Neuroscience

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; their in vitro differentiation and dedifferentiation system has not been described in detail in the literature. Thus, an in vitro differentiation and dedifferentiation system of rat Schwann cells is described in this protocol. These cultures and systems may be used to investigate the morphological and biochemical effects of drug interventions or lentivirus-mediated gene transfer on Schwann cells during differentiation or dedifferentiation.

Graphical abstract

The vestibular sensory apparatus contained in the inner ear is a marvelous evolutionary adaptation for sensing movement in 3 dimensions and is essential for an animal’s sense of orientation in space, head movement, and balance. Damage to these systems through injury or disease can lead to vertigo, Meniere’s disease, and other disorders that are profoundly debilitating. One challenge in studying vestibular organs is their location within the boney inner ear and their small size, especially in mice, which have become an advantageous mammalian model. This protocol describes the dissection procedure of the five vestibular organs from the inner ear of adult mice, followed by immunohistochemical labeling of a whole mount preparation using antibodies to label endogenous proteins such as calretinin to label Type I hair cells or to amplify genetically expressed fluorescent proteins for confocal microscopic imaging. Using typical lab equipment and reagents, a patient technician, student, or postdoc can learn to dissect and immunolabel mouse vestibular organs to investigate their structure in health and disease.

Optogenetics has the potential to transform the study of the peripheral nervous system (PNS), but the complex anatomy of the PNS poses unique challenges for the focused delivery of light to specific tissues. This protocol describes the fabrication of a wireless telemetry system for studying peripheral sensory pathways. Unlike existing wireless approaches, the low-power wireless telemetry offers organ specificity via a sandwiched pre-curved tether, and enables high-throughput analysis of behavioral experiments with a channel isolation strategy. We describe the technical procedures for the construction of these devices, the wireless power transmission (TX) system with antenna coils, and their implementation for in vivo experimental applications. In total, the timeline of the procedure, including device fabrication, implantation, and preparation to begin in vivo experimentation can be completed in ~2-4 weeks. Implementation of these devices allows for chronic (>1 month) wireless optogenetic manipulation of peripheral neural pathways in freely behaving animals navigating homecage environments (up to 8).

Peripheral nerve injury (PNI) is common in all walks of life, and the most common PNIs are nerve crush and nerve transection. While optimal functional recovery after crush injury occurs over weeks, functional recovery after nerve transection with microsurgical repair and grafting is poor, and associated with permanent disability. The gold-standard treatment for nerve transection injury is microsurgical tensionless end-to-end suture repair. Since it is unethical to do experimental PNI studies in humans, it is therefore indispensable to have a simple, reliable, and reproducible pre-clinical animal model for successful evaluation of the efficacy of a novel treatment strategy. The objective of this article is two-fold: (A) To present a novel standardized peripheral nerve transection method in mice, using fibrin glue for modeling peripheral nerve transection injury, with reproducible gap distance between the severed nerve ends, and (B) to document the step-wise description of constructing a pressure sensor device for crush injury pressure measurements. We have successfully established a novel nerve transection model in mice using fibrin glue, and demonstrated that this transection method decreases surgical difficulties and variability by avoiding microsurgical manipulations on the nerve, ensuring the reproducibility and reliability of this animal model. Although it is quite impossible to exactly mimic the pathophysiological changes seen in nerve transection with sutures, we hope that the close resemblance of our novel pre-clinical model with gold-standard suturing can be easily reproduced by any lab, and that the data generated by this method significantly contributes to better understanding of nerve pathophysiology, molecular mechanisms of nerve regeneration, and the development of novel strategies for optimal functional recovery. In case of peripheral nerve crush injury, current methods rely on inter-device and operator precision to limit the variation with applied pressure. While the inability to accurately quantify the crush pressure may result in reduced reproducibility between animals and studies, there is no documentation of a pressure monitoring device that can be readily used for real-time pressure measurements. To address this deficit, we constructed a novel portable device comprised of an Arduino UNO microcontroller board and force sensitive resistor (FSR) capable of reporting the real-time pressure applied to a nerve. This novel digital pressure sensor device is cheap, easy to construct and assemble, and we believe that this device will be useful for any lab performing nerve crush injury in rodents.

The thoracic paravertebral sympathetic chain postganglionic neurons (tSPNs) represent the predominant sympathetic control of vascular function in the trunk and upper extremities. tSPNs cluster to form ganglia linked by an interganglionic nerve and receive multisegmental convergent and divergent synaptic input from cholinergic sympathetic preganglionic neurons of the spinal cord (Blackman and Purves, 1969; Lichtman et al., 1980). Studies in the past have focused on cervical and lumbar chain ganglia in multiple species, but few have examined the thoracic chain ganglia, whose location and diminutive size make them less conducive to experimentation. Seminal studies on the integrative properties of preganglionic axonal projections onto tSPNs were performed in guinea pig (Blackman and Purves, 1969; Lichtman et al., 1980), but as mice have become the accepted mammalian genetic model organism, there is need to reproduce and expand on these studies in this smaller model. We describe an ex vivo approach that enables electrophysiological, calcium imaging, and optogenetic assessment of convergence, divergence, and studies on pre- to postganglionic synaptic transmission, as well as whole-cell recordings from individual tSPNs. Preganglionic axonal connections from intact ventral roots and interganglionic nerves across multiple segments can be stimulated to evoke compound action potential responses in individual thoracic ganglia as recorded with suction electrodes. Chemical block of synaptic transmission differentiates spiking of preganglionic axons from synaptically-recruited tSPNs. Further dissection, including removal of the sympathetic chain, enables whole-cell patch clamp recordings from individual tSPNs for characterization of cellular and synaptic properties.