Cerebrospinal fluid (CSF) analysis is a powerful tool for enhancing the detection of various processes occurring within the central nervous system, such as bleeding, inflammation, infection, tumors, and other clinical conditions. Additionally, serial evaluations of compounds in CSF (such as hormones, drugs, neurotransmitters, biomarkers, metabolites, and viruses) are equally crucial in both basic and translational research. The volume of CSF in the central nervous system is maintained stable through secretion and absorption.
I.Operational Considerations
CSF is typically collected via either a cerebral ventricle or lumbar puncture. Lumbar puncture carries a relatively lower risk of complications in non-human primates because, in most species, the lumbar puncture site is distant from the spinal cord's end. Proper anesthesia and aseptic techniques, including site preparation such as shaving, and the use of sterile materials, are necessary. At least sterile gloves, syringes, and needles should be used, with consideration for sterile surgical drapes. CSF collection is usually performed in lateral recumbency, sternal recumbency, or sitting positions, improving access to the cerebral ventricles or enlarging the intervertebral space through flexion of the neck or spine. The size and length of the chosen needle should be suitable for the species and size of non-human primates. Techniques, equipment, consumables used during the procedure, and perioperative care vary depending on the species, research objectives, and clinical veterinarian preferences.
Some research protocols may require continuous collection of CSF over time or collection of CSF samples from unanesthetized subjects. For these protocols, continuous CSF collection from awake rhesus monkeys can be achieved via long-term implanted CSF catheters connected to subcutaneous collection ports.
The author has made every effort to find the latest references regarding total CSF volume, limits or recommendations for dural punctures, collection frequency, safe withdrawal volume, and CSF replenishment rates. Literature on this aspect is limited for most non-human primates. Additionally, collection methods vary, as surveys from members of the Non-Human Primate Veterinary Association indicate that various institutions successfully use different methods for CSF collection. Therefore, the author provides considerations and recommendations for CSF collection based on the existing best evidence.
For reference, previous studies have determined that the apparent CSF volume in adult male rhesus monkeys (8.0-10.6 kg) is 10+/-0.06 mL, with a CSF flow rate of 0.018+/-0.003 mL/min. The CSF formation rate in rhesus monkeys ranges from 28.6-44.2 μL/min. Further research on non-human primates is needed in this area to facilitate the development of more instructive recommendations.
II. Potential Adverse Reactions
Although complications during CSF collection in non-human primates are rare, anatomical variations and improper or inadequate technical methods may lead to sample damage or complications such as bleeding, herniation, infection, damage to the brainstem, spinal cord, or nerve roots, and animal discomfort. While these complications are rare, they can compromise animal welfare.
Due to the risks involved, CSF should be collected by a veterinarian or trained veterinary personnel. For unsuccessful attempts at CSF collection, practices may vary. It is common practice to allow up to three unsuccessful attempts by one individual. After the third unsuccessful attempt, another qualified individual should take over. If the second individual is unable to collect CSF after three attempts, the attempts should be stopped. Meanwhile, the animal will receive appropriate analgesic treatment and recover from anesthesia. Changing the collection site (e.g., from cerebral ventricles to lumbar) after unsuccessful attempts often results in success. If a failed attempt without imaging guidance occurs, fluoroscopy-assisted needle placement can be used. Besides increasing the success rate, image guidance can also reduce the rate of traumatic punctures.
Postoperative monitoring should continue until the animal returns to its cage and can maintain an upright posture without support. Monitoring of animals should last for at least 24 hours or according to the protocol approved by the Institutional Animal Care and Use Committee (IACUC). Monitoring should include assessments of basic physiological, behavioral, and locomotor parameters. Signs of neurological dysfunction, including but not limited to lethargy, depression, coma, and signs of head pressing, should be assessed particularly. Headaches and consequent intracranial hypotension following CSF collection are not uncommon in human medicine. Considering this potential complication in non-human primates, analgesic treatment should be provided during surgery and continued for 2-3 days postoperatively, or as needed based on postoperative observations related to behaviors associated with headache or spinal pain. The most effective analgesic regimen may involve comprehensive treatment initiated preoperatively.
III. IACUC Considerations and Record Keeping
The IACUC should be familiar with the procedures related to CSF collection and all potential complications and review them at least annually. All relevant information, such as drug dosages, preoperative preparations, CSF volumes collected, collection frequency, and short-term and long-term postoperative recovery, should be described in the IACUC protocol and appropriately documented in medical records or laboratory logs.
While CSF may be crucial data in research protocols, improper collection techniques can affect animal welfare. When submitting protocols, scientific justification for the number and frequency of collections should be provided. Specifically, the IACUC should carefully evaluate each proposal involving CSF collection and consider the following:
1.Is CSF collection essential to achieving the scientific objectives outlined in the protocol?
2.Are there alternative methods and less invasive central nervous system evaluation methods (such as imaging techniques) available, and if so, have they been considered?
3.Have the operators received appropriate training and possess sufficient skills?
4.Will appropriate perioperative anesthesia and analgesic regimens be provided? Consultation with laboratory animal veterinarians should be sought regarding anesthesia and analgesic plans.
5.Does the collected CSF volume consider the age and size of the animal?
6.Are the number and frequency of dural punctures suitable for the animal, and has the risk of complications been minimized? If multiple CSF collections are part of the study, the maximum collection times for each animal should be specified.
7.Is the postoperative monitoring plan appropriate?
8.Have all possible postoperative complications been considered, and have all reasonable interventions, including euthanasia, been listed?
