In Vivo Visualization of Spontaneous Activity in Neonatal Mouse Sensory Cortex at a Single-Neuron Resolution

Summary

Primary sensory areas in the neocortex exhibit unique spontaneous activities during development. This article describes how to visualize individual neuron activities and primary sensory areas to analyze area-specific synchronous activities in neonatal mice in vivo.

Abstract

The mammalian brain undergoes dynamic developmental changes at both the cellular and circuit levels throughout prenatal and postnatal periods. Following the discovery of numerous genes contributing to these developmental changes, it is now known that neuronal activity also substantially modulates these processes. In the developing cerebral cortex, neurons exhibit synchronized activity patterns that are specialized to each primary sensory area. These patterns markedly differ from those observed in the mature cortex, emphasizing their role in regulating area-specific developmental processes. Deficiencies in neuronal activity during development can lead to various brain diseases. These findings highlight the need to examine the regulatory mechanisms underlying activity patterns in neuronal development. This paper summarizes a series of protocols to visualize primary sensory areas and neuronal activity in neonatal mice, to image the activity of individual neurons within the cortical subfields using two-photon microscopy in vivo, and to analyze subfield-related activity correlations. We show representative results of patchwork-like synchronous activity within individual barrels in the somatosensory cortex. We also discuss various potential applications and some limitations of this protocol.

Introduction

The cerebral cortex contains several sensory areas with distinct functions. The areas receive inputs originating from their corresponding sensory organs, mostly conveyed through the spinal cord or brainstem and relayed via the thalamus^1,2. Notably, neurons in each primary sensory area exhibit uniquely synchronized activity during early developmental stages, which also originate from sensory organs or the lower nervous centers, but essentially differ from the activities observed in the mature cortex³.

In neonatal rodents, for example, the primary visual area (V1) displays wave-like activity, which originates in the retina (retinal wave) and propagates through the entire visual pathway while conserving retinotopy⁴. The primary auditory area (A1) exhibits synchronous activity organized in band-shaped subregions that correspond to the isofrequency bands in the mature brain. The activity emanates from the cochlea's inner hair cells^5,6. The barrel cortex in the primary somatosensory area (S1) shows a patchwork-like activity pattern in which layer 4 neurons within individual barrels, namely, neurons responsive to individual whiskers, are synchronously activated⁷. Although proposed to originate from the trigeminal ganglion, the source of the activity remains unknown⁷. Consequently, neonatal activity patterns are specialized both within each primary sensory area and within intra-areal subfields. The simultaneous visualization of neuronal activity and structure of primary sensory areas may facilitate an inquiry into the contribution of these activity patterns to the development of sensory systems.

In this article, we summarized a series of protocols: (1) to visualize individual neuronal activities using sparse labeling of GCaMP and primary sensory areas using TCA-RFP mice that express red fluorescent protein in thalamocortical axons⁷, (2) to image single cell-level activity in neonatal mice using two-photon microscopy in vivo, and (3) to analyze the activity correlations within S1 barrel cortex. The representative results show patchwork-like synchronized activity within individual barrels of a postnatal day (P)6 mouse. Despite some limitations, this technique can be used for chronic imaging, wide-field imaging across multiple sensory areas, and various manipulation experiments. The multifaceted analysis of neuronal activity during development will enrich our comprehension of brain circuit formation mechanisms.

Protocol

All the experiments were conducted in accordance with the guidelines for animal experimentation of Kumamoto University and the National Institute of Genetics and approved by the animal experimentation committees.

1. In utero electroporation (IUE)

1. Mate male TCA-RFP mice of ICR background with female wild-type ICR mice. Observe the vaginal plug to check for mating early morning of the next day. Observe the abdomen to check for pregnancy 2 weeks later.
2. Prepare plasmid solution containing 5 ng/µL TRE-nCre, 1 µg/µL CAG-loxP-stop-loxP-GCaMP6s-ires-tTA-WPRE, and 0.02 % Trypan Blue in ddH₂O to sparsely label neurons by GCaMP using the Supernova system⁸.
3. Prepare an analgesic solution containing 0.5 mg/mL carprofen and 0.01 mg/mL buprenorphine. Protect it from light and keep it at room temperature.
4. At embryonic day (E)14, perform IUE as follows. The IUE methods are essentially the same as previous reports^9,10,11.
   1. Wipe the lab bench with 70% ethanol or glutaraldehyde solution. Sterilize all surgical instruments by autoclaving. Wear a mask and a lab coat to reduce the risk of infection to the mouse.
   2. Prepare glass micropipettes using a micropipette puller. Take the plasmid mixture solution into the glass micropipette using an aspirator tube.
   3. Induce anesthesia to the pregnant mice at E14 using isoflurane (2.0% in air). Keep the mice anesthetized during the procedure with isoflurane (1.2% in air). Pinch the toe to ensure that the anesthesia is deep enough. Use ophthalmic ointment to prevent eye dryness.
   4. Disinfect the abdominal area at least three times with alternating rounds of iodine-based scrub and 70% ethanol. Place a sterile drape to cover the surgical area. Make a midline incision and expose the uterus onto the drape.
   5. Repeat dripping warm saline onto the uterus to prevent it from drying or cooling until abdomen closure (Step 1.4.8).
   6. Inject the plasmid solution into one side of the lateral ventricle of embryos one by one using a micropipette and an aspirator tube.
   7. Use a forceps-type electrode to pinch the embryo heads and deliver square electric pulses (40 V, 50 ms) five times at 1 s intervals using an electroporator.
   8. Return the uterus to the abdominal cavity. Apply ~3 mL of warm saline into the cavity. Suture the peritoneal membrane and the abdominal skin. Administer the analgesic solution at 10 μL/g weight (carprofen at 5 μg/g weight and buprenorphine at 0.1 μg/g weight) under the skin at the back of the neck.
5. Place the mouse in a recovery cage on a heating pad (37 °C) to recover from anesthesia. Lay the mouse on its belly to prevent its tongue and saliva from choking the throat. Keep the mouse apart from other mice and monitor it until it recovers to move normally. Return the mouse to the cage.
6. After birth, perform genotyping to check for RFP allele, and euthanize the pups without the allele. This step is recommended, especially if imaging is done during the second postnatal week when the TCA-RFP signal is weak and difficult to check at step 2.2.7.

2. Cranial window surgery

1. Prepare the cortex buffer containing 125 mM NaCl, 5 mM KCl, 10 mM glucose, 10 mM HEPES, 2 mM CaCl₂, and 2 mM MgSO₄ in ddH₂O (pH adjusted to 7.4 with 1 M NaOH)¹² before the day of surgery. Sterile the buffer using a vacuum filter.
   NOTE: The buffer can be kept at 4 °C for up to 3 months. The required volume is 5-10 mL per pup.
2. Perform the following steps at P3-12. Also see the previous reports that have described this procedure^13,14.
   1. Mix 50 mg of agarose with 5 mL of cortex buffer and completely dissolve the agarose by heating. Take some of the solution into a 1.5 mL tube and keep it at 42 °C.
   2. Take some cortex buffer into a 50 mL conical tube and keep it at room temperature. Take some saline into a container (e.g., a cap of a 50 mL conical tube) and keep it at room temperature.
   3. Prepare an analgesic solution containing 0.01 mg/mL buprenorphine. Protect it from light and keep it at room temperature.
   4. Sterilize all surgical instruments by autoclaving. Disinfect the fluorescence stereo microscope with 70% ethanol.
   5. Use an isoflurane vaporizer to induce anesthesia in the pup with isoflurane (2.0% in air). Pay attention to the pup and take it out when its breathing slightly slows down.
      NOTE: If the anesthesia is prolonged, the pup's breathing may cease for a few seconds. Even if the pup's breathing is resumed, prolonged anesthesia may decrease the pup's cerebral blood flow and cause irreversible brain damage.
   6. Disinfect the pup head at least three times with alternating rounds of iodine-based scrub and 70% ethanol. Place the pup onto the heating pad (35 °C) under a fluorescence stereo microscope. Always keep the pup anesthetized with isoflurane (1.5%-2.5% in air).
   7. Select the pups expressing TCA-RFP and GCaMP by observing the fluorescence through the skull. Euthanize the pups that do not express both.
   8. Remove the scalp above the cerebral hemispheres as widely as possible carefully so as not to cause bleeding. Rub the skull with a sterile saline-soaked cotton swab to remove connective tissue.
   9. After the skull dries, adhere the incised scalp surface to the skull with a tissue adhesive.
   10. Transfer the pup onto a 37 °C heating pad to recover from anesthesia. Wait for at least 15 min to let the adhesive solidify.
       NOTE: Pause for up to 1 h before moving on to the next step. Prepare other pups in a similar manner if needed during this time.
   11. Anesthetize the pup with isoflurane. Place the pup onto the heating pad (37 °C) under a fluorescence stereo microscope while anesthetized (1.5%-2.5% isoflurane in air).
       NOTE: If the anesthesia prolongs more than the time for the endpoint (60 min), euthanize the pup by decapitation during anesthetized with isoflurane.
   12. Mark the GCaMP-expressing location on the skull with a sterile permanent marker. Apply cortex buffer onto the location.
   13. Insert the corner of a razor blade into the skull. Push the blade slowly to shave off the skull and make a hole. Pinch off the cracked skull with tweezers and remove it.
   14. Check that the cranial hole is successfully made by observing blood vessels in the hole. If bleeding occurs, quickly rinse the hole with cortex buffer using a micropipette. Repeat rinsing until the bleeding stops completely.
   15. Apply a drop of cortex buffer to the cranial hole and place in a sterile 3-mm-diameter round coverglass over the hole. Wipe away the excess buffer with non-woven fabric. Wait until the area around the glass dries.
   16. Apply warm agarose solution around the glass edge using a micropipette. Because too much agarose may reduce the fluorescent signals, remove the excess solution under the glass by gently pushing the glass from above.
   17. Remove the agarose on the glass or distant from the glass edge using tweezers. Leave agarose only around the outer glass perimeter.
   18. Wait until the agarose solidifies. If agarose shrinkage makes a space under the glass, add agarose solution from the side to cover the entire glass edge. Remove any liquid from the skull surface with non-woven fabric.
   19. Mix acrylic resin powder and liquid in a rubber container. Aspirate the mixture with a micropipette and pour it to cover the agarose surrounding the glass edge with resin.
       NOTE: Because the resin solidifies soon after mixing the powder and liquid, they need to be mixed repeatedly before applying. The amounts needed in each mixing are ~500 µL for liquid and ~0.15 g for powder.
   20. Fix a titanium bar with resin on the contralateral hemisphere. Keep the bar angle parallel to the cover glass. Fix the entire skull surface with resin.
   21. Administer the analgesic solution at 10 μL/g weight (buprenorphine at 0.1 μg/g weight) under the skin at the back of the neck. Return the pup to a 37 °C heating pad for anesthesia recovery. Wait for >60 min for resin solidification before imaging.
       ​NOTE: Pause for 1-5 h before imaging. Perform surgery on other pups in a similar manner during this time.

3. Two-photon calcium imaging

1. Attach a two-axis goniometer with a titanium plate to a stage plate with XY positioning beneath the microscope. Set up a heating pad (35 °C) on the stage.
2. Turn on the scanning software with the following conditions: Pixels, 512 x 512; Bidirectional, ON; Averaging, none; imaging area, 600 x 600 µm with a 20x objective. Set the settings so that the scan rate is faster than 1 Hz.
3. Place the pup on the heating pad, and fix the head-mount titanium bar to the titanium plate with screws. Keep the pup anesthetized by placing a tube port for isoflurane (1.5-2.0% in air).
4. Adjust the window angle horizontally by the goniometer. Turn on the backlight and observe the brain surface with a 5x objective lens, and select the imaging area by XY positioning.
5. Put eye drops on the cranial window. Switch the objective to a 20x water-immersion lens. Observe the cortical surface to confirm that blood flow is seen on the brain surface.
6. Turn off the backlight and scan the brain surface using one-photon mode. Increase the laser power to make the green autofluorescence of the glass and the brain surface visible.
7. Lower the isoflurane concentration to 1.0%-1.5%. Cover the microscope to avoid light leaks. Switch the scanning software to two-photon mode.
8. Adjust the laser power and detector gain appropriate for scanning GCaMP and RFP signals. Find the depth where TCA-RFP signals are seen. Make sure that the depth is layer 4, ~300 µm lower than the brain surface at P6. Select the imaging area where many GCaMP-expressing neurons are seen.
9. Start timelapse imaging of GCaMP and RFP signals. If two-channel simultaneous scanning is inapplicable, capture GCaMP and TCA-RFP images before the imaging.
10. Stop isoflurane to weaken the anesthesia and observe spontaneous activities for ~20 min. Monitor the pup's movements during imaging using an infrared camera. Resume isoflurane anesthesia (2% in air) immediately if any response indicating distress is observed.
11. After the pup stops moving, repeat the imaging from Step 3.9. Change the imaging area if needed.
12. Euthanize the pup with an overdose of isoflurane. Fix the brain by transcardial perfusion of saline and 4% PFA, followed by post-fixation in 4% PFA overnight to prepare tangential slices and perform immunohistochemistry. Otherwise, euthanize the pup with an overdose of anesthetic followed by decapitation.
13. If the dam has no other pups, euthanize the mother mouse with an overdose of anesthetic followed by cervical dislocation.

4. Analysis

1. Start MATLAB and run EZcalcium toolbox¹⁵ to open a graphical user interface (GUI) 'Initial GUI'.
2. Compensate for the image frame drifts due to mouse movements or other causes.
   1. Click Motion Correction in the Initial GUI to open the Motion Correction GUI. Click 'Add Files...' to load a TIF file of the imaging data.
   2. Set the settings as follows: Non-rigid Motion Correction, blank; Upsampling Factor, 50; Max Shift, 15; Initial Batch Size, 200; Bin Width, 200. Click 'Run Motion Correction' to execute the correction. Motion-corrected imaging data will be saved automatically.
      NOTE: The settings should be adjusted to the imaging data properties. If the drifts of some frames are not compensated because of nonlinear frame distortions or movement of the cortex in the depth direction, open the original imaging data without correction in ImageJ Fiji and remove the frames, and restart step 4.2.
3. Detect neurons and draw regions of interest (ROIs).
   1. Click Automated ROI Detection in the Initial GUI to open the ROI Detection GUI. Click Add Files... to load the motion-corrected imaging data.
   2. Set the settings as follows: Initialization, Greedy; Search Method, Ellipse; Deconvolution, Constrained FOOPSI-SPGL1; Autoregression, Decay; Estimated # of ROIs, 60 (more than double the number visually detected is recommended); Estimated ROI Width, 17 (~20 µm); Merge Threshold, 0.95; Fudge Factor, 0.95; Spatial Downsampling, 1; Temp. Downsampling, 1; Temp. Iterations, 5.
      NOTE: The settings should be adjusted to the imaging data properties.
   3. Click Run ROI Detection to execute the detection. ROI data will be saved automatically.
   4. Click ROI Refinement in the Initial GUI to open the ROI Refinement GUI. Click Load Data to load the ROI data. Select the ROIs that had low activity frequency (<1 Hz), located under the skull, or contained other neurons' neurites. Click Exclude ROI to exclude the ROIs from the analysis.
   5. Select the Data Export Format to XLSX and click Export Data to obtain an Excel file with raw dF/F values. The dF/F uses equation (1), where F is the average intensity of pixels in every frame, F0 is the baseline signal intensity, and Fb is the background fluorescence.
           (1)
4. Calculate the Pearson's correlation coefficient of dF/F between ROIs and make a correlation coefficient matrix. Only use dF/F after anesthesia has weakened and spontaneous activity has begun to occur (~10 minutes after stopping isoflurane).
5. Define the barrel edges from the TCA-RFP image using Fiji. Categorize the ROIs to their respective barrels or septa. Compare the pairwise correlation within the same barrel and that between different barrels.
6. Generate 1,000 to 10,000 surrogate data by randomly shuffling the correspondence between ROI and Ca²⁺ traces. Calculate the average correlation coefficient within individual barrels in each surrogate data, and evaluate the statistical significance of the correlation in the real data.
   NOTE: If one of 10,000 surrogates has a value higher than the real value, the statistical significance is 0.0001. The analyses described in steps 4.5 and 4.6 can be conducted for pooled data from multiple animals, as performed elsewhere^7,16.

Results

Figure 1 shows the representative results of layer 4 neuron activities in the barrel cortex of a P6 pup visualized using the present protocol. Two-photon images of the green channel (GCaMP) and red channel (TCA-RFP) were temporally averaged and shown in Figure 1A. Because TCA-RFP fluorescence was much weaker than GCaMP fluorescence, the GCaMP signal leaked into the red channel (Figure 1A1,A2). Fourteen ROIs were dra...

Discussion

Given that the spontaneous activities emerge from the sensory organ or lower nervous system and travel to the primary sensory area through a pathway equivalent to that of a mature nervous system³, it is crucial to define the primary sensory area and the location of imaged neurons within the area. In this protocol, we addressed this requirement by employing transgenic mice that visualize thalamocortical axons and the Supernova system that expresses GCaMP sparsely⁸. These tec...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
20× objective lens (water immersion)
250 mL Vacuum Filter/Storage Bottle System	Corning	431096
4%-paraformaldehyde phosphate buffer solution (4% PFA)	Nacalai	09154-85
Acrylic resin (UNIFAST II)	GC	N/A
Agarose	Sigma	A9793
Aspirator tube assembly	Drummond	2-040-000
CaCl2•2H2O	Nacalai	06731-05
Electroporator	BEX	GEB14
Eye drop (Scopisol)	Senju Pharmaceutical	N/A
Fluorescence stereo microscope	Leica	M165FC
Glucose	Nacalai	16806-25
Heating pad	Muromachi Kikai	FHC-HPS
HEPES	Gibco	15630-080
Isoflurane	Pfizer	N/A
KCl	Nacalai	28514-75
MgSO4•7H2O	Wako	131-00405
Micropipette puller	Narishige	PC-100
Multiphoton laser	Spectra-Physics	Mai Tai eHP DeepSee
Multiphoton microscope	Zeiss	LSM 7MP
NaCl	Nacalai	31320-05
Non-woven fabric (Kimwipe)	Kimberly Clark	S-200
Phosphate buffered saline (PBS)	Nacalai	27575-31
Plasmid: CAG-loxP-STOP-loxP-GCaMP6s-ires-tTA-WPRE	Addgene	pK175
Plasmid: TRE-nCre	Addgene	pK031
Precision calibrated micropipets	Drummond	2-000-050
Razor blade	Feather	FA-10
Rimadyl (50 mg/mL Carprofen)	Zoetis JP	N/A
Round cover glass, 3-mm-diameter	Matsunami	CS01078
Saline	Otsuka	035175315
Sodium pentobarbital	Nacalai	26427-72
Stage for imaging living pup (two single-axis translation stage for XY positioning, two-axis goniometer, base plate, adjustable pillar for z positioning)	ThorLabs	LT1/M, GN2/M, BM2060/M, MLP01/M
TCA-RFP mouse	N/A	N/A	Mizuno et al., 2018a
Tissue adhesive (Vetbond)	3M	1469SB
Titanium bar	Endo Scientific Instrument	N/A	Custom made (Mizuno et al., 2018b)
Titanium bar fixing plate		N/A	Custom made (Mizuno et al., 2018b)
Trypan blue	Sigma	T8154
Tweezers with platinum plate electrode, 5 mm diameter	BEX	CUY650P5
Wild-type ICR mouse	Nihon SLC	Slc:ICR