Protocol for investigating astrocytic mitochondria in neurons of adult mice using two-photon microscopy

Summary

Under pathological conditions, astrocytes can transfer mitochondria to neurons, where they exert neuroprotective effects. In this context, we present a protocol for capturing astrocytic mitochondria in neurons of adult mice using a two-photon microscope. We describe an approach for constructing a mouse model with combined labeling of astrocytic mitochondria and neurons. We then detail procedures for the preparation of a coverslip with a customized titanium ring and cranial window for two-photon microscopy scanning.

For complete details on the use and execution of this protocol, please refer to Zhou et al. ¹

Graphical abstract

Highlights

• •Steps for astrocytic mitochondria labeling via PhAM^floxed mice
• •Instruction for AAV-based neuronal labeling
• •Procedures for preparing optimal cranial window
• •Guidance for intravital imaging of astrocytic mitochondria in neurons

Publisher’s note: Undertaking any experimental protocol requires adherence to local institutional guidelines for laboratory safety and ethics.

Under pathological conditions, astrocytes can transfer mitochondria to neurons, where they exert neuroprotective effects. In this context, we present a protocol for capturing astrocytic mitochondria in neurons of adult mice using a two-photon microscope. We describe an approach for constructing a mouse model with combined labeling of astrocytic mitochondria and neurons. We then detail procedures for the preparation of a coverslip with a customized titanium ring and cranial window for two-photon microscopy scanning.

Before you begin

The protocol described here outlines the construction and imaging a model of astrocytic mitochondria transfer to neurons in vivo. Additionally, it is worth noting that the protocol is adaptable for cerebral vascular observation, brain blood flow monitoring, calcium signal measurement and continuous evaluation of newly formed tissue growth.

Institutional permissions

All animal studies were approved by the Southwest Medical University Biomedical Ethics Committee in Sichuan, China (Protocol# SYXK(Chuan) 2018-065). All experimental procedures were performed in accordance with the National Institutes of Health Guidelines for the Care and Use of Laboratory Animals. Any experiments involving animals require prior approval from the appropriate animal ethics committee at your research institution.

Adeno-associated virus transfection

Timing: 10 weeks

This section describes stable mouse model construction using adeno-associated virus (AAV).

• 1.
  Preparation and identification of PhAM^floxed mice (Figures 1A and 1B).

  Note: PhAM^floxed (photo-activatable mitochondria) mice are genetically engineered to express a mitochondrial-targeted version of Dendra2, a green/red photoswitchable monomeric fluorescent protein. In these mice, Cre-mediated excision of a floxed stop sequence enables the expression of green fluorescence specifically in mitochondria. When exposed to 405 nm laser light, the green fluorescence in the mitochondria is irreversibly converted to red, allowing for the assessment of mitochondrial fusion and transport. More detail information could find in the ref. ² PhAM^floxed mice are purchased from Jackson Laboratory.

  • a.
    House mice in a vivarium under a 12-h light/dark cycle at a temperature of 22°C, with unrestricted access to food and water.
  • b.
    Randomly assign animals to each treatment group and perform all experiments under standard laboratory procedures of randomization and blinding.
  • c.
    Identify the genotype of PhAM^floxed mice by PCR using mouse tail DNA.

    • i.
      Extract genomic DNA with a DNA isolation kit (DC102, Vazyme Biotech, China).
    • ii.
      Amplify DNA with a PCR amplification mix (P112, Vazyme Biotech, China).
    • iii.
      Perform electrophoresis using a 1% agarose gel with a DNA marker (DL5000, Vazyme Biotech, China).

      Optional: Primers used for genotyping were listed in Table 1. Expected results are mutant (203 bp), heterozygote (203 bp and 361 bp) and wild type (361 bp). Mice of both mutant and heterozygous genotypes are suitable for subsequent AAV administration.

• 2.
  Injection of AAV.

  • a.
    Preparation of AAV.

    Note: AAVs are obtained from OBiO Technology.

    • i.
      Use an AAV vector carrying a Cre recombinase transgene under control of the astrocyte-specific GfaABC1D promoter (AAV2/PHP.eB-GfaABC1D-Cre) to catalyze recombination at loxP sites, enabling green fluorescent labeling of mitochondria in astrocytes.
    • ii.
      Identify neurons by red fluorescence after transduction with an AAV vector expressing tdTomato under the synapsin promoter (AAV2/PHP.eB-Syn-tdTomato).

      Optional: AAV vectors can be obtained commercially or produced in the laboratory. The key components for generating the desired AAV constructs include the Cre recombinase coding sequence and the tdTomato fluorescent reporter gene are detailed in Table 2.

  • b.
    Injection of AAV.

    • i.
      Place mice in an anesthesia induction chamber to induce anesthesia with isoflurane at 3 mL/h and to maintain anesthesia at 1.5 mL/h.
    • ii.
      Use proparacaine hydrochloride eye drops (0.5% proparacaine hydrochloride ophthalmic solution, Alcon, USA) to accomplish ophthalmic anesthesia.
    • iii.
      Use 29-gauge insulin syringes (Ultra-Fine Insulin Syringe, BD, USA) to finish the injection.

      CRITICAL: To determine the optimal transfection titer of different vectors, it is better to administer a concentration gradient of AAV particles, followed by immunofluorescence analysis of endogenous fluorescence co-staining with GFAP and NeuN (Figures 1C and 1D).

Figure 1.

Preparation and identification of mice

(A) The breeding arrangement of mice.

(B) The genotype identification of mice. +/− is heterozygote, +/+ is mutant, and −/− is wild type.

(C and D) Immunofluorescence confirmed the AAVs were separately expressed in astrocytes (GFAP, C) and neurons (NeuN, D). Scale bar, 5 μm.

Table 1.

Primer sequences for mouse genotyping

Primers	Forward	Reverse
Dendra 2 WT	CGCGACACTGTAATTTCATACTG	CTTCCCTCGTGATCTGCAAC
Dendra 2 Loxp	GGACATCCCCGACTACTTCA	GCTCCCACTTCAGGGTCTTC

Table 2.

Sequences for AAV

Key component	Coding sequence
Cre	cccaagaagaagaggaaggtgtccaatttactgaccgtacaccaaaatttgcctgcattaccggtcgatgcaacgagtgatgaggttcgcaagaacctgatggacatgttcagggatcgccaggcgttttctgagcatacctggaaaatgcttctgtccgtttgccggtcgtgggcggcatggtgcaagttgaataaccggaaatggtttcccgcagaacctgaagatgttcgcgattatcttctatatcttcaggcgcgcggtctggcagtaaaaactatccagcaacatttgggccagctaaacatgcttcatcgtcggtccgggctgccacgaccaagtgacagcaatgctgtttcactggttatgcggcggatccgaaaagaaaacgttgatgccggtgaacgtgcaaaacaggctctagcgttcgaacgcactgatttcgaccaggttcgttcactcatggaaaatagcgatcgctgccaggatatacgtaatctggcatttctggggattgcttataacaccctgttacgtatagccgaaattgccaggatcagggttaaagatatctcacgtactgacggtgggagaatgttaatccatattggcagaacgaaaacgctggttagcaccgcaggtgtagagaaggcacttagcctgggggtaactaaactggtcgagcgatggatttccgtctctggtgtagctgatgatccgaataactacctgttttgccgggtcagaaaaaatggtgttgccgcgccatctgccaccagccagctatcaactcgcgccctggaagggatttttgaagcaactcatcgattgatttacggcgctaaggatgactctggtcagagatacctggcctggtctggacacagtgcccgtgtcggagccgcgcgagatatggcccgcgctggagtttcaataccggagatcatgcaagctggtggctggaccaatgtaaatattgtcatgaactatatccgtaacctggatagtgaaacaggggcaatggtgcgcctgctggaagatggcgattag
tdTomato	atggtgagcaagggcgaggaggtcatcaaagagttcatgcgcttcaaggtgcgcatggagggctccatgaacggccacgagttcgagatcgagggcgagggcgagggccgcccctacgagggcacccagaccgccaagctgaaggtgaccaagggcggccccctgcccttcgcctgggacatcctgtccccccagttcatgtacggctccaaggcgtacgtgaagcaccccgccgacatccccgattacaagaagctgtccttccccgagggcttcaagtgggagcgcgtgatgaacttcgaggacggcggtctggtgaccgtgacccaggactcctccctgcaggacggcacgctgatctacaaggtgaagatgcgcggcaccaacttcccccccgacggccccgtaatgcagaagaagaccatgggctgggaggcctccaccgagcgcctgtacccccgcgacggcgtgctgaagggcgagatccaccaggccctgaagctgaaggacggcggccactacctggtggagttcaagaccatctacatggccaagaagcccgtgcaactgcccggctactactacgtggacaccaagctggacatcacctcccacaacgaggactacaccatcgtggaacagtacgagcgctccgagggccgccaccacctgttcctggggcatggcaccggcagcaccggcagcggcagctccggcaccgcctcctccgaggacaacaacatggccgtcatcaaagagttcatgcgcttcaaggtgcgcatggagggctccatgaacggccacgagttcgagatcgagggcgagggcgagggccgcccctacgagggcacccagaccgccaagctgaaggtgaccaagggcggccccctgcccttcgcctgggacatcctgtccccccagttcatgtacggctccaaggcgtacgtgaagcaccccgccgacatccccgattacaagaagctgtccttccccgagggcttcaagtgggagcgcgtgatgaacttcgaggacggcggtctggtgaccgtgacccaggactcctccctgcaggacggcacgctgatctacaaggtgaagatgcgcggcaccaacttcccccccgacggccccgtaatgcagaagaagaccatgggctgggaggcctccaccgagcgcctgtacccccgcgacggcgtgctgaagggcgagatccaccaggccctgaagctgaaggacggcggccactacctggtggagttcaagaccatctacatggccaagaagcccgtgcaactgcccggctactactacgtggacaccaagctggacatcacctcccacaacgaggactacaccatcgtggaacagtacgagcgctccgagggccgccaccacctgttcctgtacggcatggacgagctgtacaag

Prepare customized coverslips

Timing: 2 days

• 3.
  Prepare titanium rings and coverslips (Figures 2A–2C).

  • a.
    Titanium ring specifications: outer diameter of 4.5 mm, inner diameter of 3.3 mm, radial width of 0.6 mm, and thickness of 0.5 mm.
  • b.
    Section the coverslips with a thickness of 0.13–0.17 mm into wafers with a diameter of 3.6 mm using a precision cutting instrument.

CRITICAL: Titanium rings are non-ferromagnetic, making them a superior choice compared to other ferromagnetic metals for applications.

• 4.
  Fabricate customized titanium rings and coverslips (Figures 2D–2F).

  • a.
    Clean and dry titanium rings and coverslips.
  • b.
    Glue titanium rings and coverslips.

    • i.
      Smear adhesive (Norland optical adhesive 6.1) and put the coverslip on one side of the ring.
    • ii.
      Expose to ultraviolet (UV) light using a 365 nm wavelength light-emitting diode (LED) curing lamp.
    • iii.
      Soak in alcohol.

Figure 2.

Preparation of customized coverslips

(A) Anterior view of the titanium ring.

(B) Lateral perspective of the titanium ring.

(C) Frontal aspect of the coverslip.

(D) Titanium ring post-adhesive application.

(E) Titanium ring with integrated coverslip.

(F) UV light using a 365 nm wavelength light-emitting diode (LED) curing lamp.

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Antibodies
NeuN monoclonal antibody (1:500)	Abcam	Cat# ab104224; RRID: AB_10711040
GFAP (GA5) monoclonal antibody (1:500)	Cell Signaling Technology	Cat# 3670; RRID: AB_561049
Goat pAb to Ms IgG Alexa Fluor 647 (1:200)	Abcam	Cat# ab150115; RRID: AB_2687948
Bacterial and virus strains
AAV2/PHP.eB-GfaABC1D-Cre-WPRE	OBiO	N/A
AAV2/PHP.eB-Syn-tdTomato-WPRE	OBiO	N/A
Chemicals, peptides, and recombinant proteins
Dental cement	This paper	N/A
Cyanoacrylate glue	This paper	N/A
0.5% proparacaine hydrochloride ophthalmic solution	Alcon	N/A
Agarose	BIOWEST agarose	Cat#BY-R0100
Optical adhesive 6.1	Norland	Cat# 289
Isoflurane	RWD	Cat# R510-22-10
Critical commercial assays
Mounting medium with DAPI-aqueous, fluoroshield	Abcam	Cat# ab104139
DNA isolation kit	Vazyme Biotech	Cat# DC102
Taq master mix	Vazyme Biotech	Cat# P112
DL5000 DNA marker	Vazyme Biotech	Cat# MD102-01
Experimental models: Organisms/strains
Male and female C57BL/6 mice with PhAMfloxed genotype (10–12 weeks old, for breeding)	The Jackson Laboratory	Strain# 018385; IMSR_JAX 018385
Male C57BL/6 mice with PhAMfloxed genotype (5–6 weeks old, for AAV injection)	This paper	N/A
Oligonucleotides
Primers for qPCR, see Table 1	This paper	N/A
AAV sequence, see Table 2	This paper	N/A
Software and algorithms
ImageJ	National Institutes of Health	https://imagej.nih.gov/ij
Imaris software	Bitplane	https://imaris.oxinst.com/
NIS-Elements Viewer	Nikon	https://www.microscope.healthcare.nikon.com
Other
29G Ultra-fine insulin syringe	BD	N/A
Microforceps	Ideal-tek	Cat# 5B.SA
High-speed drill	RWD	Cat# 78001
Carbide cutter (drill size: 0.5-mm diameter)	Meisinger	Cat# HM1-005-HP
Carbide cutter (drill size: 0.6-mm diameter)	Meisinger	Cat# HM1-006-HP
Stereotaxic frame	RWD	Cat# 68045
Small animal anesthesia machine	RWD	Cat# R500IE
Confocal microscope	Olympus	Cat# CSU-W1
Ni-E Microscope	NiKON	Cat# A1R MP
Stereomicroscopes	Daofeng	N/A
Peristaltic pump	Kamoer	Cat# NCKP-S04B
Stainless-steel screw (size: 0.8-mm diameter)	This paper	N/A
Head plate	This paper	N/A
Titanium ring	This paper	N/A
Coverslip	This paper	N/A

Step-by-step method details

Retro-orbital injections of AAV

Timing: 3 weeks

This section describes the steps of retro-orbital injections of AAV.

CRITICAL: AAV-injected mice are used to complete all experiments 3 weeks after infection.

• 1.Dilute 2 $\times$ 10¹¹ vg of each AAV in 100 μL normal saline for one mouse. Troubleshooting 1.
• 2.
  Injection of AAV.

  • a.
    Anesthetize mouse.

    • i.
      Place mouse in an anesthesia induction chamber to induce anesthesia with isoflurane at 3 mL/h.
    • ii.
      Maintain anesthesia via inhalation of isoflurane at 1.5 mL/h through the mask.
    • iii.
      Accomplish ophthalmic anesthesia using proparacaine hydrochloride eye drops.

      Optional: Other topical anesthetics may be employed for inducing anesthesia in murine palpebral tissues, such as 4% oxybuprocaine hydrochloride eye drops.

  • b.
    Use 29-gauge insulin syringes to finish the retro-orbital injection. Troubleshooting 2.

    CRITICAL: The needle bevel orientation is a critical determinant of successful injection. We developed this protocol based on previous ref.^3,4

    • i.
      Remove excess ophthalmic anesthetics by holding an absorbent gauze pad to the medial canthus.
    • ii.
      Introduce the needle into the medial canthus at an angle of approximately 30° with bevel down.

      CRITICAL: The majority of injections are performed with the bevel of the needle oriented upwards. However, for retro-orbital injections, orienting the needle so that the bevel faces down has been demonstrated to reduce the risk of damage to the eyeball.

    • iii.
      Inject slowly and smoothly, with a rate of less than 10 μL/s.

      CRITICAL: In general, no pullback is conducted prior to the injection to avoid vascular collapse.

    • iv.
      Withdraw the needle slowly and smoothly after injection.
  • c.
    Placed mouse on a warming pad for recovery.

Cranial window preparation

Timing: 1 day

This section delineates the methodology for cranial window preparation, encompassing bone window localization, craniotomy procedures, titanium ring fixation and head plate implantation. All mice are allowed 24 h for postoperative recovery in scanning prior.

• 3.
  Anesthetize mouse.

  • a.
    Place mouse in an anesthesia induction chamber to induce anesthesia with isoflurane at 3 mL/h.
  • b.
    Position mouse in a stereotaxic frame with anesthesia via inhalation of isoflurane at 1.5 mL/h through the mask.

CRITICAL: Ensuring the cranium is parallel alignment with the table.

• 4.
  Prepare zone of operation.

  • a.
    Remove the hair using an electric shaver for pets.
  • b.
    Perform strict disinfection.
  • c.
    Ophthalmic ointment is applied to prevent corneal drying and abrasion.
• 5.
  Bone window craniotomy. Troubleshooting 3.

  • a.
    Make an incision along the midline skin.
  • b.
    Debride periosteum using scalpel scraping.
  • c.
    Expose the bregma through sequential cleansing of the cranial surface with hydrogen peroxide and sterile saline solution.
  • d.
    Locate bregma and mark a circle centered at AP(Anterior-Posterior) −2.0 mm, ML(Medial-Lateral) 1.5 mm.
  • e.
    Drill a hole with a diameter of 3 mm under the stereomicroscope.

    • i.
      Initiate partial-thickness craniotomy along predetermined markings using carbide cutters (drill size: 0.5 or 0.6-mm diameter) (Figure 3A).

      CRITICAL: Following each 30 s drilling interval, wipe the operative site with saline to avoid heat-induced brain injury.

    • ii.
      Utilize a 29-gauge needle to carefully dissect between the external and internal plates of the skull along the craniotomy margins.

      Optional: Dip in saline for 60 s may facilitate bone flap mobilization.

    • iii.
      Elevate the bone flap using a 29-gauge needle in conjunction with microforceps (Figure 3B).

      CRITICAL: During bone flap elevation, maintain a minimal angle to prevent contralateral dural trauma.

    • iv.
      Keep the bone flap properly in the sterile 48-orifice plate and make records.
• 6.
  Fix customized titanium ring. Troubleshooting 4.

  • a.
    Place the customized titanium ring over the cranium window with coverslip downward.
  • b.
    Employ the stereotaxic microinjector holder to apply gentle downward pressure on the titanium ring.

    CRITICAL: The titanium ring should be carefully positioned to ensure optimal apposition with the cranium window. Excessive compression should be avoided to mitigate the risk of hemorrhage.

  • c.
    Bond the titanium ring and skull using cyanoacrylate glue.
  • d.
    Remove the stereotaxic microinjector holder (Figures 3C and 4A).
• 7.
  Affix custom-made head plate. Troubleshooting 5.

  • a.
    Drill a hole near the head plate using a carbide cutter (drill size: 0.6-mm diameter).
  • b.
    Screw a stainless-steel screw (size: 0.8-mm diameter) into the hole.
  • c.
    Utilize cyanoacrylate glue to affix the screw.
  • d.
    Apply moderate dental cement to encompass the extramural region of the titanium ring, including the screw fixtures, ensuring robust fixation of the customized platform (Figure 3D).

CRITICAL: Excessive application of dental cement should be avoided to prevent contamination of the coverslip and minimize the overall mass of the apparatus, thereby reducing potential discomfort to the mice.

• 8.Clean and disinfect the wounds, dry and then apply an appropriate amount of erythromycin ointment.
• 9.Switch off the isoflurane and inject 4 mg/kg/day meloxicam subcutaneously on the back of the mice when the mouse is close to recovery.

CRITICAL: The dosage of meloxicam should be adjusted in accordance with the condition of the mice and be administered for a period of three consecutive days in accordance with the experimental arrangement.

• 10.House all mice individually during recovery to avoid removal of head plate by other animals.

Figure 3.

Bone window craniotomy followed with customized titanium ring fixation

(A) Disruption of the cranial outer plates.

(B) Excision of the bone flap.

(C) Affixation of titanium ring.

(D) Installation of the stabilization apparatus.

(E) Positioning of mouse on the imaging platform.

(F) Microscopic examination of the prepared mouse.

Figure 4.

Cases of adequacy and inadequacy

(A) Adequate installation of titanium ring.

(B) Inadequate installation with bleeding 1.

(C) Inadequate installation with bleeding 2. The hemorrhagic foci are marked with red arrowheads.

Intravital two-photon microscopy

Timing: 1 h

• 11.
  Anesthetize mouse.

  • a.
    Place mouse in an anesthesia induction chamber to induce anesthesia with isoflurane at 3 mL/h.
  • b.
    Position mouse on a customized microscope stage with anesthesia via inhalation of isoflurane at 1.5 mL/h through the mask (Figures 3E and 3F).

CRITICAL: While two-photon microscopy requires a low-temperature environment, maintenance of body temperature of mouse can be achieved through a heating pad.

• 12.Clean the coverslip using saline.
• 13.Employ a 40 $\times$ 0.8 NA water immersion objective lens for scanning.
• 14.Set magnification to 40 $\times$ at a 1024 $\times$ 1024 scan size.
• 15.Set suitable excitation wavelength of IRNDD model as 920 nm with a 1-mm step size for z-stack.
• 16.Set the emission wavelength of FITC to 525 nm and tdTomato to 575 nm.
• 17.
  Follow-up arrangement of mouse after scanning.

  • a.
    Schedule other experiments for mouse.

    • i.
      Slowly remove the brain plate and titanium ring.
    • ii.
      Put the bone flap back and close the wound.
    • iii.
      Return the mouse to the home cage after it gains full consciousness.
    • iv.
      Meloxicam is administered subcutaneously daily for 3 days.
  • b.
    Schedule no experiments for mouse.

    • i.
      Deeply anesthetize the mouse with 100 mg/kg pentobarbital sodium.
    • ii.
      Transcardially perfuse with pre-cooled PBS (pH 7.4) and subsequently with 4% paraformaldehyde through a peristaltic pump (NCKP-S04B, Kamoer, China).
    • iii.
      Remove the brain and immerse in 4% paraformaldehyde for 16–17 h post-fixation.
    • iv.
      Sequentially dehydrate for 24 h in 10%, 20%, and 30% sucrose in PBS (pH 7.4).
    • v.
      Froze the perfused brain for further detection.

Result reviewing

Timing: 10 min

• 18.View all results with NIS-Elements Viewer and ImageJ.
• 19.
  Render 3D models with Imaris software (Bitplane).

  • a.
    Import and convert image stacks into .ims files in Arena modules of Imaris 10.0.
  • b.
    Select the data to be analyzed into Surpass modules.
  • c.
    Use the Spots in Scene modules to simulate mitochondria and the Surfaces to simulate neurons.

    Optional: Crop the region of interest using Crop 3D, ensuring the same size is used across different groups to maintain consistency in the scale.

  • d.
    Turn off the original fluorescence in Display Adjustment modules after simulation.
  • e.
    Use Color modules to mark mitochondria at different locations (Figure 5).
  • f.
    Use Animation modules to record a dynamic rotation video to better present the relationship between mitochondria and neurons.

Figure 5.

Example of astrocytic mitochondrial visualization in mouse neurons

Astrocytic mitochondria (green) in neurons (tdTomato, red) of mice were measured by 2-photon laser scanning microscopy. The 3D reconstruction of astrocytic mitochondria in neurons is illustrated by the yellow dots. Scale bar, 3 μm.

Expected outcomes

Building upon the injection of AAV based on the PhAM^floxed mice, this protocol enables users to capture imaging of astrocytic mitochondria in neurons for in vivo functional experiments.⁵ We used 2 $\times$ 10¹¹ vg AAV to target astrocytes directed by the astrocytic-specific GfaABC1D promoter.¹ However, it is imperative to empirically determine the appropriate number of AAV particles for each experiment, depending on the fluorescence intensity.

Skull penetration is a major challenge for in vivo mitochondrial fluorescence imaging of mouse brain tissue. To address this issue, we developed a novel imaging window consisting of a titanium ring integrated with a cover glass mounted on a custom-designed brain platform (Figures 3D and 4A). This device provided a stable and transparent interface for two-photon microscopy. When used in conjunction with fluorescently labeled transgenic mice, this setup allowed clear visualization of astrocytic mitochondria and neurons in the mouse brain.¹

It should be noted that the quality and depth of two-photon imaging is critically dependent on two factors: (1) the absence of bleeding in the cranial window and (2) the stability of the imaging platform. Therefore, refinement of the surgical technique is essential for optimal results. Further practice and optimization of these procedures may be required to achieve consistently high-quality deep tissue imaging. In addition, the customized cranial window is adaptable for cerebral vascular observation, brain blood flow monitoring, calcium signal measurement and continuous evaluation of newly formed tissue growth.

Limitations

There are several limitations in our protocol. First, the restricted view through the customized titanium ring limits observations to localized areas and precludes a comprehensive view of the entire region. Second, AAV-mediated gene delivery may result in off-target neuronal expression. Third, craniotomy procedures require crucial operator expertise and skill acquired through extensive practice. In addition, current technological limitations make prolonged dynamic observation difficult.

Troubleshooting

Problem 1

Astrocytic and neuronal mitochondria are not fluorescently labeled (see step 1).

Potential solution

Optimizing the injection dose of AAV prior to the trial can mitigate problems caused by variations in viral concentration or purity. In addition, ensuring effective intravascular delivery is critical. If retro-orbital injection is technically challenging, tail vein administration can be used as an alternative route of delivery. However, it is essential to validate the efficacy of the injection concentration through preliminary studies prior to new implementation.

Problem 2

In cranial thinning procedures, excessive drilling may result in penetration of the internal skull plate, potentially resulting in injury to the dura mater or underlying cerebral parenchyma (see step 5e i).

Potential solution

Slowly drill the skull, and timely cool the skull.

Problem 3

The skull is difficult to remove or tends to bleed (Figures 4B and 4C) (see step 5e iii).

Potential solution

The outer and inner plates of the bone flap are completely separated using a 29G needle. A small portion of the bone flap is then elevated using the needle tip, followed by complete elevation of the flap using forceps. In the event of bleeding, the bone spurs are carefully arranged at the flap margins. Hemostasis is achieved by local application of a small saline-soaked pledget for 30 s. Excess fluid is then absorbed by applying dry pledget. If the initial hemostatic efforts proved inadequate, the duration of saline application could be extended appropriately and the procedure repeated until satisfactory hemostasis was achieved.

Problem 4

The titanium ring did not adhere adequately to the cranial surface, allowing cyanoacrylate glue to infiltrate the cranial window during subsequent fixation procedures. This compromised the clarity of the imaging field (see step 6b and 6c).

Potential solution

During cranial polishing, an outward expansion technique should be employed to ensure the cranial window diameter meets or slightly exceeds 3 mm. Prior to utilizing the stereotaxic microinjector holder, the coverslip should be precisely aligned with the cranial window. Subsequently, appropriate pressure should be applied to ensure complete insertion of the coverslip into the cranial window.

Problem 5

Detachment of the custom-made head plate and respiratory-induced motion artifacts result in compromised imaging field stability (see step 7).

Potential solution

As described in 7a and 7b, a screw is added to the posterior side of the mouse head plate.

Resource availability

Lead contact

Further requests for information should be directed to the lead contact, Tao Li (scutaoli1981@scu.edu.cn).

Technical contact

Requests for technical information should be directed to technical contact, Jian Zhou (zhoujian2019@swmu.edu.cn) and Fengling Du (dufengling00@swmu.edu.cn).

Materials availability

This study did not generate new unique reagents.

Data and code availability

This study did not generate any codes.

Acknowledgments

This work was supported by grants from the National Natural Science Foundation of China (82371310, 82370260, 82271306, 81971132, and 82401715), the Young Elite Scientist Sponsorship Program by the China Association for Science and Technology (YESS20200178), the Sichuan Province Science and Technology Support Program (2023YFH0069, 2023NSFSC0028, 2023NSFSC1559, 2022YFS0615, and 2022NSFSC1421), and the Clinical Research Special Project of Southwest Medical University (2024LCYXZX32). Some graphical elements are created with Biorender.com and ScienceSlides suite.

Author contributions

J.Z., F.D., and X.Z. performed the AAV injection. J.Z., F.D., F.Z., and T.T. performed bone window craniotomy. Y.W. and L.Z. performed mice reproduction and identification. P.L. performed intravital two-photon microscopy. J.Z. and F.D. prepared figures for the manuscript. J.Z., Y.J., J.P., and T.L. edited the manuscript. J.Z., J.P., L.Z., T.L., and Y.J. allocated funding and finalized the manuscript.

Declaration of interests

The authors declare no competing interests.

Declaration of generative AI and AI-assisted technologies in the writing process

During the preparation of this manuscript, the authors used ChatGPT in order to refine the content. After utilizing this service, the authors carefully reviewed and revised the manuscript as necessary and take full responsibility for the content of the publication.