Animals
This study followed some applicable aspects of the PREPARE89 planning guidelines checklist, such as the formulation of the in vivo study, dialogue between scientists and the animal facility, and quality control of the in vivo components in the study. All animals were born, bred and housed at the Karolinska Institutet, Comparative Medicine Biomedicum animal facility (KMB). Mouse brain tissues (postnatal days P0, P2, P5, P7, P10 and P21) were obtained from a mouse line generated by crossing Sox10:cre animals (The Jackson Laboratory, 025807) on the C57BL/6j genetic background with RCE:loxP (eGFP) animals (The Jackson Laboratory, 32037-JAX) on a C57BL/6xCD1 mixed genetic background. Females with a hemizygous cre allele were mated with males lacking the cre allele, while the reporter allele was kept in hemizygosity in both females and males. In the resulting Sox10:cre-RCE:loxP (eGFP) progeny, the entire OL lineage was labelled with eGFP.
None of the experimental animals in this study were subjected to previous procedures before enrolment in the study. All of the animals were free from mouse viral pathogens, ectoparasites, endoparasites and mouse bacterial pathogens. Mice were kept under the following light–dark cycle: dawn, 6:00–7:00; daylight, 7:00–18:00; dusk, 18:00–19:00; night, 19:00–6:00; a maximum number of 5 mice were housed per cage in individually ventilated cages (IVC Sealsafe plus GM500, Tecniplast). General housing parameters such as relative humidity, temperature and ventilation follow the European Convention for the Protection of Vertebrate Animals used for experimental and other scientific purposes treaty ETS 123, Strasbourg 18.03.1996/01.01.1991. In brief, consistent relative air humidity of 55 ± 10% was maintained at 22 °C and the air quality was controlled with the use of stand-alone air handling units supplemented with HEPA filtrated air. Monitoring of husbandry parameters was done using ScanClime (Scanbur) units. Cages contained hardwood bedding (TAPVEI), nesting material, shredded paper, gnawing sticks and card box shelter (Scanbur). The mice received a regular chow diet (CRM(P) SDS and CRM(P), SAFE). Water was provided by using a water bottle, which was changed weekly. The cages were changed every other week. Cage changes were done in a laminar air-flow cabinet (NinoSafe MCCU mobile cage changing unit) with a HEPA H14 EN1822 filter (0.3 μm particle size). Facility personnel wore dedicated scrubs, socks and shoes. Respiratory masks were used when working outside of the laminar air-flow cabinet. Both sexes were included in the study. Randomization was performed to assign samples to time-point groups.
All experimental procedures on animals were performed following the European directive 2010/63/EU, local Swedish directive L150/SJVFS/2019:9, Saknr L150, Karolinska Institutet complementary guidelines for procurement and use of laboratory animals, Dnr. 1937/03-640 and Karolinska Institutet Comparative Medicine veterinary guidelines and plans (version 2020/12/18). The procedures described were approved by the regional committee for ethical experiments on laboratory animals in Sweden (Stockholm Norra Djurförsöksetiska Nämnd, 1995/2019 and 7029/2020).
Tissue and slide preparation
For all postnatal collection points except for P21, we performed a decapitation and extracted the brain, immediately placing the tissue in Tissue Tek O.C.T. compound over a bath of dry ice and 70% ethanol. For all mice older than P21 we anaesthetized the mice and then performed a transcardiac perfusion with gassed, ice-cold artificial cerebral spinal fluid. Next, we decapitated the mice and removed the brain and followed the same embedding procedure as for the neonates.
All tissues were stored at −80 °C until further usage. Fresh 50 × 75 mm (Ted Pella, NC1811932) 0.01% poly-L-lysine-coated (Sigma-Aldrich, P1524-100) slides were prepared and stored at 4 °C and used within 1 week for mounting tissue. For brain tissue sectioning, the tissue was incubated for 30 min in the cryostat chamber at −20 °C. The slides were simultaneously precooled (poly-L-lysine coated for DBiT and charged Superfrost Plus (Eprendia, 4951PLUS-001) for CODEX) and kept in the cryostat for the entire procedure. The sections were cut using an antiroll plate, to a thickness of 10 µm. Tissue collection was done by placing the cold slide in the correct position on top of the tissue then placing over a gloved backhand until the tissue was fully bound. The slide was immediately placed back in the cryostat chamber until sectioning was complete, at which point all slides were stored at −80 °C.
LPC mouse model (1% LPC injection)
Sox10:cre-RCE:loxP (eGFP) mice of mixed sex, at around P85, were used in the LPC study. In brief, under an aseptic technique, mice received an intraperitoneal injection of a non-steroidal anti-inflammatory analgesic (Carprofen; 5 mg kg−1), and were then deeply anaesthetized with isoflurane; an ophthalmic ointment was then applied in both eyes to prevent corneal desiccation, and the mouse was placed over a warm pad for the entire procedure to prevent hypothermia. The site of the incision was shaved and sanitized with 2% chlorhexidine before cutting a 0.5 cm incision with a scalpel to expose the cranium. Using a stereotaxic frame, the rough coordinates of −0.8 mm posterior and 0.8 mm lateral were measured for a skull drilling perforation. After completing, the 1% LPC solution was loaded into a pulled-glass micropipette and the coordinates were recalculated. The micropipette was then slowly inserted 1.3 mm deep (CC below cingulum bundle). The 2 µl injection was performed using an injection speed of 5 nl s−1. After the injection was complete the micropipette was kept in place for an additional 5 min to prevent efflux and then slowly pulled out of the brain. Monitoring of reflexes, respiratory rate and breathing pattern was performed during the entire surgical procedure. The skin was sutured with a non-absorbable suture (Vicryl, Ethicon), and the mouse was observed until regaining full consciousness and mobility. Post-operative care was administered daily for the next 72 h. Mice were euthanized at 5, 10 and 21 days after surgery using the sampling procedure and sample collection method as for the P21 mice (above).
Retro-AAV-eGFP injection
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