Neuroscience

Sciatic nerve injury is a prevalent traumatic condition that significantly impacts a patient's quality of life. The sciatic nerve compression injury model is among the most commonly utilized models for investigating nerve repair and regeneration. Within this context, the degree III sciatic nerve injury model is frequently employed in scientific research due to its clinical relevance and its suitability for studies focused on functional recovery. However, a standardized approach for accurately assessing the success of constructing the degree III sciatic nerve injury model remains lacking. Traditional macroscopic observation methods exhibit limitations, whereas neurophysiological testing serves as a highly sensitive and objective evaluation technique that can directly reflect changes in nerve conduction function, thus providing reliable quantitative evidence for the successful establishment of the model. This study aims to offer a comprehensive description of the application of neurophysiological techniques in evaluating the construction of the degree III sciatic nerve injury model, thereby ensuring the success of model preparation.

The neuromuscular junction (NMJ) is a peripheral synaptic connection between a lower motor neuron and skeletal muscle fibre that enables muscle contraction in response to neuronal stimulation. NMJ dysfunction and morphological abnormalities are commonly observed in neurological conditions, including amyotrophic lateral sclerosis, Charcot–Marie–Tooth disease, and spinal muscular atrophy. Employing precise and reproducible techniques to visualise NMJs in mouse models of neuromuscular disorders is crucial for uncovering aspects of neuropathology, revealing disease mechanisms, and evaluating therapeutic approaches. Here, we present a method for dissecting the deep lumbrical and flexor digitorum brevis (FDB) muscles of the mouse hind paw and describe the process of whole-mount immunofluorescent staining for morphological analysis of NMJs. Similar whole-mount techniques have been applied to other muscles, such as the diaphragm; however, dense connective tissue in adult samples often impedes antibody penetration. Moreover, large hind limb muscles, including the gastrocnemius and tibialis anterior, are commonly used to examine NMJs but require embedding and cryosectioning. These additional steps increase the complexity and duration of the protocol and can introduce sectioning artefacts, including transection of NMJs and disruption of morphology. Using small hind paw muscles enables whole-mounting, which completely eliminates the requirement for embedding and cryosectioning. As a result, the entire neuromuscular innervation pattern can be visualised, allowing a more accurate assessment of NMJ development, denervation, and regeneration in mouse models of neurological disease and nerve injury, which can be applied across all postnatal ages.

Changes in neuronal conduction are common in disease states affecting peripheral nerves. These alterations can significantly impact nerve function and lead to sensorimotor disabilities. In vivo electromyography recording is a well-established electrophysiological method that has been used for decades to assess sensory and motor functions in the nervous system. Nerve studies are challenging to conduct in vivo in rodents, and the involvement of muscle activity makes it difficult to isolate and assess nerve function independently. This protocol provides a comprehensive guide for accurate ex vivo sciatic nerve dissection and handling from mice. It includes the creation of a three-compartment chamber and the establishment of electrophysiological protocols, which enable differential recordings and the analysis of compound action potentials from various nerve fibers. This setup allows researchers to study the specific effects of drugs and pathologies on nerves from a mechanistic perspective. The setup is a stand-alone apparatus that does not require the use of suction electrodes and the maintenance of negative pressure, which can affect the signal-to-noise ratio and recording stability.

Measuring signal propagation through nerves is a classical electrophysiological technique established decades ago to evaluate sensory and motor functions in the nervous system. The whole-nerve preparation provides a valuable model to investigate nerve function ex vivo; however, it requires specific knowledge to ensure successful and stable measurements. Although the methodology for sciatic nerve recordings has long existed, a method for reliable and long-lasting recordings from myelinated and non-myelinated (nociceptive) fibers still needs to be adapted for pharmacological testing. This protocol takes benefits from epineurium sheath removal for pharmacological tests and provides a detailed description of how to make accurate nerve preparations, from the dissection and handling of nerves to epineurium cleaning, fabrication of adaptable suction electrodes for appropriate fiber stimulation and recordings, setting of electrophysiological protocols for compound action potential (CAP) recordings to distinguish between myelinated and non-myelinated (nociceptive) fibers, and finally to the analysis of the datasets of CAP components. We also demonstrate the feasibility of CAP recordings from individual branches in epineurium-free nerve preparations and provide clues to help retain nerve viability and maintain stable recordings over time. Although a sciatic nerve preparation was used here, the methodology can be applied to other nerve-type preparations.

Key features

• Detailed and simplified protocol for peripheral nerve preparation for recording sensory inputs ex vivo.

• Recordings from myelinated and non-myelinated (nociceptive) fibers can be performed hours after nerve preparation.

• The protocol involves the epineurium removal to facilitate drug permeability into nerve tissue for pharmacological tests.

• The protocol allows physiological and pathological studies (pain/chronic pain conditions).

Graphical overview

Preparation and recordings from the sciatic nerve, including myelinated and non-myelinated (nociceptive) fibers

This paper presents versatile protocols to prepare primary human Schwann cell (hSC) cultures from mature peripheral nervous system tissues, including fascicles from long spinal nerves, nerve roots, and ganglia. This protocol starts with a description of nerve tissue procurement, handling, and dissection to obtain tissue sections suitable for hSC isolation and culturing. A description follows on how to disintegrate the nerve tissue by delayed enzymatic dissociation, plate the initial cell suspensions on a two-dimensional substrate, and culture the primary hSCs. Each section contains detailed procedures, technical notes, and background information to aid investigators in understanding and managing all steps. Some general recommendations are made to optimize the recovery, growth, and purity of the hSC cultures irrespective of the tissue source. These recommendations include: (1) pre-culturing epineurium- and perineurium-free nerve fascicles under conditions of adherence or suspension depending on the size of the explants to facilitate the release of proliferative, in vitro–activated hSCs; (2) plating the initial cell suspensions as individual droplets on a laminin-coated substrate to expedite cell adhesion and thereby increase the recovery of viable cells; and (3) culturing the fascicles (pre-degeneration step) and the cells derived therefrom in mitogen- and serum-supplemented medium to accelerate hSC dedifferentiation and promote mitogenesis before and after tissue dissociation, respectively. The hSC cultures obtained as suggested in this protocol are suitable for assorted basic and translational research applications. With the appropriate adaptations, donor-relevant hSC cultures can be prepared using fresh or postmortem tissue biospecimens of a wide range of types and sizes.

This paper introduces simple analytical methods and bioassays to promptly assess the identity and function of in vitro cultured human Schwann cells (hSCs). A systematic approach is proposed to unequivocally discriminate hSCs from other glial cells, non-glial cells, and non-human SCs (authentication), identify hSCs at different stages of differentiation, and determine whether individual hSCs are proliferative or senescent. Examples of how to use distinct cell-based approaches for quality control and routine troubleshooting are provided to confirm the constitution (identity, purity, and heterogeneity) and potency (bioactivity) of hSC cultures from multiple sources. The bioassays are valuable for rapidly gauging the responses of hSCs to mitogenic and differentiating factors and ascertaining the cells’ basic properties before performing co-culture or cell grafting studies. The assays are image based and use adherent hSCs established in monoculture to simplify the experimental setup and interpretation of results. Finally, all sections contain thorough background information, notes, and references to facilitate decision making, data interpretation, and ad hoc method development for diverse applications.

This manuscript describes step-by-step procedures to establish and manage fresh and cryopreserved cultures of nerve-derived human Schwann cells (hSCs) at the desired scale. Adaptable protocols are provided to propagate hSC cultures through serial passaging and perform routine manipulations such as enzymatic dissociation, purification, cryogenic preservation, live-cell labeling, and gene delivery. Expanded hSCs cultures are metabolically active, proliferative, and phenotypically stable for at least three consecutive passages. Cell yields are expected to be variable as determined by the rate of growth of individual batches and the rounds of subculture. The purity, however, can be maintained high at >95% hSC regardless of passage. The cells obtained in this manner are suitable for various applications, including small drug screens, in vitro modeling of neurodevelopmental processes, and cell transplantation. One caveat of this protocol is that continued expansion of same-batch hSC populations is eventually restricted due to senescence-linked growth arrest.

Different regions of the gastrointestinal tract have specific functions and thus distinct motility patterns. Motility is primarily regulated by the enteric nervous system (ENS), an intrinsic network of neurons located within the gut wall. Under physiological conditions, the ENS is influenced by the central nervous system (CNS). However, by using ex vivo organ bath experiments, ENS regulation of gut motility can also be studied in the absence of CNS influences. The current technique enables the characterisation of small intestinal, caecal, and colonic motility patterns using an ex vivo organ bath and video imaging protocol. This approach is combined with the novel edge detection script GutMap, available in MATLAB, that functions across Windows and Mac platforms. Dissected intestinal segments are cannulated in an organ bath containing physiological saline with a camera mounted overhead. Video recordings of gut contractions are then converted to spatiotemporal heatmaps and analysed using the GutMap software interface. Using data analysed from the heatmaps, parameters of contractile patterns (including contraction propagation frequency and velocity as well as gut diameter) at baseline and in the presence of drugs/treatments/genetic mutations can be compared. Here, we studied motility patterns of female mice at baseline and in the presence of a nitric oxide synthase inhibitor (N_ω-Nitro-L-arginine; NOLA) (nitric oxide being the main inhibitory neurotransmitter of gut motility) to showcase the application of GutMap. This technique is suitable for application to a broad range of animal models of clinical disorders to understand underlying biological pathways contributing to gastrointestinal dysfunction.

Key features

• Enhanced video imaging analysis of gut contractility in rodents using a novel software interface.

• New edge detection algorithm to accurately contour curvatures of the gastrointestinal tract.

• Allows for output of high-resolution spatiotemporal heatmaps across Windows and Mac platforms.

• Edge detection and analysis method makes motility measurements accessible in different gut regions including the caecum and stomach.

Graphical overview

Enhancing axon regeneration is a major focus of peripheral nerve injury research. Although peripheral axons possess a limited ability to regenerate, their functional recovery is very poor. Various activity-based therapies like exercise, optical stimulation, and electrical stimulation as well as pharmacologic treatments can enhance spontaneous axon regeneration. In this protocol, we use a custom-built cuff to electrically stimulate the whole sciatic nerve for an hour prior to transection and repair. We used a Thy-1-YFP-H mouse to visualize regenerating axon profiles. We compared the regeneration of axons from nerves that were electrically stimulated to nerves that were not stimulated (untreated). Electrically stimulated nerves had longer axon growth than the untreated nerves. We detail how variations of this method can be used to measure acute axon growth.

In the peripheral nervous system, Schwann cells are the primary type of glia. This protocol describes an in vitro differentiation and dedifferentiation system for rat Schwann cells. These cultures and systems can be used to investigate the morphological and biochemical effects of pharmacological intervention or lentivirus-mediated gene transfer on the process of Schwann cell differentiation or dedifferentiation.

Graphical abstract