Intrathecal Injection of Newborn Mouse for Genome Editing and Drug Delivery
Summary
The present protocol outlines step-by-step instructions for performing intrathecal injections in neonatal mice for gene editing and drug delivery.
Abstract
Intrathecal injection is a commonly employed procedure in both pediatric and adult clinics, serving as an effective means to administer medications and treatments. By directly delivering medications and treatments into the cerebrospinal fluid of the central nervous system, this method achieves higher localized drug concentrations while reducing systemic side-effects compared to other routes such as intravenous, subcutaneous, or intramuscular injections. Its importance extends beyond clinical settings, as intrathecal injection plays a vital role in preclinical studies focused on treating neurogenetic disorders in rodents and other large animals, including non-human primates. However, despite its widespread application, intrathecal injection in young, particularly neonatal pups, poses significant technical challenges due to their small size and fragile nature. Successful and reliable administration of intrathecal injections in newborn mice requires meticulous attention to detail and careful consideration of various factors. Thus, there is a crucial need for a standardized protocol that not only provides instructions but also highlights key technical considerations and good laboratory practices to ensure procedural consistency, as well as the safety and welfare of the animals.
To address this unmet need, we present a detailed and comprehensive protocol for performing intrathecal injections specifically in newborn pups on postnatal day 1 (P1). By following the step-by-step instructions, researchers can confidently perform intrathecal injections in neonatal pups, enabling the accurate delivery of drugs, antisense oligos, and viruses for gene replacement or genome editing-based treatments. Furthermore, the importance of adhering to good laboratory practices is emphasized to maintain the well-being of animals and ensure reliable experimental outcomes. This protocol aims to address the technical challenges associated with intrathecal injections in neonatal mice, ultimately facilitating advances in the field of neurogenetic research that aims to develop potential therapeutic interventions.
Introduction
Intrathecal (IT) injection is a common clinical procedure used to administer medications, collect cerebrospinal fluid, and maintain intracranial pressure in both pediatric and adult patients in clinics1,2. The administration of medications via intrathecal injection is an effective approach for increasing medication concentrations in the central nervous system (CNS) while minimizing systemic exposure. Consequently, this method enhances therapeutic efficacy and reduces side effects, especially for temperature-sensitive and short half-life drugs3.
In preclinical studies testing new drugs and treatments using rodent models, it is imperative to employ a reliable method of drug administration that offers greater precision and result reproducibility4,5. For preclinical studies evaluating new treatments for neurogenetic and neurodevelopmental disorders, early treatment is crucial for initial proof-of-concept studies because earlier interventions are typically predicted to yield more favorable outcomes6,7,8.
Compared to conventional intracerebroventricular (ICV) injections, IT injections carry significantly lower risks since they obviate the need for direct penetration through the cerebral cortex. This advantage substantially reduces the potential damage to regional cortical tissue and surrounding nerves. Furthermore, IT injections allow for at least a fivefold increase in the administrable volume of medications through a single injection, greatly enhancing the feasibility of repeated administrations. However, due to the small size and fragile nature of newborn mice, performing intrathecal injections in newborn pups is technically challenging and requires specialized techniques, equipment, and meticulous handling.
This article provides a detailed protocol with step-by-step instructions for performing intrathecal injections in P1 newborn pups. The key considerations and good laboratory practices are emphasized here to ensure the consistency of administration and the safety and well-being of the animals during the procedure. By following this protocol, researchers can confidently conduct experiments with precision and reproducibility while minimizing any potential risks or discomfort to the animals.
Protocol
The described procedures and protocols were in compliance with the guidelines outlined in the National Institutes of Health Guide for the Care and Use of Laboratory Animals. Additionally, the procedures received approval from the Animal Care and Use Committee at Yale University School of Medicine. Newborn wild-type (WT) C57BL/6J male and female mice were used for presented study. The animals were obtained from a commercial source (see Table of Materials).
1. Preparation of the workspace
- Prepare the following items first: wet ice for cryo-anesthesia, an empty cage to separate the pups from the dam, a dissecting microscope, a light source, a clean surface for placing the animal during the injection, cotton swabs, a heating pad, a 25/10 µL syringe, and a 34 G/ 0.375"/ 12 DEG needle (see Table of Materials).
NOTE: Cryo-anesthesia for mouse pups using wet ice is an optional step intended to facilitate handling, reduce pup movement, and minimize potential animal discomfort. This cryo-anesthesia step may also provide the benefit of lowering intracranial pressure and reducing volume-related complications9,10. - Move the pups to a separate cage away from the dam while handling them.
- Weigh each pup and document their weight.
- Wipe the back of the mouse using gauze and ethanol. Confirm the intervertebral space or, at a minimum, the midline of the spinal canal (which should appear red in P1 pups) using the dissecting microscope (Supplementary Video 1).
2. Injection procedure
- To anesthetize a single pup, gently place it on a water-proof barrier such as a latex sleeve or aluminum foil on an ice bath for 3-5 min. It's important to avoid leaving the animal on the ice for an extended period, as doing so could pose potential risks of hypothermia-related complications, including ventricular fibrillation, tissue hypoxia, and metabolic acidosis.
NOTE: The duration of 3-5 min may vary on a case-by-case basis. Assess the signs of anesthesia, such as unresponsiveness to a toe pinch, to determine the appropriate duration. - While the animal is on the ice, load the syringe with 10 µL of the drug formulation, virus preparation, or control artificial spinal fluid, etc.
NOTE: During the learning phase, consider the option of injecting the same volume of mixed 1% Fast Green dye with the delivery materials (see Table of Materials). This can aid in visualizing the injection process and assist in learning and refining the technique. Pups injected with Fast Green Dye or similar materials should be euthanized shortly after injection per the approved protocol, as these materials may result in inflammatory reactions or other side effects in animals. - Once the animal is fully anesthetized, as confirmed by reduced or absent body movement, gently position the pups under the microscope.
- With the left index finger and thumb, carefully palpate the intervertebral space along the midline, situated between the bilateral pelvic girdles (Supplementary Video 1). Gently rotate the base of the tail slightly to help identify the midline of the spine.
- Adjust the needle bevel toward the head of the animal before injection.
- Carefully insert the needle, tilting it slightly to an angle of 70°-80° at the point where the indentation intersects, while ensuring the syringe remains aligned with the central sagittal plane. As the needle makes contact with the bone, gradually decrease the angle to approximately 30°, then advance the needle about 2 mm into the intervertebral space.
NOTE: The ability of the needle to slightly lift the entire body is a sign of successful entry into the intradural space. - Slowly inject up to 10 µL volume within 50-60 s. Keep the needle in place for 10-20 s after delivery is completed. Withdraw the needle with a gentle rotation to avoid leaking.
NOTE: The cerebellum will turn green before withdrawing the needle. Also, the slow push is critical to prevent an increase in intracranial pressure associated with delivery and to minimize potential complications. Based on our experience with injections in more than 500 pups, delivering a 10 µL volume over 50-60 s is optimal.
3. Post injection
- Apply a cotton swab to the injection site if there is any leaking or blood.
NOTE: There should not be any in most cases. From our experience, pups treated with traces of leaking or blood are still usable, but consideration of a reduced dose of drugs or treatments may be necessary during data analysis. - Place the pup on a heating pad and allow 10-15 min for the pups to fully recover and re-warm. Carefully observe the pups to ensure they are alert and actively moving before returning them to their home cage. Adequate recovery of a mouse is indicated by the restoration of pink skin color, increased spontaneous body movement, and responsive reactions to touch.
- Place the pup back into the home cage and ensure that the pup is properly covered with bedding, nestlet, or both. This ensures that the pup receives the necessary maternal care from the dam.
- Assess the general appearance and activity daily for at least 3 days post-injection. A sick appearance and reduced activity may raise the possibility of infection, treatment-associated side effects, or other complications, etc. If necessary, consult with veterinary care.
Representative Results
Successful intrathecal injection immediately resulted in the widespread distribution of the administered solution, although the actual cellular penetration depended on the nature of the delivered drugs and materials. In this study, we used Fast Green to visualize the immediate results after intrathecal injection (IT) in wild-type neonates (Figure 1A–K) and compared it with conventional intracerebroventricular (ICV) injection (Figure 1L–N). The long-term results (10 days after injection) were also examined using YFP reporter mice activated by the delivery of CRISPR/Cas9-based gene editing7. YFP expression was observed widely across the entire mouse brain compared to non-CRISPR/Cas9-treated mice (Figure 2). The expression of YFP was observed in the majority of cells under higher magnification. Injections were performed in more than 500 newborn pups, and over 98% of the injected pups survived the procedure. No harmful effects on the long-term survival and health of treated pups were observed (Supplementary Figure 1).
Figure 1: Temporal and spatial distribution of Fast Green dye in mouse brains. (A) Gross observation of mice, comparing injected and non-injected mice 5 min after intrathecal injection. (B) Visualization of the distribution of Fast Green dye in the mouse brain before dissection. (C–E) Distribution of the dye in dissected brains 5 min after intrathecal administration. (F–H) Distribution of the dye in dissected brains 30 min after intrathecal administration. (I–K) Distribution of the dye in dissected brains 60 min after intrathecal administration. (L–N) For comparison, the distribution of the dye in dissected brains 40 min after intracerebroventricular administration. Scale bar: 1 mm. Please click here to view a larger version of this figure.
Figure 2: Long-term effects of intrathecally administered CRISPR/Cas9 gene editing. Widespread expression of the YFP reporter in the mouse brain following intrathecal injection of CRISPR/Cas9 gene editing: cerebellum (A–F), posterior cortex (D–F), and prefrontal cortex (G–I). Scale bar: 50 µm. Please click here to view a larger version of this figure.
Supplementary Figure 1: Survival curve of Angelman Syndrome mice after intrathecal CRISPR gene editing. Survival curve depicting the outcomes of Angelman Syndrome mice receiving CRISPR gene editing through intrathecal administration, compared to non-treated and wild-type mice. Please click here to download this File.
Supplementary Video 1: Intrathecal injection procedure in neonatal mice. Video demonstrating the process of delivering intrathecal injections to neonatal mice. Please click here to download this Video.
Discussion
Described is a step-by-step procedure for intrathecal injection in neonatal mice (P1), resulting in widespread drug distribution in their brains. In comparison to the common intracerebroventricular injection method for delivering medication to neonatal mice, which involves piercing the cerebral cortex11, intrathecal injection avoids direct injury to the neonatal mouse brain due to needle penetration. Due to minimal invasiveness, intrathecal injection can be performed repeatedly when necessary, simulating repeated administrations in humans in a clinical setting12.
Changes in intracranial pressure are commonly associated with intrathecal injection13, potentially leading to the dam’s rejection and refusal to feed due to behavioral changes in the pups. However, acute changes in behavior or reduced survival among injected pups have not been observed. Similarly, unusual or abnormal behaviors associated with intrathecal injections in adults have not been noticed (data not shown).
Several technical tips may have contributed to the success and are worth emphasizing. The slower rate of injection is likely an important factor. Additionally, cryo-anesthesia may reduce intracranial pressure before the injection, minimizing backflow during the intrathecal injection and reducing other complications. Lastly, the precision of the injection site may also impact the success rate.
To achieve the best efficacy via intrathecal administration, performing the procedure as soon as possible after the pups are delivered is critical. Drugs and other substances delivered via intrathecal injection enter the intrathecal space, which is the space between the arachnoid mater and pia mater layers of the meninges that surround the brain and spinal cord. Therefore, the drugs delivered via intrathecal injection pass through these layers of the meninges12. In rodents, as in humans, the meninges consist of three layers: the dura mater, arachnoid mater, and pia mater14. These membranes form during embryonic development and are fully mature by postnatal day 2 (P2)15. Therefore, using a rapid genotyping protocol to assign the pups to experimental groups within a few hours is recommended, especially for experiments involving animal genotypes, such as gene-editing experiments. The earlier the pups are injected, the better the outcome. The injections are typically completed within 3 h after birth. This time window allows the injected drugs to follow the flow of cerebrospinal fluid into the brain parenchyma while the ependymal lining is still immature and less affected by the size of drug particles. It’s worth noting the developmental differences between humans and mice. P1 neonatal mice correspond to the late gestational stage of human brain development16. Results from experiments on P1 neonatal mice serve as valuable proof of concept, but caution should be exercised when extrapolating these results to humans in translational study designs.
This technique is challenged by a constrained administration time window and the requirement for highly skilled experimenters. High mortality may be associated with the procedure if the experimenter lacks experience. However, the narrow timeframe demands a heightened level of precision and repeatability within and between studies. Furthermore, with adequate practice, the proficiency and success rate of this method can be significantly enhanced.
If the injections are performed correctly, the survival of injected pups is primarily affected by maternal care. Preparing foster female pairs for your targets is recommended. If the targeted pups do not have a milk spot on their bellies in the afternoon of the injection day, they should be transferred to the foster female immediately. Female mice recognize their babies by odor. Therefore, it is critical to avoid introducing unfamiliar odors from experimenters or unrelated dams to the pups during and after the procedure. However, the necessity of using foster females should be assessed for individual experiments. Performing the procedure in a well-ventilated room, ideally in a biological fume hood, is also recommended. Mixing the pups with the dam’s bedding and excrement is also helpful. After the initial post-procedure check, minimizing disturbance to the dam for at least 3 days to reduce stress is recommended. Like any surgical procedure, the risk of post-procedure infection should be considered. Thus, strict adherence to good laboratory practices for sterile procedures should be followed during the injection. It should be noted that intrathecal pressure is higher than the external environment, providing natural protection against infection. Experience indicates that the rate of post-injection infection is rare. However, daily monitoring of the general appearance and activity of pups for at least 3 days post-injection to detect signs and symptoms of infection or other complications is recommended. In special cases, consultation with veterinary services may be warranted instead of euthanizing pups with significant complications.
Divulgations
The authors have nothing to disclose.
Acknowledgements
XNL is supported by Foundation for Angelman Syndrome Therapeutic (FAST) Postdoctoral Fellowship. YHJ is also support by FAST and NIH Grant R01HD110195 and R01MH117289.
Materials
| Balance | Ohaus Corporation | 30253017 | |
| C57BL/6J mice | The Jackson Laboratory | 000664 | |
| Digital Microscope | RWD | DOM-1001 | |
| DPBS | ThermoFisher | 14190144 | |
| Fast Green | Sigma | F7252-5G | |
| Heating pad | RWD | 69020 | |
| Needles | Hamilton | 6PK (34/0.375”/4/12DEG)S | |
| Syringe | Hamilton | 1702RN | |
| Syringe Filters | Sigma | SLGVM33RS |
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