Aug 29, 2025
Cerebrospinal Fluid (CSF) Collection in mouse models
- Lilia Crew1,
- Alyssa Seerley2,
- Serena McElroy2,
- Andrea Grindeland Panter1,2
- 1Touro University College of Osteopathic Medicine, Great Falls, MT, United States;
- 2Weissman Hood Institute at Touro University, McLaughlin Research Institute, Great Falls, MT, United States
- Lilia Crew: Co-first author;
- Alyssa Seerley: Co-first author
- Andrea Grindeland Panter: Senior author
Protocol Citation: Lilia Crew, Alyssa Seerley, Serena McElroy, Andrea Grindeland Panter 2025. Cerebrospinal Fluid (CSF) Collection in mouse models . protocols.io https://dx.doi.org/10.17504/protocols.io.36wgqpnwxvk5/v1
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: August 28, 2025
Last Modified: August 29, 2025
Protocol Integer ID: 225742
Keywords: Cerebrospinal fluid, CSF, mouse models, Neurodegeneration, cerebrospinal fluid, csf, mouse model, collection in mouse model, mouse
Funders Acknowledgements:
NIH General Medical Sciences
Grant ID: 5P20GM152335
Abstract
This protocol describes a method for the collection of Cerebrospinal fluid (CSF) from a mouse able to be used for downstream analysis.
Materials
Large Scissors
Small Scissors
70% EtOH
Fine Forceps (Dumont Curved)
Capillary tube made of polished borosilicate glass with filament, size O.D.:1.0mm, I.D.: 0.78mm.
Dissection microscope
Troubleshooting
Terminal Cerebrospinal Fluid (CSF) Collection
Apply 70% ethanol (EtOH) to the surgical area to wet down the hair of the mouse.
Using a #10 blade, make a midline skin incision over the occiput to the second cervical vertebrae (C2).
Incise between the cervical muscles on the midline over the occiput to C2, using sharp dissection. The cervical muscles rhomboideus cervicis, the cervical part of the trapezius, splenius capitis, semispinalis capitis, and erector spinae muscles will all need to be separated at midline to expose the cisterna magna; however, the deeper muscles which are difficult to visualize with the eye, may be separated under the dissecting microscope in step 6 using blunt dissection techniques.
Place the ventral surface of the thorax of the mouse on the weight boat under the microscope.
Stabilize the mouse, ensuring that the skull is hyper flexed for maximum access to the cisterna magna. See Fig. 1
Fig. 1 Mouse Positioning to obtain CSF. Anesthetized mouse positioned for cerebrospinal fluid (CSF) collection, demonstrating hyperflexion in the cranial-occipital region. The mouse is secured with orange tape to maintain the necessary posture for effective CSF extraction and hyperflexion is induced via a weigh boat and tape roll or any object that will hold the correct stabilization and placement of the head.
Bluntly dissect and separate at midline the interior muscles remaining from step 3 down to the cisterna magna to visualize a transparent membrane. This is the arachnoid membrane covering the cisterna magna, which is a large pocket of CSF within the subarachnoid space. If hemorrhage from the musculature occurs, blot it with a cotton swab prior to puncturing the arachnoid membrane as it is critical to avoid blood contamination in the CSF.
Once the arachnoid membrane is visualized, use small thumb forceps to move the muscles laterally if needed for complete visualization.
With the capillary tube, puncture the arachnoid membrane at a 45° angle, using a gentle spinning motion.
Note
Using a micropipette puller, make a needle from a 10 cm length capillary tube made of polished borosilicate glass with filament, size O.D.:1.0mm I.D.: 0.78mm. The end of the needle will need to be cut to a diameter of 0.2mm, as the larger diameter will allow the CSF to flow into the tube by capillary action. See Fig 2.
Fig.2 Capillary tube recommendations. Borosilicate glass with filament [O.D.: 1.0mm, I.D.: 0.78mm, 10cm length] capillary tube pulled into a needle and cut leaving a 0.2mm diameter opening for puncture of the arachidonic membrane.
Allow CSF to flow into the tube by capillary action. The approximate amount of CSF collected is between 4-10µL. See Fig 3.
Fig. 3 Capillary tube containing CSF.
Place the blunt end of the capillary tube into the rubber bulb capillary dispenser instrument and expel the sample into the desired sample tube. See Fig. 4
Fig. 4Transference of Collected CSF. Using a rubber bulb capillary dispenser into a storage vial in a laboratory setting. This process ensures the integrity and sterility of the sample for subsequent analyses.
Acknowledgements
Colony management for the mouse models included in this study was provided by the
McLaughlin Research Institute - Gene Editing and Mouse Models Assessment (GEMMA) Core
Facility within the Center for Integrated Biomedical and Rural Health Research,
RRID: SCR_027045, 1P20GM152335. The authors thank Rose Pitstick and Kaela Davey for exceptional care of the mice. We would also like to thank student intern research assistant Bridget Gray and federal work study Touro COM-MT students Dalia Shaaban and Clairissa Kaylor for thoughtful discussions regarding planning and presentation of dissection methods in the mouse brain microdissections.