吸入麻醉方法 z-bo 2020-10-27 443

　　Introduction: Understanding the mechanisms of brain function requires revealing the structure, connections, and activities of neuronal networks. Recombinant DNA technology has enabled the development of powerful genetic coding tools that can be used for visualization of cell populations, editing of genomic DNA, and tracking of neuronal circuits. These tools can be injected stereotactically into the brain to enter specific brain regions of adult rodents in the form of recombinant viruses. This widely used technique includes surgical opening of the skull before injection. It takes about 40 minutes per animal, but it is very efficient and provides reliable nerve conduction to adult animals. Research using newborn rodents is not so common, but it is equally valuable. It can analyze developmental processes, long-term transgene expression, and use young tissues for sensitive electrophysiological experiments. However, it is complicated by the lack of experimental equipment used by newborns, and the key is the difficulty of anesthesia for these animals. As we all know, neonatal anesthesia is unreliable and uncontrollable, which brings huge welfare challenges to its use. Low temperature is used for anesthesia, and the pups are incubated on ice for about 15 minutes, but the depth of low temperature anesthesia is unknown and difficult to control. It is not clear whether hypothermia produces anesthesia or simply immobilization, and it has been shown that recovering from hypothermia can cause pain and damage. Therefore, according to the 3Rs principle, this method is now restricted. The procedure of inhalation anesthesia for newborn mice was performed at P0-2. Using 3D printing equipment for anesthesia delivery, we detailed the use of isoflurane for controlled maintenance and reliable recovery from inhalation anesthesia. This method can be easily applied in any laboratory, and provides important improvements to the welfare of newborn rodents and promotes their application in a range of scientific disciplines. Using this method, we injected adeno-associated virus (AAV) into the brain of P0-2 mice. A litter of complete postpartum animals can be injected safely and reliably in less than 2 hours, thus providing a fast and effective means for brain region-specific transduction.

　　Stereotactic injection procedure: use a small animal stereotactic frame for injection, and use a Hamilton syringe for virus delivery through a microinjection adapter. It is recommended to use coarse adjustment handles to facilitate virus delivery. The anesthesia mold is located under the frame, and the required height can be reached using a plastic box.

　　Syringe preparation: Use the one-step heating protocol on the pipette to pull the borosilicate glass capillary into the conical glass needle. Use tweezers with a diameter of ∼50 to 80μm to break the glass to form a beveled tip. The size of the tip of each pipette is confirmed under a magnifying glass (for example, using a micro lens). The air in the syringe is eliminated to ensure accurate volume distribution; therefore, the syringe is completely filled with mineral oil. Use a Luer syringe and a 30 G needle to backfill the needle, and use a glass pipette adapter to fix it in the Hamilton syringe and place it on the frame. AAV is inserted into the glass needle through the tip to minimize the amount of wasted virus. A drop of AAV containing PBS is dropped on the secondary membrane below the needle tip, and the Hamilton syringe plunger is retracted using the stereotactic frame to draw the required volume into the needle. The volume of AAV should not exceed the volume of the glass pipette to prevent cross-contamination between experiments. The glass needle should be replaced after injection of 3 animals to prevent blockage or passivation. Changing the needle after each animal requires a lot of preparation time, prolongs the operation time, and has the risk of rejection by the mother, so it is not recommended.

　　Anesthesia induction: Just before starting the operation, move the entire nest (including pups and litter) from the mother cage to an empty cage. The induction of anesthesia is carried out in a small induction chamber, and the flow of oxygen-4% isoflurane is 2L/min for 3-4.5min. After 3 minutes, gently pinch the foot with forceps at 30-second intervals to test the anesthesia state. Then transfer the newborn to an anesthesia mold with 4% isoflurane at a flow rate of 2L/min, and fix it with a tissue covering and microporous tape. In order to minimize the experimenter's exposure to isoflurane and animal allergens, it is recommended to operate on a downward suction workbench.

　　Intracerebral injection: measure the injection position of a specific brain area relative to lambda. Under LED lighting, the pipette is located above the injection site, and the depth axis (z) returns to zero at the skin height. The injection is performed by quickly and firmly lowering the glass needle to penetrate the skin and skull. After penetration, adjust the injection depth to the desired position, and manually control the slow injection of the virus. We use 0.5μL of virus injection, about 1×1012 GC ml-1 per hemisphere. Gently retract the glass pipette from the skull and repeat the injection in the second hemisphere if necessary.

　　Recovery: The total duration of anesthesia is about 6-10 minutes. After the injection, the tail of the animal was marked with a pen to distinguish it from the uninjected animal and returned to the nest with the remaining litters to recover from anesthesia. Subsequent animals were injected according to the same procedure. Once all injections have been completed, check whether the pups have recovered successfully (usually 10-20 min), as determined by spontaneous limb movement. After the animal recovers successfully, it is returned to the mother cage. From setup to recovery, the injection of a litter of 6-8 pups usually takes about 1.5-2 hours.

　　Result: Neonatal inhalation anesthesia induction: We aim to develop a powerful and reliable technique to anesthetize newborn mice as an alternative to hypothermic therapy. We monitored the induction of anesthesia in P0-2 mice using 4% isoflurane in the induction room, which is usually used for anesthesia of adult mice. It takes at least 3 minutes of isoflurane exposure to obtain a reliable and sufficient depth of anesthesia, which is obviously longer than that of adult mice. However, gentle pinching must be used to confirm the duration of each animal, because the exercise level of newborn mice may be very low even without complete induction of anesthesia. We have noticed that the extension of anesthesia time may be related to irreversibility, but for ethical reasons, this has not been experimentally checked. We induce the cub for no more than 4.5 minutes. If the response to pinching is still obvious, the animal will not accept the procedure. With 3 minutes to 4.5 minutes of induction, well-trained users can expect the smallest incidence of non-recovery.

　　Use a custom frame to maintain anesthesia: We designed and produced a 3D printing gas flow device, including a newborn-shaped mold and a nose cone composed of concentric rings for anesthesia delivery. The inflow (central) and outflow (external) paths are connected to standard anesthesia equipment via metal adapters embedded in the frame.

　　Consideration of the age of newborn mice: In order to limit the loss of injected pups, especially due to the cannibalism of female mice, we and others injected with P1 instead of P0, because infant mortality within 24 hours after birth can be avoided . For adult animals, it is necessary to drill a hole in the skull to enter the brain, and at P0−2, a glass injection needle can be used to penetrate the soft skull. This greatly simplifies the procedure, greatly shortens the duration of the test, and allows a complete litter of pups (6-8 pups) to be injected within 1.5-2 hours. It takes about 1.5-2 hours from setting to recovery. Due to the thickening of the skull, this method can only be used for animals below P3.

　　Intracerebral injection for neuron labeling: We use two experimental methods to demonstrate this process. First, we inject AAVs expressing Cre-dependent tdTomato (lox-Stop-lox box) and Cre-EGFP into two discrete brain regions at a ratio of 10000:1 for sparse fluorescent labeling of neuronal populations. Inject 0.5μl of virus mixture into the hippocampus or frontal cortex of each hemisphere, with a total concentration of 1.5×1012 GCml−1. When the P8 brain is dissected, the corresponding brain area has obvious strong positive expression.

　　Intracerebral injection for organotypic tissue preparation: hippocampal organotypic slice culture is a powerful method for in vitro analysis of neuron physiology, but tissue needs to be collected around P7. Neonatal injections can be used for in vivo tissue transduction before harvest, increasing the flexibility of experimental time. This is especially important for genome operations such as Cre-dependent gene resection, which can take several weeks to remove highly abundant or stable target proteins.

　　We have demonstrated the value of intracerebral injection in neonates for in vivo tissue transduction before organotypic culture, and we have adopted this method in the study of synaptic receptors. Our technology provides a fast and effective neuronal transduction program, and the published brain map of newborns will help stereotactic any brain area. This technology may provide an alternative method for the development of neuroscience research (such as in utero electroporation) and has substantial benefits for mouse welfare. The protocols and devices we report have greatly improved the welfare of rodents and can be easily adjusted to increase the use of newborn animals in neuroscience and broader scientific research.