Guide/Surgery and baseplating

Workflow

Prerequisites — complete these first:

Guide/Experiment Planning for Freely Behaving Animals

Next steps — these build on this guide:

Guide/Animal Handling & Habituation

Review interval	12 months
Status	No reviews recorded yet — needs first review

Overview

Miniscope imaging for freely moving behavior requires three sequential surgical procedures: virus injection, GRIN lens implantation, and baseplate attachment. Each procedure is described in its own section below. If you are using a transgenic animal expressing a calcium indicator, the virus steps will not apply to you.

These procedures have cumulative dependencies. A lens that is well placed but over an area of poor viral expression will yield no usable data. A well-expressed viral population paired with a lens that is off-target by even a few hundred microns may produce nothing. Read the full guide before beginning any surgeries on experimental animals. For guidance on GRIN lens selection, viral construct optimization, and the required viral dilution study, see Guide/Experiment Planning for Freely Behaving Animals first.

Before beginning: Complete the viral dilution study described in Guide/Experiment Planning for freely behaving animals Step 3 before operating on any experimental cohort animals. Beginning surgeries without confirmed optimal titer and expression is the most common cause of cohort-level imaging failure.

For video demonstrations of surgery and baseplating, see the online workshop(https://sites.google.com/metacell.us/miniscope-workshop-2021/home). For a general introduction to rodent stereotaxic surgery, see the JoVE stereotaxic surgery protocol.

Surgical timeline

The table below summarizes typical time and inter-procedure intervals. These are estimates; recovery and expression timelines vary by brain region, viral construct, and animal.

Procedure	Time per mouse	Interval before next step
Virus injection (separate surgery)	~1 hr	3 days - 1 week before lens implant (to allow tissue recovery)
Lens implant	2–3 hrs	4-6 weeks before baseplating (to allow expression)
Virus injection + lens implant (combined, small lenses only)	>3 hrs	4-6 weeks before baseplating (to allow expression)
Baseplating	~1 hr	1–3 days recovery before behavior

This guide is part of the Freely Moving Behavior Workflow. Prerequisites: Guide/Pre-Experiment Checklist. Next step: Guide/Animal Handling & Habituation.

General Surgical Preparation

Equipment

All three surgical procedures require the same core setup.

Item	Notes
Small animal stereotaxic frame	Non-rupture mouse earbars recommended; robotic stereotax (e.g. Neurostar) useful for accurate coordinate targeting and leveling
Isoflurane vaporizer with nose cone and scavenger	Gas anesthesia is strongly preferred over injectable anesthesia for the long duration of these surgeries; ensure scavenger is on and isoflurane tank is filled before starting
Homeothermic monitoring system and surgical heat pad	Maintain body temperature throughout; have a separate recovery heating pad
Stereo surgical microscope with light source	Essential for all steps; aspiration and lens implantation cannot be done accurately without magnification
Microdrill with appropriate burr sizes	0.8 mm burr for skull screw holes; fine burr or trephine for craniotomy
Bead sterilizer	Autoclave all instruments before the session; re-sterilize between animals
Micromanipulator or stereotaxic arm for lens holder	Required for controlled, measured lens lowering
Miniscope V4 stereotaxic holder	For baseplating; mount the Miniscope on the stereotax for precise positioning

For a complete parts list see the master parts list.

Consumables and reagents

Fine forceps (Dumont 5 and Dumont 7 ceramic-coated for lens handling)
Curved Graefe forceps
Vannas spring scissors
Scalpel and blades
Small skull screws (00-96 x 1/16; sterilize before use)
Cyanoacrylate (super) glue plus accelerant/activator (e.g. Vetbond surgical adhesive — strongly recommended over standard cyanoacrylate for lens fixation)
Dental cement
Metabond (recommended as primer layer before dental cement)
Kwik-Sil silicone elastomer (for protecting the lens between surgeries and sessions)
Gel foam absorbable sponge (for bleeding control)
Cortex buffer / ACSF (recipe: 7.888 g NaCl, 0.372 g KCl, 1.192 g HEPES, 0.264 g CaCl₂, 0.204 g MgCl₂ in 1000 ml milliQ water; prepare 3 ml per surgery)
Sterile 0.9% saline (0.5 ml for subcutaneous injection during surgery)
Betadine scrub solution
70% ethanol
Hair removal cream (e.g. Nair) and electric razor
Hydrogen peroxide
Topical lidocaine
Eye lubricant (ophthalmic ointment)
Parafilm and mineral oil (for Nanoject virus loading)
Lens paper and optical cleaning solution
Kimwipes

Pre-operative drug doses

Administer the following 15–30 minutes before the start of surgery:

Drug	Dose	Route	Notes
Carprofen (Rimadyl)	5 mg/kg	Subcutaneous	Analgesic; administer before surgery and every 12–24 hours for 48 hours post-operatively
Dexamethasone	0.2 mg/kg	Subcutaneous	Anti-inflammatory; administer before surgery and every 12–24 hours for 48 hours post-operatively (or longer depending on procedure)
Lidocaine (optional)	5 mg/kg	Subcutaneous at incision site	Topical analgesia at the surgical site; reduces depth of general anesthesia required

Standard skull preparation

The following steps apply to all three surgical procedures.

[IMAGE: Hair removal — photograph showing Nair cream applied to the mouse scalp with a cotton-tip applicator, alongside an electric razor. Shows the scalp before and after hair removal. Source: Crete Workshop slide 17, credit Susie Yu Feng.]

Retrieve animals from the animal facility and allow 30 minutes in the surgery room before starting
Anesthetize with 4–5% isoflurane in the induction chamber; confirm surgical anesthesia with a toe pinch test
Administer pre-operative drugs; apply ophthalmic lubricant generously to both eyes
Check the O₂ tank and replace if pressure is below 200 psi
Place the animal in the stereotaxic frame; maintain at 1–1.5% isoflurane throughout
Secure with ear bars and nose clamp; test skull stability by pressing firmly with a clean cotton-tip applicator — the head must not move
Apply hair removal cream (Nair) to the scalp with a cotton-tip applicator; wait 1–2 minutes and remove thoroughly; follow with an electric razor if needed

[IMAGE: Skull exposure — photograph showing the midline incision made and the skull surface being cleaned with hydrogen peroxide. Bregma and lambda are visible. Source: Crete Workshop slide 18, credit Susie Yu Feng.]

Disinfect the exposed scalp with 3 alternating scrubs of betadine and 70% ethanol
Make a midline incision with a scalpel; for lens implant surgeries, remove a larger skin flap to expose the full skull
Swab hydrogen peroxide over the skull with a cotton-tip applicator to remove connective tissue and clearly expose bregma and lambda
Use a cotton-tip applicator to thoroughly remove all periosteum from the skull surface

[IMAGE: Skull scoring — photograph showing forceps scoring the cleaned skull surface in a crosshatch/grid pattern. Source: Crete Workshop slide 19, credit Susie Yu Feng.]

Score the skull surface in a grid pattern with forceps to increase surface area for dental cement (avoid sutures and any prior injection sites)
Detach neck muscle from the posterior skull edge (ear to ear) using a scalpel with the blade kept flush with the skull — this prevents muscle pull on the implant and reduces later muscle growth that can destabilize the headcap; do not push the blade downward

[IMAGE: Skin gluing — two photographs showing cyanoacrylate being applied along the perimeter of the scalp incision to glue the skin edge flat against the skull. The second photo shows the completed glued edge. Caption note: "Make sure perimeter of incision is completely dry before applying glue." Source: Crete Workshop slides 20–21, credit Susie Yu Feng.]

Apply a thin bead of cyanoacrylate glue along the full perimeter of the incision to adhere the skin edge to the skull — ensure the perimeter is completely dry before applying glue; this step prevents later skin ingrowth under the headcap

[IMAGE: Completed skull preparation — photograph showing the fully prepped skull with scored surface, perimeter skin glued down, clean exposed bone, and landmarks visible. Source: Crete Workshop slide 22, credit Susie Yu Feng.]

Level the skull: if using a Neurostar robotic stereotax, open the software and confirm that stereotax axis readings align with the software display before proceeding; measure depth at bregma and lambda and adjust until the difference is less than 0.1 mm in the A-P axis; confirm M-L leveling by measuring at equal distances left and right of the midline

Critical: Connective tissue, muscle, skin, and fur must not contact the dental cement that will form the headcap. Any tissue contact is the primary cause of headcap failure and loosening, which increases motion artifact and risks losing the implant entirely.

Building an aspirator

An aspirator is required for tissue removal during GRIN lens implantation when using larger diameter lenses or approaching cortical-depth targets. A simple aspirator can be built from: a vacuum line or pump, a liquid trap flask, a 1 ml syringe with a side hole for suction control by finger, blunt needles (27G and 30G), tubing, and connectors.

[IMAGE: Aspirator diagram — schematic illustration of the full aspirator assembly showing the vacuum filter flask, tubing, the 1 ml syringe with the side-hole suction control, and the needle at the end. Source: Crete Workshop slide 25, credit Susie Yu Feng.]

[IMAGE: Needle size comparison — side-by-side diagram of a 27G and 30G blunt needle, showing the relative sizes used at different aspiration stages. Source: Crete Workshop slide 25.]

Control suction pressure by partially covering the side hole with a finger. Test suction by briefly approaching the saline bubble over the skull — large and immediate changes in volume indicate excess pressure; reduce before approaching tissue.

Building a GRIN lens holder

A vacuum-based lens holder provides secure control during implantation and releases the lens without mechanical force when suction is released.

Take two 1 ml micropipette tips
Cut the first so its tip opening is just slightly larger than the diameter of your GRIN lens
Cut the second so its opening is just slightly smaller than the lens diameter
Insert the smaller tip into the larger tip
Connect the assembly to a 1 ml syringe attached to the vacuum line
Fix the holder to the stereotaxic arm with tape or a clamp

[IMAGE (optional): 3D-printed lens holder — photograph or render of the 3D-printed stereotaxic lens holder, showing the tip with a smaller inner diameter that prevents the lens from being accidentally sucked up into the holder. Caption: "3D-printed lens holder. Inner tip diameter is slightly smaller than the lens." Source: Crete Workshop slide 30.]

Commercial lens holders are available for different lens sizes (e.g. KORF Model 1770 for 0.5 mm, Thorlabs XCL for 1 mm). The Golshani Lab stereotaxic holders repository includes designs compatible with V4 lenses.

GRIN lens selection by brain region

[IMAGE: Lens implant regions diagram — three-panel histology image showing example GRIN lens placements in cross-section for three different brain regions, one with a 1 mm lens, one with a 0.5 mm lens, and one with a 1 mm lens. Each panel is annotated showing the lens positioned ~150–200 µm above the viral injection site. Caption: "Example lens implants in different brain regions. Lens implanted ~150–200 µm above viral injection. Lenses can be reused." Source: Crete Workshop slide 13/14.]

Brain region	Suggested lens (diameter / length)	Notes
dCA1	1.8 mm / 4.3 mm (UCLA V3) or 1 mm / 4 mm (UCLA V4)	Full aspiration required; corpus callosum serves as clear aspiration landmark
CA2	1 mm / 4 mm	Angled approach (30° away from midline) required; craniotomy must be enlarged accordingly (craniotomy ~2.0–3.0 mm ML to accommodate angle)
DG (dentate gyrus)	0.5 mm / 6.1 mm	No aspiration required; make track with sharpened 27G blunt needle
vCA1	0.5 mm / 6.1 mm	No aspiration required
mPFC (prelimbic, PrL)	1 mm / 4 mm	Full aspiration required
IL, DP	0.5 mm / 6.1 mm	No aspiration required
NAc	0.6 mm / 7.3 mm	Partial aspiration recommended
VTA	0.6 mm / 7.3 mm	Dopaminergic cells are sensitive to implant; expect higher attrition in this region
VMH	0.5 mm / 8.4 mm	No aspiration required

The lens should be long enough to reach the target area while leaving 2–4 mm above the skull post-implantation. The focal plane of standard GRIN lenses sits approximately 150–200 µm below the bottom surface of the lens — position the lens 150–200 µm above the GCaMP-expressing cell population.

Note on lenses: GRIN lenses can be reused across animals after cleaning. See the Guide/Pre-Experiment Checklist for lens cleaning and inspection steps.

Stereotaxic coordinates

The table below gives Golshani Lab coordinates for hippocampal targets. All coordinates in mm; DV is measured from skull surface. These are starting points — verify and adjust based on histology from your first cohort.

Brain region	AP		ML		DV		Notes
Brain region	Virus	Lens	Virus	Lens	Virus	Lens	Notes
CA2	−1.8	−1.8	−1.9	Craniotomy −2.0 to −3.0	−1.7	−1.7	30° angle away from midline; large craniotomy required to accommodate angle
dCA1	−2.1	−2.1	2.0	1.8	−1.65	−1.25	Full aspiration to corpus callosum; target lens depth 1.35 mm below skull
DG	−1.8	—	−0.75	—	−2.25	—	Virus injection coordinates only; no lens implant coordinates currently listed

These coordinates are specific to the Golshani Lab mouse colony. Coordinates vary across colonies due to differences in animal age, weight, and skull geometry. Always verify lens placement relative to expressing cells by post-hoc histology on your first cohort before committing to full-scale surgeries.

Aspiration strategy by lens size

Lens diameter	Aspiration approach	Rationale
1.8 mm or 1 mm	Full aspiration required	Lens creates too much tissue pressure if pushed through intact cortex
0.6 mm at cortical depths	Partial aspiration recommended	Reduces out-of-focus background fluorescence above the lens
0.5 mm or 0.6 mm at depths > 2 mm	No aspiration; sharp needle tract	Cavity too narrow to visualize and clear safely

Part 1 — Virus Injection

When to perform

For 1.8 mm or 1 mm diameter lenses, perform virus injection as a separate surgery 4–7 days before lens implantation. This prevents aspiration of recently injected virus before it has had time to diffuse into tissue.

For 0.5 mm or 0.6 mm diameter lenses, virus injection and lens implantation can be performed in a single combined surgery. Inject virus, wait 30–60 minutes, then proceed to lens implantation. Performing both in one session ensures consistent skull leveling and landmark identification, which simplifies accurate targeting and reduces animal use.

Virus injection procedure

After completing standard skull preparation:

Mark the injection site by moving the leveling device or drill to the target AP and ML coordinates and marking with a fine-tip pen
Drill a small hole at the injection coordinates using a fine drill bit; penetrate the skull but do not pierce the dura — piercing the dura will cause bleeding
Prepare the Nanoject syringe pump in advance (Nanoject manual): pull a glass capillary and slowly fill with mineral oil ensuring no air bubbles; load the capillary onto the Nanoject metal needle and tighten securely
Place a strip of Parafilm over the ear bar and flatten; pipette approximately twice the required volume of virus onto the Parafilm
Aspirate virus into the capillary at 1000 nl/min; ensure no bubbles enter; mark the oil-virus interface in the capillary
Mount the Nanoject on the stereotaxic arm; zero at bregma; move to target AP and ML coordinates
Lower the needle tip to skull level at the craniotomy and record the DV coordinate
Lower slowly at 0.1 mm/sec to the target DV; for large injections (≥1 µl), lower an additional 0.1 mm past target then raise back — this creates space and reduces pressure-driven backflow
Inject at 1 nl/sec
After injection is complete, wait 10 minutes before withdrawing — do not withdraw immediately
Withdraw slowly at 0.1 mm/sec
Inject 0.5 ml sterile saline subcutaneously

Virus handling: Dispose of everything that contacts virus as biohazard waste in bleach. Place the bleach-containing waste bottle in the biohazard bin. Follow your institutional biosafety guidelines.

For the injection site offset relative to the lens implant site, see Guide/Experiment Planning for freely behaving animals — for most preparations we recommend injecting 200–250 µm lateral to the planned lens center to avoid imaging damage from the injection tract.

Post-operative care (virus injection only)

Return the animal to a clean, warmed recovery cage on a heating pad until ambulatory
Carprofen (5 mg/kg s.c.) every 12–24 hours for 48 hours
Dexamethasone (0.2 mg/kg s.c.) every 12–24 hours for 48 hours
Amoxicillin (0.25 mg/ml in drinking water) for 7 days; or TMS (1 ml in 100 ml drinking water) for 7 days as an alternative; replace with normal water after 7 days
Complete the "Investigator Will" card or equivalent post-operative monitoring documentation per your institution's requirements
Monitor daily for signs of infection or distress

Part 2 — GRIN Lens Implantation

Surgical preparation

Perform the standard skull preparation steps above. If this is a second surgery after a prior virus injection, reopen the original incision and clean the skull surface before proceeding.

Drill a skull screw hole in a region where the lens will not be placed — typically the left posterior skull. Drill and size the hole to fit the anchor screw snugly; sterilize the screw; insert with forceps and screwdriver. The screw should not protrude above the final height of the implanted lens above the skull.

Clear all bone debris from screw drilling before beginning the craniotomy.

Craniotomy

[IMAGE: Craniotomy diagram — schematic top-down illustration showing four small guide holes drilled in a circle and connected to outline a circular craniotomy. Shows the progression from four holes to a complete circular opening. Source: Crete Workshop slide 23, credit Susie Yu Feng.]

[IMAGE: Craniotomy photo — photograph through the surgical microscope showing a completed craniotomy with the dura visible, irrigated with cortex buffer. Source: Crete Workshop slide 24, credit Susie Yu Feng.]

Irrigate the skull with cortex buffer before drilling to prevent tissue overheating
Drill and connect four guide holes to outline a circular craniotomy of sufficient diameter to accommodate the lens — the craniotomy must be at least as large as the lens diameter; for CA2 angled approaches, the craniotomy must cover ~2.0–3.0 mm ML to accommodate the 30° angle
Thin the skull until it is flexible but intact; do not drill through at this stage
Use a bent 27G needle or Bonn micro-probe to break through the remaining thin skull and loosen the skull cap
Immediately apply cortex buffer to the exposed tissue
Remove any remaining small bone fragments with a bent needle or micro-curette
Perform durotomy: use the bent needle to pierce the dura and slowly peel it back; dura has a yellow tint compared with white brain tissue underneath
Irrigate immediately after durotomy and continue irrigating throughout

If bleeding occurs during or after durotomy: Apply a saline-soaked piece of gel foam and wait. Do not proceed until bleeding has completely stopped. Blood clots under the lens will occlude the FOV permanently. If bleeding cannot be controlled, do not implant — allow recovery and reassess.

Tissue aspiration

Full aspiration (1.8 mm and 1 mm lenses)

[IMAGE: Active aspiration — photograph through the surgical microscope showing a 27G blunt needle inside the craniotomy aspirating cortex, with cortex buffer being continuously applied. Source: Crete Workshop slide 26, credit Susie Yu Feng.]

Attach a blunt 27G needle bent at ~45° to the vacuum line; test suction by approaching the saline bubble — volume changes should not be immediately visible
Slowly lower the needle toward tissue while continuously irrigating with cortex buffer; aspirate in slow circular motions
Some bleeding is normal — adjust irrigation to maintain visibility and prevent large clot formation; gel foam can control bleeding at cavity edges

[IMAGE: Corpus callosum landmark — photograph through the surgical microscope showing the corpus callosum fibers visible at the base of the aspirated cavity. The image should show the banded fiber structure. Caption: "Look for horizontal and diagonal bands of the corpus callosum. Stop when vertical fibers appear." Source: Crete Workshop slide 27, credit Susie Yu Feng. This is one of the most important images in the guide — the CC landmark is difficult to describe in words alone.]

For dCA1 and mPFC: Aspirate with the 27G needle, watching for the corpus callosum. The CC appears in three sequential stages as you approach correct depth:
Stage 1 — Horizontal striations: Fibers running left-to-right across the field. Switch to a 30G blunt needle at this point.

Stage 2 — Diagonal striations: Fibers running at an angle; continue careful aspiration.

Stage 3 — Vertical striations: STOP. Vertical fibers indicate you are at the correct depth. Do not aspirate further — disturbing the vertical fibers causes irreversible tissue damage.

For mPFC: aspirate ~1 mm depth with 27G then switch to 30G.
Wait 10 minutes to confirm complete hemostasis before placing the lens — do not place a lens into a field that is still bleeding

Partial aspiration (0.6 mm lenses at cortical depths)

Mark 1 mm from the tip of a 27G blunt needle; aspirate to 1 mm depth
Switch to a 30G needle marked at approximately half the planned implant depth; continue to that depth
Use a sharpened needle to make a tract and retract; wash and clear the cavity until bleeding stops

No aspiration (0.5 mm and 0.6 mm, deep targets)

Use a sharpened needle lowered to target depth to create a tract, then retract slowly
Wash the drill hole with cortex buffer and aspirate fluid until no active bleeding

Lens implantation

[IMAGE: Lens insertion — photograph through the surgical microscope showing the GRIN lens being lowered into the aspirated cavity on the vacuum holder, with the lens tip approaching the target depth. Source: Crete Workshop slide 31, credit Susie Yu Feng.]

Place the GRIN lens on the lens cloth from the optical cleaning kit; use heat-shrink-covered forceps or the lens holder to handle it — never touch the optical surfaces with unprotected metal
Use compressed air to remove dust from lens surfaces
Secure the lens in the holder on the stereotaxic arm; visually confirm it is level before lowering
Zero the lens above bregma without touching the skull; move to the target coordinates
Confirm all bleeding has stopped before lowering
Lower the lens at a constant rate of 100–200 µm/min in 10–20 µm steps — 100 µm/min is preferred; slower lowering gives tissue time to part rather than compress
Target depth: 150–200 µm above the GCaMP-expressing cell population (for dCA1, target depth is typically 1.35 mm below skull surface; see coordinates table above)
Brain tissue and small vasculature should remain visible under the lens throughout lowering

Critical: Lower the lens slowly. Rapid insertion is the primary cause of excessive tissue damage, autofluorescence, and loss of dynamic signal. If unexpected bleeding occurs during lowering, stop and wait — do not continue through active bleeding.

Securing the lens

[IMAGE: Removing the lens holder — photograph showing suction being released and the lens holder being carefully lifted away from the implanted lens, which remains seated in the craniotomy held in place by cyanoacrylate. Source: Crete Workshop slide 32, credit Susie Yu Feng.]

[IMAGE: Sealing with glue — photograph showing cyanoacrylate being applied around the base of the lens and the skull surface. Source: Crete Workshop slide 33, credit Susie Yu Feng.]

[IMAGE: Applying dental cement — photograph showing dental cement being built up around the lens base and over the anchor screw, covering the full skull surface. Source: Crete Workshop slide 34, credit Susie Yu Feng.]

[IMAGE: Applying Kwik-Sil — photograph showing the completed headcap with Kwik-Sil silicone elastomer applied over the top of the GRIN lens to protect it. Source: Crete Workshop slide 35, credit Susie Yu Feng.]

Wick away excess cortex buffer from around the lens base using a Kimwipe — do not apply the Kimwipe directly to the lens
Apply a small ring of cyanoacrylate or surgical adhesive (Vetbond strongly recommended) around the lens base and skull; avoid contact with the lens holder; allow to cure fully (~5 minutes) before proceeding
Apply cyanoacrylate accelerant to speed curing if needed
Release suction and carefully remove the lens holder — the lens should not move
Apply additional super glue around the lens-skull junction, then cover the entire exposed skull area with dental cement; cover the anchor screw
Seal the cement edges with a thin additional layer of super glue
Apply Kwik-Sil over the top of the lens to protect it
Smooth all headcap edges; no cement should contact skin or muscle

Post-operative care

Same regimen as for virus injection surgery above. Allow 3–6 weeks for expression and tissue recovery before attempting baseplating. The exact window depends on your construct and target region — confirm this with your expression window from the dilution study. Do not attempt baseplating until dynamic calcium activity is verified through the lens.

Lens implant troubleshooting

Issue	Solutions
Kwik-Sil keeps coming off / lens is scratched or damaged	Consider an alternative lens protector design. Ensure Kwik-Sil is applied in a thin, even layer and cured fully before the animal recovers. Mechanical damage during recovery is common if Kwik-Sil is too thick or poorly adhered.
Lens is loose or comes out	Make a smaller craniotomy — a tight fit between the craniotomy edge and the lens base provides mechanical support for the glue. Ensure skull surface is completely dry and free of periosteum before applying cyanoacrylate. Confirm the anchor screw is seated fully.
Too much bleeding during aspiration	Slow down irrigation and aspiration pace. Bleeding obscuring the CC landmark is common — irrigate constantly and wait for hemostasis before proceeding. If bleeding cannot be controlled, close and allow recovery.
Unable to visualize horizontal or vertical CC bands	Re-configure the microscope to be more zoomed in and positioned directly over the craniotomy center. Go slower — aspirating too quickly removes too much tissue per pass to reliably identify the bands. If coordinates may be off, adjust ML or DV slightly and verify by post-hoc histology.
No imaging signal after verified lens placement	May need to try different coordinates to avoid sutures or adjacent structures. Perfuse and assess histology to confirm lens position relative to expressing cells. Adjust DV for remaining cohort.

Important tips

Keep isoflurane as low as possible; surgeries take 2–6 hours and prolonged deep anesthesia delays recovery
Infections destroy imaging quality and the ability to track the same cells across sessions — maintain strict sterility
Depth is trial and error for new brain regions; if the first few animals do not yield good images, verify lens placement relative to expressing cells by post-hoc histology and adjust DV coordinates accordingly
If the lens is placed too far from the expressing population in any axis, cells will not be in the focal plane — histology is the only reliable way to diagnose this

Part 3 — Checking Expression and Attaching the Baseplate

When to proceed

Do not attach the baseplate until you can confirm through the GRIN lens that:

Distinct GCaMP-expressing cell bodies are clearly visible
Dynamic calcium transients are present — not static fluorescence
Vasculature is visible and can serve as a landmark for across-session FOV alignment

Typically 5–6 weeks after virus injection for most brain regions; as early as 2–3 weeks for superficial cortical targets; up to 8–12 weeks for some deep brain regions. If cells are not yet distinct at first check, reapply Kwik-Sil and recheck 2–3 weeks later.

Do not baseplate if the fluorescence signal is static (no dynamic activity), consists of spherical bright spots without the honeycomb appearance of healthy GCaMP expression (indicating tissue damage or cell death), or the field of view is completely dark (possible blood clot). Perfuse and assess histology before deciding whether to adjust coordinates for the remaining cohort.

Checking expression (V4)

Anesthetize the animal and secure in the stereotaxic frame
Carefully remove the Kwik-Sil from the top of the GRIN lens with blunt forceps — do not scratch the lens
Drill away any dental cement that is obstructing the area around the lens top; this may be needed to allow the baseplate to sit flat
If residue (Kwik-Sil, dental cement) is present on the lens surface, gently remove with a Kimwipe and a small amount of acetone; wipe laterally only, never apply downward pressure
Clear any remaining debris from the lens with compressed air
Connect the Miniscope V4 assembly to the DAQ and cable; run the Miniscope DAQ software on the recording computer and confirm the Miniscope is detected
Screw the baseplate onto the bottom of the Miniscope, ensuring it sits flush
Mount the Miniscope and holder onto the stereotaxic arm with the baseplate screw facing toward the rear of the mouse — this orientation simplifies future removal and reattachment during behavioral sessions
Visually align the Miniscope over the lens; begin with the Miniscope approximately 3–4 mm above the lens top
Turn on the LED; set gain to 2–3 rather than maximizing LED brightness — this preserves LED longevity while maintaining visibility. Typically: gain 2 and LED 30–60% is a good starting range
Ensure the electrotunable lens (ETL) is set to 0 at this stage
Slowly lower the Miniscope toward the lens using the stereotaxic arm until cells come into focus; take time adjusting physical height for optimal cell visibility — do not adjust the ETL during this process

Troubleshooting — unable to see cells: Virus may need more time to express (2–3 weeks minimum). Try increasing the LED (usually higher than you expect). Some neurons are not active under anesthesia — a gentle tail pinch can sometimes stimulate activity. If virus appears bleached during the procedure, expression typically recovers over a few days; recheck before deciding to abandon baseplating.

Baseplating procedure (V4)

[IMAGE: Miniscope holder setup — photograph showing the miniscope mounted in its stereotaxic holder, positioned above the animal. This is the "sturdy holder" setup referenced throughout the baseplating protocol. Source: Crete Workshop slide 38.]

[IMAGE: Baseplating overview — photograph showing the full baseplating setup with the Miniscope lowered toward the GRIN lens implant on an anesthetized mouse in the stereotaxic frame. Source: Crete Workshop slide 39.]

[VIDEO: Baseplating tutorial — embed the YouTube video at https://youtu.be/GoDJGfqO3po showing the full V4 baseplating procedure. This is the primary step-by-step video reference for this section.]

See also the Aharoni group baseplating protocol and video tutorial for an additional reference.

Tip: Practice removing and reattaching the Miniscope to find the same field of view several times before applying any cement. You should be able to reliably return to the same cells and vasculature pattern before committing.

Once the Miniscope is correctly positioned with cells clearly visible, turn the LED off
Mix dental cement to a relatively viscous consistency — thick enough to shape, not runny
Apply dental cement around the baseplate; ensure enough cement fully covers all gaps between the baseplate and the skull to minimize light contamination. Build up gradually rather than all at once
Every few minutes during cementing, turn the LED on to verify the field of view has not shifted
Ensure no cement contacts the Miniscope assembly itself
Once all gaps are filled and the baseplate is fully covered on all sides, allow the cement to harden completely — typically 15–20 minutes
Periodically turn the LED on during curing to confirm the field of view remains stable throughout
Once cement is fully hardened, loosen the baseplate screw with the hex key
Gently pull the Miniscope upward while applying light downward pressure to the animal's head — do not pull at an angle
Attach the protective cap to the baseplate
Allow the animal to recover for 2–3 days before beginning handling and habituation sessions

Critical: Cement that is too runny can flow onto the lens imaging surface. Cement that is too solid will not adhere properly, causing movement between sessions. Improper baseplate attachment is the leading cause of motion artifact across recording sessions.

Baseplating troubleshooting

Issue	Solutions
Unable to see cells	Virus may need more time (2–3 weeks minimum). Increase LED brightness (usually higher than expected). Some neurons are not active under anesthesia — a tail pinch can sometimes stimulate activity.
Cell visibility changes over the course of baseplating surgery	Virus may bleach transiently during the procedure; expression will recover over a few days. As dental cement dries it can slightly shift baseplate placement — check the FOV every few minutes during cementing and curing.
Baseplate falls off or comes loose when attaching scope	The skull scoring, set screw, and perimeter skin glue steps in skull preparation are critical for long-term adhesion. Confirm the baseplate screw is tightened securely before removing the Miniscope. If the baseplate repeatedly fails, verify no tissue or moisture was present at the cement interface.
FOV during behavior differs from baseplating FOV	Cement shrinkage as it dries can shift the baseplate — check the FOV frequently during curing. Experimental conditions (e.g. water deprivation, stress) can change the ETL focal depth; use the ETL range to recover the target FOV rather than re-baseplating.

Relay baseplating for thin lenses

[VIDEO: Relay baseplating — embed the YouTube video at https://youtu.be/b-LrnOBX3j8 demonstrating the relay baseplating technique for thin (0.5 mm or 0.6 mm) lenses, including the option to cement the objective lens directly to the headcap.]

When using a thin relay lens (0.5 mm or 0.6 mm) in combination with a separate objective GRIN lens, two options exist:

Option 1: Secure the objective lens into the Miniscope and baseplate as described above. The objective lens must be glued very securely — any movement changes the focal plane across sessions.
Option 2 (recommended for stable longitudinal cell tracking): Cement the objective lens directly onto the animal's headcap during baseplating, rather than mounting it in the Miniscope. This produces more stable recordings of the same cells across weeks. See the relay baseplating video above.

Expression outcome guide

What you observe	Action
Low fluorescence but dynamic activity at early timepoint (2–3 weeks)	Reapply Kwik-Sil; recheck at 5–6 weeks. Early timepoints often show low brightness that improves with continued expression.
Distinct cell bodies with clear dynamic transients	Proceed with baseplating.
Static fluorescence; spherical bright spots; no transients	Likely autofluorescence from tissue damage or cell death. Perfuse and assess histology. Do not baseplate.
Black visual field	Possible blood clot occluding the lens. Perfuse and examine tissue under the lens.
Dynamic activity absent under anesthesia	Try stimulating activity with a gentle tail pinch or loud stimulus. Deep isoflurane can suppress activity in some populations — briefly reducing anesthesia depth may reveal activity.
No detectable signal after 8–12 weeks	Perfuse and check lens placement relative to expressing cells by histology. Adjust DV coordinates for remaining cohort animals.

Post-surgical monitoring

Monitor all animals daily from surgery through the end of the experiment. Criteria for immediate veterinary contact and potential euthanasia:

Any sign of infection at the surgical site (discharge, redness, swelling)
Weight loss greater than 20% of pre-surgical body weight that does not resolve
Signs of pain or distress not resolved by the analgesic protocol
Visible damage, cracking, or displacement of the GRIN lens or baseplate

Do not attempt field re-cementation of a loose baseplate. Contact your veterinary staff.

References / Further Reading

Resendez SL, Jennings JH, Ung RL, Namboodiri VMK, Zhou ZC, Otis JM, Nomura H, McHenry JA, Kosyk O, Stuber GD. (2016) Visualization of cortical, subcortical and deep brain neural circuit dynamics during naturalistic mammalian behavior with head-mounted microscopes and chronically implanted lenses. Nat Protoc 11(3):566–597. https://doi.org/10.1038/nprot.2016.021
Zhao P, Aharoni D, Golshani P. (2025) GRIN lens implantation strategies for in vivo calcium imaging using miniature microscopy. PLoS One 20(5):e0323256. https://doi.org/10.1371/journal.pone.0323256 — UCLA Miniscope group protocol covering lens selection by brain region, three aspiration strategies (full, partial, none) by lens diameter, multi-lens implantation for bilateral mPFC or mPFC + NAc, and post-operative care. Full step-by-step protocol also available on protocols.io: https://doi.org/10.17504/protocols.io.ewov12jyogr2/v1
Thapa R, Liang B, Liu R, Li Y. (2021) Stereotaxic Viral Injection and Gradient-Index Lens Implantation for Deep Brain In Vivo Calcium Imaging. J Vis Exp (176):e63049. https://doi.org/10.3791/63049
Aharoni D, Hoogland TM. (2019) Circuit Investigations With Open-Source Miniaturized Microscopes: Past, Present and Future. Front Cell Neurosci 13:141. https://doi.org/10.3389/fncel.2019.00141
UCLA Miniscope Project — Online Workshop. https://sites.google.com/metacell.us/miniscope-workshop-2021/home — Video recordings of the 2021 Virtual Miniscope Workshop (organized by Daniel Aharoni, Denise Cai, and Tristan Shuman), covering Miniscope hardware, surgery and baseplating, animal behavior, and data analysis.