Integrated Protocol for Assessing Meningeal Lymphatics: Cisterna Magna Injection, Lymph Node Imaging, Meningeal Dissection, and Whole-Mount Staining

Summary

Here, a protocol is presented to assess mouse meningeal lymphatic vessels (MLVs), which play an important role in multiple neurologic diseases. The goal of this protocol is to provide an integrated method that enables both functional evaluation of tracer drainage and detailed morphological characterization of the dorsal MLVs network.

Abstract

Meningeal lymphatic vessels (MLVs) have emerged as critical regulators of central nervous system (CNS) homeostasis, and MLVs dysfunction has been implicated in various neurological disorders. A comprehensive assessment of MLVs' morphology and function is therefore essential for understanding their role in CNS physiology and disease. This protocol describes an integrated method for evaluating MLVs in mice using four key techniques: Intra-cisterna-magna (i.c.m.) injection of fluorescent tracers, live imaging of cervical lymph nodes (CLNs), meningeal dissection, and whole-mount staining. First, the 30 G needle is connected to the syringe and infusion pump via a polyethylene (PE) tube. After the mouse is anesthetized, the dura mater is exposed by blunt dissection of the neck muscles, and the injection needle is inserted into the cisterna magna (CM) to deliver the tracer into the cerebrospinal fluid (CSF) with precise control. Tracer drainage to the CLNs can be visualized in real time, while subsequent meningeal dissection and whole-mount staining enable detailed morphological analysis of the dorsal MLVs. The entire procedure can be completed within 3 h, encompassing i.c.m. injection, tissue perfusion, skull isolation, and placement in fixative solution. This protocol can also be adapted for intracranial drug delivery or combined with molecular analyses, as the meningeal dissection procedure is compatible with fresh, unfixed tissues suitable for RNA and protein extraction.

Introduction

MLVs are located within the dural sinuses and are able to carry CSF components and immune cells from the CNS to the CLNs^1,2. The discovery of MLVs has challenged the traditional concept of central nervous system "immune privilege" and has emerged as a research hotspot in recent years³. Studies have shown that impaired MLVs function is closely associated with various diseases, including Alzheimer's disease, age-related cognitive decline, brain tumors, and CNS infections^4,5,6,7. As research in this field advances, there is a growing need for integrated, reproducible, and well-standardized experimental methods.

Most existing protocols for lymphatic vessel research focus on peripheral lymphatic vessels^8,9,10. In studies related to MLVs, A study described a straightforward CM cannulation procedure that enables direct access to the CSF without damaging the skull or brain parenchyma¹¹. When combined with imaging techniques, this method can effectively assess glymphatic system function and CSF dynamics. However, detailed experimental protocols for studying the function and morphology of MLVs have not been well documented.

The present protocol integrates functional imaging with morphological analysis, enabling simultaneous acquisition of drainage function and structural morphology data from the same animal, thereby reducing the number of animals used and improving data comparability and consistency. Overall, this protocol can be used to study structural and functional changes in MLVs in various disease models, as well as to evaluate the effects of pharmacological or genetic interventions on MLVs. The protocol provides detailed descriptions of the operational steps, key precautions, and troubleshooting guidance. In addition, the i.c.m. injection technique described in this protocol can be independently applied for intracranial drug delivery, and the meningeal dissection procedure is compatible with fresh tissue for subsequent RNA or protein extraction, offering considerable flexibility and scalability.

Protocol

The mouse procedures described in this protocol were performed in compliance with the protocol approved by the Institutional Animal Care and Use Committee (IACUC) of Shanghai University (Approval No. ECSHU 2023-099). 

1. Preparation of surgical instruments and buffers

CAUTION: Operate in a chemical fume hood and wear gloves to minimize the potential risks when working with PFA, a moderately toxic and carcinogenic substance. All operations involving Isoamyl alcohol must be carried out in a chemical fume hood away from ignition sources, while wearing gloves and a mask, to reduce exposure risks.

1. Prepare surgical instruments, such as one pair of straight scissors, two pairs of curved fine-tipped forceps, two pairs of blunt-tipped forceps, one pair of micro scissors, and one pair of straight micro forceps with smooth (non-serrated) tips.
   NOTE: Wipe surgical instruments with 75% ethanol and keep them dry both before and after the experiment.
2. Prepare buffers.
   1. 1x PBS.
   2. 4% Paraformaldehyde Fix Solution (4% PFA).
   3. Blocking solution (1x PBS containing 3% bovine serum albumin (BSA), 0.5% Triton X-100, and 2% fetal bovine serum).
   4. Antibody dilution buffer (1x PBS containing 3% BSA and 0.5% Triton X-100).
   5. Antifade Mounting Medium.
      ​NOTE: The prepared reagent can be stored at 4 °C for one month.
3. Prepare 2,2,2-Tribromoethanol anesthetic solution (Avertin, 1.25%, 50 mL)
   1. Weigh 0.625 g Tribromoethanol into a 2 mL tube, add 1.25 mL Isoamyl alcohol, and dissolve thoroughly (37 °C shaker).
   2. Transfer the fully dissolved Tribromoethanol-isoamyl alcohol solution to a 50 mL conical tube, add 1x PBS to bring the total volume to 50 mL, and mix thoroughly.
   3. Filter the solution through a 0.22 µm filter in a biological safety cabinet.
4. Prepare Ovalbumin, Alexa Fluor 647 Conjugate (OVA-647) solution (2 mg/mL). Dissolve 2 mg of OVA-647 in 1 mL of artificial cerebrospinal fluid (aCSF).

2. Assembly of the injection needle and connection to the microinjection pump

1. Use blunt forceps to break off the 30 G needle and retain the tip¹¹ (outer diameter 0.30 mm, inner diameter 0.15 mm, length 8 mm).
2. Use a 30 cm length of PE-10 tubing (outer diameter 0.64 mm, inner diameter 0.28 mm) to connect the broken 30 G needle to the microinjection needle containing a 30 G needle (Figure 1).
3. Mount the microinjection needle, with the tubing and 30 G needle attached, onto the microinjection pump.

3. i.c.m. injection and live imaging

CAUTION: If the needle is inserted too deeply and penetrates the brain parenchyma, the injectate will be delivered into the brain tissue rather than the CSF compartment, compromising the accuracy of the result and potentially leading to mouse mortality. During the surgical procedure, continuous physiological monitoring of the mouse is required. Anesthesia depth should be assessed every 15 min using the toe-pinch reflex to ensure the animal remains pain-free. Body temperature should be maintained at 37 °C ± 0.5 °C using a heating pad to prevent hypothermia and its associated effects on physiological function. Additionally, respiration and heart rate should be recorded every 30 min, assessed by observing thoracic movements and palpating the cardiac apex. During i.c.m. injection, if the needle is inserted too deeply and causes bleeding, this indicates a failed injection. Terminate the experiment and euthanize the animal.

1. Weigh the mouse and administer 200 mg/kg Avertin via intraperitoneal injection to anesthetize the mouse.
   NOTE: Alternative anesthetics (e.g., ketamine/xylazine) can be used.
2. Apply ophthalmic ointment to prevent corneal drying.
3. Apply depilatory cream to the cervical region to remove hair over the lymph nodes when the toe pinch reflex ceases, and respiration becomes slow and steady.
4. Shave the dorsal neck region.
5. Place the mouse in a stereotaxic frame on a heating pad with the head tilted at a 120° angle to the trunk (Figure 2A).
6. Clean the area sequentially with povidone-iodine and 75% ethanol.
7. Make a midline incision in the skin of the neck and bluntly separate the muscle layers.
   NOTE: Using two pairs of blunt-tipped forceps, separate the muscle layers to minimize bleeding.
8. Dissect through the muscle with fine-tipped forceps until the translucent dura mater overlying the CM becomes visible (Figure 2B, C).
   NOTE: Use a fine-tipped curved forceps to retract the muscles and maintain consistent exposure of the CM.
9. Under a stereomicroscope, use blunt forceps to hold the needle and insert it at a 45° angle to the plane of the dura mater to a depth of approximately 1–2 mm. Position the needle bevel parallel to the dura mater so that it lies completely beneath the dura mater (Figure 2D).
   NOTE: Apply gentle force with the needle to puncture the dura mater. Successful puncture is indicated by a palpable loss of resistance, along with visualization of the needle tip beneath the dura mater and the outflow of CSF.
10. Use absorbent paper to dry any leaking CSF.
11. Set the microinjection pump to deliver at 1 µL/min. Inject 5 µL of OVA-647 solution (2 mg/mL) into the CM⁴ (Figure 2E).
12. Leave the needle in place for 2 min to prevent backflow.
13. Apply one drop of biological glue onto the meninges.
14. Close the surgical incision.
15. While still under anesthesia, perform in vivo imaging of the mouse to monitor dye intensity in the CLNs at various time points (30 min, 60 min). Use the following imaging parameters: exposure time: 0.5 s; excitation filter: 640 nm; emission filter: 680 nm. Continue to monitor anesthetic depth and supplement with additional anesthetics if needed.

4. Isolating the dorsal aspect of the mouse skull

NOTE: Confirm the absence of the toe pinch reflex and the cessation of spontaneous respiration before initiating thoracic dissection.

1. Euthanize the mouse with an overdose of Avertin (600 mg/kg) and perform cardiac perfusion with 20 mL 1x PBS or normal saline.
2. Decapitate the mouse to isolate the head.
3. Make an incision along the midline of the skull to open the scalp.
4. Use two curved hemostats to separate the scalp bilaterally by gentle retraction. Subsequently, insert the curved hemostats into the orbital fossae and carefully strip off the residual scalp from the nasal and oral regions (Figure 3A–C).
   NOTE: This method enables rapid and complete separation of the scalp.
5. Remove the mandible while keeping the skull intact (Figure 3D).
6. Remove the bone from the base of the skull to expose the ventral aspect of the brain (Figure 3E).
7. Carefully extract the brain with blunt forceps, avoiding contact with the dura mater of the parietal bones (Figure 3F).
8. Completely immerse the dorsal skull in 5 mL, 4% PFA and fix for 24 h at 4 °C (Figure 3G).
   PAUSE POINTS: Store fixed skulls at 4 °C for no longer than one week. Remove the sample from 4 °C and allow it to return to room temperature before meningeal dissection.

5. Dissection of meninges from the mouse skull

1. Wash the fixed skull with 5 mL 1x PBS, repeating the wash three times (10 min each).
2. Place the skull dorsal-side down in a 10-cm dish filled with 1x PBS (Figure 4A).
3. Use the tip of fine-tipped forceps to grasp or press down on the tympanic bulla and remove it from the skull (Figure 4B).
   NOTE: Removing the tympanic bulla is a critical step to avoid meningeal tearing caused by adhesion during separation from the skull.
4. Trim closely along the ventral surface of the skull for a distance of about 2 mm with scissors to smooth them (Figure 4C).
5. Use fine forceps, meticulously clear residual white tissue from the confluence of sinuses (COS) region of the meninges (Figure 4D).
   NOTE: Incomplete removal of residual tissue is a major cause of nonspecific staining in immunolabeling.
6. Use fine micro forceps to gently lift one edge of the meninges from the skull margin, then lift the opposite edge (Figure 4E, F).
7. Grasp the bowl-shaped rim of the meninges and pull to separate it from the skull in the COS region (Figure 4G).
8. Remove the maxillary bone with scissors (Figure 4H).
9. Grasp the bowl-shaped rim of the meninges again and pull to separate the meninges from the skull at the frontal bone area (Figure 4I).
10. Place the intact meninges into a 24-well plate containing 1 mL 1x PBS, ready for staining.
    NOTE: If the meninges rupture during the autopsy, especially at the junction of the superior sagittal and transverse sinuses, the sample may not be suitable for further analysis. If high non-specific background staining is observed, add a quenching step after PFA fixation (e.g., incubate with 0.1 M glycine solution for 40 min¹²) to further reduce the background signal.

6. Blocking, primary antibody, and secondary antibody staining

1. Use a 1 mL pipette to aspirate the 1x PBS so that the meninges adhere to the bottom of the well plate.
2. Block the whole-mount meninges with 200 µL blocking solution in a 24-well plate for 1 h at room temperature.
   NOTE: Gently spread the meninges completely in the blocking solution using forceps to ensure even exposure.
3. Use a pipette to remove the blocking solution.
4. Incubate the whole-mount meninges with 200 µL of diluted anti-LYVE-1 (lymphatic vessel-specific marker¹³) antibody at a 1:200 dilution (final concentration 1 µg/mL) overnight at 4 °C.
5. Use a pipette to remove the antibody solution.
6. Wash three times with 1x PBS (10 min each).
7. Incubate whole-mount meninges with the DAPI (final concentration 1 µg/mL) and Alexa Fluor 488 donkey anti-rat IgG (final concentration 2 µg/mL) for 1 h at room temperature in 1x PBS.
   NOTE: Protect from light during antibody incubation.
8. Use a pipette to remove the antibody solution.
9. Wash three times with 1x PBS (10 min each).

7. Mounting and imaging

1. Pipette 100 µL of 1x PBS onto the center of a slide.
2. Float and spread the whole-mount meninges within the droplet (Figure 5A).
3. Aspirate the 1x PBS to allow the tissue to adhere flatly to the slide (Figure 5B).
4. Apply 20 µL of mounting medium onto the whole-mount meninges.
5. Place a coverslip over the tissue (Figure 5C).
6. Air-dry the slides at room temperature and store the slides at 4 °C.
   PAUSE POINTS: The slides can be stored at 4 °C for up to one month. Remove the slide from 4 °C and allow it to return to room temperature. Remove any condensation from the slide surface with absorbent paper.
7. Use a 20x objective lens to acquire MLVs images. Set the channel to FL Green with an emission wavelength of 518 nm and an exposure time of 100 ms.

Results

Following i.c.m. injection of OVA-647, in vivo imaging of small animals revealed distinct oval-shaped fluorescent signals distributed bilaterally in the cervical region (Figure 6A). In successful injections, the tracer was clearly visible in the CLNs after injection, with fluorescence intensity peaking at approximately 60 min (Figure 6B). In addition, fluorescent signals were observed in the isolated superficial and deep cervical lymph nodes (

Discussion

This protocol provides a comprehensive method for evaluating MLVs function and morphology by integrating i.c.m. injection, in vivo imaging of CLNs, meningeal dissection, and whole-mount staining. A key advantage of this integrated approach over conventional methods is that it enables simultaneous assessment of lymphatic drainage function and vessel morphology within the same animal, eliminating the need for separate cohorts and thereby reducing experimental variability while improving data comparability. Further...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
ACSF (Artificial Cerebrospinal Fluid)	Tocris	Cat# 3525
Alexa Fluor 488 donkey anti-rat IgG (1:1000 dilution)	Invitrogen	Cat# A-21208
Antifade Mounting Medium	Beyotime	Cat# P0126
Blunt-tipped forceps	RWD Life Science	Cat# F12011-10
BSA	Beyotime	Cat# ST023
Coverslips	CITOTEST	Cat# 80312-3161
Curved sharp forceps	RWD Life Science	Cat# F11003-11
DAPI	Sigma-Aldrich	Cat# D9542
Depilatory cream	Veet	N/A
Dish-10 cm	Thermo Fisher Scientific	Cat# 150464
Disposable intravenous (IV) infusion needle	Zhejiang Kindly Medical Devices	N/A
Ethanol-75%	Sinopharm Chemical Reagent	Cat# 80176961
Fetal Bovine Serum	Excell	Cat# FSD500
Filter-0.22 µm	Merck Millipore	Cat# SLGPR33RB
Glass slides	CITOTEST	Cat# 80340-3610
Isoamyl alcohol	Sinopharm Chemical Reagent	Cat# 10003218
IVIS Spectrum	PerkinElmer	N/A
Micro forceps	Shinva Medical Instrument	Cat# ZD372RB
Micro scissors	Shinva Medical Instrument	Cat# ZO13056R
Microinjection needle (25 µL)	Shanghai Gaoge Industry & Trade	N/A
Microinjection pump	Voyage Power	Cat# QHZS-001A
Mouse: C57BL/6 wild-type, 8 weeks old, male	Cavens	N/A
Needle-30G	Zhejiang Kindly Medical Devices	N/A
Normal Saline	Beyotime	Cat# ST341
Ovalbumin, Alexa Fluor™ 647 Conjugate	Thermo Fisher Scientific	Cat# O34784
Paraformaldehyde Fix Solution-4%	Beyotime	Cat# P0099
PE-10 tubing	Shanghai Qiujing Biochemical Reagent and Instrument	N/A
Phosphate-buffered saline (1x PBS)	Beyotime	Cat# ST447
Rat anti-LYVE-1 (1:200 dilution)	Santa Cruz	Cat# sc-65647
Stereomicroscope	Jonec	Cat# JSZ6
Stereotaxic frame	RWD Life Science	Cat# 68030
Straight scissors	RWD Life Science	Cat# S12007-10
Syringe-1-mL	Zhejiang Kindly Medical Devices	N/A
Tissue Adhesive I	3M Vetbond	Cat# 1469SB
Tribromoethanol	Sigma-Aldrich	Cat# T48402
Triton X-100	Sigma-Aldrich	Cat# T8787
VS120 High-throughput fluorescence imaging system	Olympus	N/A