successive imaging sessions.

For subsequent imaging sessions, previously defined imaging fields were re-identified using a stepwise alignment procedure to reproduce the field of view and imaging depth established during the initial imaging session.

1. The mesoscope objective was first returned to the reference tilt (R1 and R2) and stage coordinates (x and y) established during habituation, providing an initial estimate of the imaging location.

2. A live epifluorescence image of the cranial window was acquired and aligned to the reference epifluorescence image from the initial imaging session using superficial vascular landmarks to recover the targeted cortical regions.

3. A two-photon image of the cortical surface was then acquired and matched to the corresponding surface reference image from the initial imaging session to refine field-of-view alignment.

4. Finally, the imaging plane was adjusted in the x-, y-, and z-axes until cellular features and other anatomical landmarks matched those in the reference images acquired at the target imaging depth, enabling reliable field-of-view and cell matching across imaging sessions.

Imaging parameters¶

After stabilizing the imaging plane, PMT gain and laser power were adjusted to maximize signal-to-noise ratio and dynamic range while limiting saturation (<1000 saturated pixels per frame). A predefined power lookup table guided parameter selection. Signal intensity between planes was balanced by adjusting beam power while monitoring pixel intensity histograms.

Laser power was selected from the depth-dependent lookup ranges.

Two-photon imaging data (512 × 512 pixels; 11 Hz per plane for multi-plane acquisitions), eye tracking (30 Hz), and behavioral video (30 Hz) were recorded simultaneously and continuously monitored. Recording sessions were approximately 60 minutes in duration, with total imaging sessions lasting up to 75 minutes including setup and calibration. Sessions were terminated early if animals exhibited signs of stress (e.g., excessive periocular secretion, abnormal posture) or if data quality was compromised by technical issues, including loss of synchronization between data streams, photomultiplier tube (PMT) signal instability, or dropped imaging frames.

A total of 15 mice were allocated for the 2-photon workflow. The Sequence & Motor cohorts each consists of 5 completed data sets with each mouse undergoing 8 imaging sessions. Additional sessions were acquired as needed to replace datasets that failed quality control. 5 mice were removed from the workflow due to health-related issues.

Quality control for two-photon calcium imaging¶

Quality control metrics were evaluated after each session. Sessions failing any criterion were repeated.

1. Image saturation: Initial frames were inspected to ensure fewer than 1000 saturated pixels and adequate use of the detector dynamic range.

2. Photobleaching: Baseline fluorescence at the beginning and end of the session was compared; sessions with >20% signal loss were excluded.

3. Targeting accuracy: Imaging locations were registered to ISI-derived maps to confirm correct visual area targeting.

4. Z-axis stability: Mean images from the first and last 5 minutes were compared to a post hoc z-stack (±30 µm, 0.75 µm steps) to estimate drift. Sessions with >10 µm drift were excluded.

5. Animal well-being: Behavioral videos were reviewed for signs of stress (e.g., excessive secretion, orbital tightening, abnormal posture). Animals exhibiting sustained stress responses were removed from the experiment.

6. Temporal synchronization: Alignment across all recorded data streams was verified.

7. System integrity: Data streams were assessed for hardware or software failures affecting data quality.

8. Motion artifacts: Residual motion was evaluated after motion correction.

9. Interictal activity: Full-field fluorescence traces (first 10,000 frames) were analyzed for abnormal transient events. Sessions with potential interictal activity were manually reviewed and excluded if necessary Steinmetz et al., 2017.

Metrics used for each criteria are available on the AWS S3 bucket in a qc.json file

SLAP2 dendritic imaging.¶

Dual-color imaging of synaptic glutamate and somatic calcium in single neurons was performed using SLAP2. SLAP2 allows for simultaneous measurement of arbitrarily-shaped ROIs across two imaging planes. Citation needed: add the SLAP2 methods paper when available. We recorded from Layer 2/3 pyramidal neurons in the visual cortex. We imaged from soma and several peri-somatic dendritic segments in one plane, and imaged several apical dendritic segments on the second plane, typically achieving recordings of >100 synapses at >200 Hz each. Imaging was motion stabilized by using SLAP2’s image-based online motion correction.

Acquisition of reference stacks and ROI selection¶

Prior to functional imaging, structural reference stacks were acquired for motion correction and ROI definition. For each imaging plane, a z-stack consisting of 21 optical sections spaced 1 um apart was collected, averaging 35--45 repeated acquisitions per section . One reference stack was centered on the soma and proximal dendrites, whereas the second stack was centered on apical dendritic regions.

Reference stacks were aligned during the imaging session and used to define imaging ROIs. ROIs were drawn manually encompassing dendritic segments containing visually identifiable spines and algorithmically refined to exclude dark background pixels. Restricting ROIs to dendritic shafts and spines reduces the number of imaged pixels and increases sampling rate.

The reference stacks were also used for online motion correction during functional imaging. Lateral (x-y) displacements were estimated by registering incoming data to the reference volumes and were used to update the DMD illumination patterns in real time. Axial (z) motion was compensated independently using a remote-focusing system, allowing ROIs to remain aligned to the targeted neuronal structures throughout the recording session.

Data processing¶

Neuropixels extracellular electrophysiology.¶

Raw data was processed using the AIND ephys pipeline Allen Institute for Neural Dynamics, 2026 on the Code Ocean platform. In brief, the pipeline is implemented in Nextflow DSL2 and each probe was processed in parallel with the following steps:

• Preprocessing: including phase shift, highpass filter, bad channel detection and removal, common median reference

• Spike Sorting with Kilosort version 4

• Postprocessing: computing additional extensions (e.g., waveforms, spike amplitudes, PCA scores) and quality metrics

• Curation: applying quality-metric based thresholding and UnitRefine pre-trained classifiers Jain et al., 2025

• Visualization and QC: generation of plots for quality control of raw and spike sorted data

For more details, please refer to Buccino et al., 2026

Identification of brain areas associated with Neuropixels nodes¶

After brains are processed in the imaging pipeline, neuroglancer (https://neuroglancer-demo.appspot.com) is used to reconstruct probe tracks in the brain. Points are placed along the length of the probe track and are closely inspected to ensure annotation of the probe tip and each track is assigned to a particular day of recording.

Electrophysiology features recorded from neural probes are aligned with anatomical landmarks based on the Allen Mouse Brain Common Coordinate Framework (CCFv3) Wang et al., 2020. For this, we use the IBL ephys alignment GUI (https://github.com/AllenNeuralDynamics/ibl-ephys-alignment-gui). Based on firing rate, LFP power, as well as spike and LFP cross correlograms, we place reference lines delineating borders of areas. Through this procedure, we consistently align the top of the cortex (Layers 2/3, where we expect spiking activity) using cross correlograms. Other alignment locations such as white matter tracks, thalamus, and subcortical regions are only made when there are clear electrophysiological landmarks (i.e. a stark increase in firing rate for thalamus, a stark decrease in activity in white matter, etc). If features such as these are not present, the GUI’s probe track interpolation is accepted. Notably, this interpolation is accepted for cortical layer assignments between Layers 2/3 and the bottom of cortex. We also ensure that the channel assignments on the probe track are within 10% of the expected scaling factor.

Two-photon mesoscope calcium imaging.¶

Raw two-photon calcium imaging data were processed using the AIND planar optical physiology pipeline (aind-pophys-pipeline v11 and v13; https://github.com/AllenNeuralDynamics/aind-pophys-pipeline) on the Code Ocean platform. The pipeline is implemented in Nextflow DSL2 and processes each imaging plane independently and in parallel through the following steps:

• Data conversion: For multiplane mesoscope data acquired with the dual-beam configuration, interleaved TIFF files were de-interleaved into individual imaging planes and stored as separate HDF5 timeseries using the aind-pophys-converter-capsule (https://github.com/AllenNeuralDynamics/aind-pophys-converter-capsule).

• Motion correction: Non-rigid (piecewise rigid) motion correction was performed on each plane using Suite2p Pachitariu et al., 2016 (https://github.com/MouseLand/suite2p), implemented in the aind-ophys-motion-correction capsule (https://github.com/AllenNeuralDynamics/aind-ophys-motion-correction). Default parameters included a maximum registration shift of 10% of the field of view (maxregshift = 0.1), Gaussian spatial smoothing with σ = 1.15 pixels, a maximum non-rigid shift of 5 pixels per block (maxregshiftNR = 5), and a signal-to-noise threshold of 1.2 for block smoothing (snr_thresh = 1.2). Frames were processed in batches of 500.

• Decrosstalk: Because the dual-beam mesoscope acquires pairs of imaging planes with temporally offset excitation beams that are separated by analog demultiplexing, the incomplete capture of fluorescence decay within the 6.25 ns integration window produces approximately 10% inter-plane signal crosstalk. To correct this residual bleed-through, paired planes were identified from session metadata using the aind-ophys-group-planes capsule (https://github.com/AllenNeuralDynamics/aind-ophys-group-planes), and an independent component analysis (ICA)-based demixing algorithm was applied to the motion-corrected plane pairs using the aind-ophys-decrosstalk-roi-images capsule (https://github.com/AllenNeuralDynamics/aind-ophys-decrosstalk-roi-images).

• Cell segmentation and trace extraction: Regions of interest (ROIs) corresponding to individual neurons were detected using a cell detection algorithm, implemented in the aind-ophys-extraction capsule (https://github.com/AllenNeuralDynamics/aind-ophys-extraction).

  The default configuration used Suite2p’s sparse detection mode (init = sparsery) with automatic diameter estimation (diameter = 0), a cell probability threshold of 0.0 (cellprob_threshold = 0.0), threshold scaling of 1, and a maximum overlap of 75% between ROIs (max_overlap = 0.75). For each detected ROI, a raw fluorescence trace was computed by averaging pixel intensities within the ROI footprint. Neuropil contamination was estimated from a surrounding annular region and subtracted using a neuropil correction coefficient determined by minimizing the mutual information between the corrected trace and the neuropil signal. Suite2p’s built-in classifier was used to provide an initial cell/non-cell classification for each ROI.

• ROI classification: Following extraction, a GPU-accelerated ROI classification step was applied to each imaging plane using a pre-trained classifier (aind-ophys-classifier capsule). The classifier, trained using the ROICaT framework (Region Of Interest Classification and Tracking; https://github.com/richiehakim/ROICaT), categorized each detected ROI as a cell or non-cell based on learned spatial features. The classification results were stored in a separate classification.h5 file for each plane and used to label ROIs in the final NWB output.

• ΔF/F computation: Baseline-corrected fluorescence traces (ΔF/F) were computed from the neuropil-corrected traces using the aind-ophys-dff capsule (https://github.com/AllenNeuralDynamics/aind-ophys-dff). The algorithm proceeded as follows: (1) the noise standard deviation σ was estimated using the median absolute deviation (MAD) method; (2) an initial baseline b was estimated; (3) active frames were identified as outliers exceeding b + 3σ and masked; (4) the baseline fluorescence F₀ was obtained by median-filtering the trace over a 60 s sliding window using only inactive frames, with interpolation across masked segments; (5) ΔF/F was computed as (F − F₀) / F₀. A short window of 3.333 s was used for local noise estimation, and the inactive percentile was set to 10.

• Event detection: Deconvolved neural events were extracted from the ΔF/F traces using the OASIS algorithm Friedrich et al., 2017; (https://github.com/j-friedrich/OASIS), implemented in the aind-ophys-oasis-event-detection capsule (https://github.com/AllenNeuralDynamics/aind-ophys-oasis-event-detection). OASIS performs nonnegative deconvolution of calcium fluorescence traces to infer the underlying spike-related activity, modeling the calcium dynamics as an autoregressive process with an exponential decay kernel. The decay time constant, baseline, and sparsity penalty (Lagrange multiplier for the noise constraint) were automatically estimated from the data’s autocovariance. The outputs include the inferred deconvolved activity (event rates), the denoised fluorescence trace, and the estimated model parameters for each ROI.

SLAP2 dendritic imaging.¶

Post-hoc motion correction¶

Images were generated with an 80 Hz query timebase, and aligned using a custom algorithm (MultiRoiRegistration; AllenNeuralDynamics/GIAnT-MATLAB (2026)), as conventional motion correction algorithms are unstable when correcting motion in thin strip fields of view (Pnevmatikakis & Giovannucci, 2017). Our algorithm was implemented in MATLAB. In brief, the algorithm first initializes a template by using NoRMCorre (Pnevmatikakis & Giovannucci, 2017) to align 42 evenly-spaced frames throughout a trial. Each frame is then aligned against the template by finding the X and Y displacement that yields the maximum cross-correlation among imaged pixels. The pixel values and template values are square root transformed as variance stabilizing technique. Subpixel shifts are determined by fitting a 2-dimensional quadratic to the cross-correlation function around the maximum. With the displacement estimate, a motion corrected frame is generated by linear interpolation, weighted by the freshness of each contributing observation. The motion corrected frame is then averaged into the template to dynamically update it as frames become aligned.

Source extraction¶

Source extraction was performed with a custom algorithm (SILo; AllenNeuralDynamics/GIAnT-MATLAB (2026)), implemented in MATLAB. This algorithm takes advantage of the fact that glutamate release events are spatiotemporally sparse. Specifically, we take inspiration from superresolution localization microscopy methods (Lelek et al., 2021; Chen et al., 2025) to precisely identify source locations despite the reduced effective resolution produced by the integration over pixels. We model a single event as having a spatiotemporal profile of a small Gaussian dot (standard deviation of 1.33 pixels) modulated over time by a decaying exponential of a time constant matched to the glutamate indicator (for iGluSnFR4f we use a decay constant of $\tau = 20$ ms). We perform event detection by convolving this shape with the movie to identify local maxima in space and time (i.e., a 3D matched filter). These events are weighted by their intensity in the filtered movie and aggregated into a summary image, which we term the activity image. The activity image shows localized densities around active sources. We then identify the centroids of these densities by fitting each to a symmetric Gaussian to establish the location of each synapse. As a final step, we use the established source locations as the initialization for constrained non-negative matrix factorization to fine tune the spatial profiles and extract their corresponding time traces.

NWB data packaging¶

Eye tracking¶

At different points in each modality’s respective pipelines, eye tracking information is extracted from the raw behavior videos. A standardized capsule, aind-capsule-eye-tracking (https://github.com/AllenNeuralDynamics/aind-capsule-eye-tracking) was built for fitting ellipses to the pupil, eye (visible perimeter of the eyeball), and corneal reflection of the right eye, based on points tracked using the open source software DeepLabCut (Mathis et al. 2018). DeepLabCut, which uses a pre-trained ResNet 50 deep residual network, was used to track (up to) 12 points along the perimeters of the eye, pupil, and corneal reflection. Ellipses were then fit to the tracking points and the ellipse fit parameters were saved to disk. Validation against hand-annotated ‘ground truth’ frames confirmed that a single ‘universal’ model, trained on a broad selection of data samples, robustly generalized on held-out data across different physiology rigs and individual animals

Synchronized Stimulus Table Generation - Ephys¶

The alignment of stimulus information to the electrophysiology recording is done using the legacy version of the aind-metadata-mapper (https://github.com/AllenNeuralDynamics/aind-metadata-mapper/tree/legacy) requires two input files: a sync file (HDF5, .sync) containing a binary counter sampled at approximately 100 kHz, where each bit encodes the state of a labeled digital line, and a stimulus pickle file (.stim.pkl) containing the intended stimulus parameters and timing as programmed by the stimulus control software (CamStim https://github.com/AllenNeuralDynamics/camstim). The pickle file stores, for each stimulus block, a sweep table of parameter values (e.g., orientation, contrast, spatial frequency), an array of sweep frame indices in stimulus-local coordinates, and a display sequence specifying when each block was presented in seconds relative to session start.

Frame times are extracted from the sync file using two digital lines: the vsync line (stim_vsync), whose falling edges mark nominal monitor frame boundaries, and the photodiode line (stim_photodiode), which records the output of a photodiode affixed to the stimulus monitor that toggles state every 60 frames. Because the photodiode signal reflects actual light output, it captures true frame timing including irregular refresh intervals that the vsync signal does not. The alignment procedure detects all photodiode transitions, trims pulses outside the vsync range, corrects for missing or spurious edges, and then allocates individual frame timestamps within each 60-frame photodiode interval using a trimmed-mean estimate of frame duration. Clock counter rollovers are detected and corrected by adding 2^32 offsets where negative intervals appear. The result is a one-dimensional array of frame times in seconds on the sync clock, one entry per monitor frame, with sub-millisecond precision.

To map stimulus parameters onto these frame times, each stimulus block’s display sequence is converted from seconds to global frame indices using the monitor frame rate and pre-blank duration recorded in the pickle file. The stimulus-local sweep frame indices are then shifted into the global frame domain by apply_display_sequence(). Finally, convert_frames_to_seconds() indexes into the photodiode-corrected frame time array to assign each sweep a start time and stop time in sync-clock seconds. The output is a single stimulus table (saved as a CSV) in which each row corresponds to one stimulus sweep, with columns for start time, stop time, start frame, end frame, stimulus name, and all associated stimulus parameters. Gaps between stimulus blocks are filled with rows labeled as spontaneous activity.

Synchronized Stimulus Table Generation - Mesoscope¶

The mesoscope alignment uses the same, aind-metadata-mapper (https://github.com/AllenNeuralDynamics/aind-metadata-mapper) and the same two input files: a sync file containing digital line states sampled at approximately 100 kHz, and a stimulus pickle file containing the programmed stimulus sweep parameters and timing. The processing diverges from the ephys pipeline in how frame times are derived from the sync data.

Rather than using the photodiode to reconstruct per-frame timing, the mesoscope pipeline takes the falling edges of the vsync line as the base frame times directly. The photodiode is instead used to measure the delay between when the computer issues a vsync signal and when the monitor actually displays the frame. This is done by extract_frame_times_with_delay(), which detects a characteristic three-pulse photodiode pattern marking stimulus onset and offset, then computes, for each photodiode transition in between, the difference between the photodiode rising edge and the nearest vsync falling edge. The median of these per-pulse delays — typically around 35.6 ms — is added uniformly to all vsync-derived frame times. If the measurement is unreliable (standard deviation exceeding 2 ms, or missing data), a hardcoded fallback delay of 0.0356 seconds is applied instead.

Once delay-corrected frame times are established, the mapping from stimulus sweeps to seconds proceeds identically to the ephys case: display sequences are converted to global frame indices, local sweep frames are remapped into the global domain, and frame indices are converted to seconds by lookup in the frame time array. The mesoscope pipeline produces two output tables. The primary table uses the delay-corrected frame times and is appropriate for analyses correlating neural calcium signals with stimulus events, since both the imaging and the stimulus share the same monitor display latency. The secondary table uses raw vsync times without the delay correction, which is useful for analyses tied to hardware trigger signals. Both tables have the same columnar structure: one row per sweep, with start time, stop time, frame indices, stimulus name, and stimulus parameters.

Mesoscope 2-Photon Imaging NWB Packaging Pipeline¶

All processed data were packaged into Neurodata Without Borders (NWB) format Rübel et al., 2022. NWB packaging was performed as an integrated step of the 2-Photon processing pipeline (mentioned above) which produced NWBs in the Zarr format containing the processed 2-Photon data.

A secondary pipeline, also run in nextflow DSL2 on the CodeOcean platform (https://codeocean.allenneuraldynamics.org/capsule/8980147/tree) consolidated all processing outputs together with raw synchronization data and behavioral data into a single NWB file per session. This included several steps run in parallel, whose outputs were merged into the final combined NWB:

• Converting the Pophys processing pipeline’s Zarr NWB to an h5py NWB using a capsule NWB-Zarr-HDMF-Conversion (https://github.com/AllenNeuralDynamics/NWB-Zarr-HDMF-Conversion)

• Generating a synchronized stimulus table that is time-aligned to the recorded ophys timing. Described above under “Synchronized Stimulus Table Generation - Mesoscope” This is then split into several intervals tables based on the type of stimulus shown and packaged into a copy of the ophys NWB.

• Processing the behavior videos of the eye into eye tracked output. Described above in “Eye Tracking”

• Aligning the eye tracking output to the recorded synchronized timing and packaging it into a copy of the ophys NWB, using a capsule aind-eye-tracking-nwb (https://github.com/AllenNeuralDynamics/aind-eye-tracking-nwb)

• Aligning the running speed traces to the recorded synchronized timing and packaging it into a copy of the ophys NWB, using a capsule aind-running-speed-nwb (https://github.com/AllenNeuralDynamics/aind-running-speed-nwb)

• Generating a new data description.json that references the input data, and includes the name of the newly generated data. Aggregating processing.json output from the eye tracking processing and the stim table generation into a new processing.json. Copying the remaining inputted metadata jsons (including procedures.json, quality_control.json, rig.json, session.json, subject.json)

• Finally, merging the resulting NWBs; the stimulus table NWB, the running speed NWB, and the eye tracking NWB, into one complete NWB using aind-nwb-utils (https://github.com/AllenNeuralDynamics/aind-nwb-utils)

• The NWBs were then uploaded to their dandiset using the DANDI command line interface (https://github.com/dandi/dandi-cli)

Neuropixels Ephys NWB Packaging Pipeline¶

The processed data were also packaged into an NWB file during their respective processing pipeline mentioned above. This pipeline included two capsules which were run in sequence, appending processed ecephys information to a base subject NWB. They were aind-ecephys-nwb (https://github.com/AllenNeuralDynamics/aind-ecephys-nwb), which packaged LFP, electrode information, and device metadata, followed by aind-units-nwb (https://github.com/AllenNeuralDynamics/aind-units-nwb) which packaged the units table of the units outputted from kilosort and their associated processed and postprocessed metric into the NWB.

A secondary pipeline, also run on the CodeOcean platform, took the output spike sorted NWB and the raw synchronization and behavior data to produce a complete NWB. This was done by appending data to the NWB in a sequence of processing capsules. This included;

• Converting the Spike sorting processing pipeline’s Zarr NWB to an h5py NWB using a capsule NWB-Zarr-HDMF-Conversion (https://github.com/AllenNeuralDynamics/NWB-Zarr-HDMF-Conversion)

• Using a synchronized stimulus table that is time-aligned to the recorded ecephys timing, this was generated prior to CodeOcean upload performed on Allen Institute rig computers. This process is described above under “Synchronized Stimulus Table Generation - Ephys” This table is then split into several intervals tables based on the type of stimulus shown and packaged into the NWB

• Processing the behavior videos of the eye into eye tracked output. Described above in “Eye Tracking”.

• Aligning the eye tracking output to the recorded synchronized timing and packaging it into the NWB, using a capsule called aind-eye-tracking-nwb (https://github.com/AllenNeuralDynamics/aind-eye-tracking-nwb)

• Aligning the running speed traces to the recorded synchronized timing and packaging it into the NWB, using a capsule called aind-running-speed-nwb (https://github.com/AllenNeuralDynamics/aind-running-speed-nwb)

• The NWBs were then uploaded to their dandiset using the DANDI command line interface (https://github.com/dandi/dandi-cli)