神经科学

Since the discovery that astrocytes are characterized by Ca²⁺-based excitability, investigating the function of these glial cells within the brain requires Ca²⁺ imaging approaches. The technical evolution from chemical fluorescent Ca²⁺ probes with low cellular specificity to genetically encoded indicators (GECIs) has enabled detailed analysis of the spatial and temporal features of intracellular Ca²⁺ signal. Different imaging methodologies allow the extraction of distinct information on calcium signals in astrocytes from brain slices, with resolution ranging from cell populations to single cells up to subcellular domains.

Here, we describe 2-photon laser scanning microscopy (2PLSM) Ca²⁺ imaging in astrocytes from the somatosensory cortex (SSCx) of adult mice in ex vivo acute cortical slices, performed using two genetically encoded Ca²⁺ indicators, i.e., cytosolic GCaMP6f and endoplasmic reticulum-targeted G-CEPIA1er. The main advantage of the 2PLSM technique, compared to single-photon microscopy, is the possibility to go deeper in the tissue while avoiding photodamage, by limiting laser excitation to a single focal plane. The fluorescent signal of the indicator is analyzed offline in different compartments—soma, proximal processes, and microdomains—for GCaMP6f experiments and in the perinuclear, somatic area for G-CEPIA1er. The analysis of Ca²⁺ signal from different compartments, although not providing a value of absolute concentration, allows a critical comparison of the degree of astrocyte activation between different experimental conditions or mouse models. Moreover, the analysis of G-CEPIA1er signal, which reveals metabotropic receptor activation as a dynamic decrease in free Ca²⁺ in the endoplasmic reticulum (ER), can provide information on possible alterations in this critical second messenger pathway in astrocytes, including, for example, steady-state ER Ca²⁺ levels and kinetics of Ca²⁺ release.

Understanding dendritic excitability is essential for a complete and precise characterization of neurons’ input-output relationships. Theoretical and experimental work demonstrates that the electrotonic and nonlinear properties of dendrites can alter the amplitude (e.g., through amplification) and latency of synaptic inputs as viewed in the axosomatic region where spike timing is determined. The gold-standard technique to study dendritic excitability is using dual-patch recordings with a high-resistance electrode used to patch a piece of distal dendrite in addition to a somatic patch electrode. However, this approach is often impractical when distal dendrites are too fine to patch. Therefore, we developed a technique that utilizes the expression of Channelrhodopsin-2 (ChR2) to study dendritic excitability in acute brain slices through the combination of a somatic patch electrode and optogenetic activation. The protocol describes how to prepare acute slices from mice that express ChR2 in specific cell types, and how to use two modes of light stimulation: proximal (which activates the soma and proximal dendrites in a ~100 µm diameter surrounding the soma) with the use of a high-magnification objective and full-field stimulation through a low-magnification objective (which activates the entire somato-dendritic field of the neuron). We use this technique in conjunction with various stimulation protocols to estimate model-based spectral components of dendritic filtering and the impact of dendrites on phase response curves, peri-stimulus time histograms, and entrainment of pacemaking neurons. This technique provides a novel use of optogenetics to study intrinsic dendritic excitability through the use of standard patch-clamp slice physiology.

Neurons are complex cells with two distinct compartments: the somatodendritic and the axonal domains. Because of their polarized morphology, it is challenging to study the differential cellular and molecular mechanisms that occur in axons and impact the soma and dendrites using conventional in vitro culture systems. Compartmentalized cultures offer a solution by physically and chemically separating the axonal from the somatodendritic domain of neurons. The microfluidic chamber model presented in this work is valuable for studying these mechanisms in primary cortical cultures derived from rat and mouse. In addition, this chamber model is compatible with various microscopy methods, such as phase contrast, and fluorescence imaging of living and fixed cells.

Key features

• Preparation and attachment of PDMS microfluidic chambers to glass coverslips.

• Primary culture of cortical neurons and plating cortical neurons in microfluidic chamber.

• Confirmation of compartmentalization using the retrograde transport of the fluorescently labeled form of cholera toxin subunit B (f-Ctb).

• Immunofluorescence and multilabeling of compartmentalized cortical neurons.

• Retrograde transport of fluorescently labeled BDNF.

The activity of numerous autophagy-related proteins depends on their phosphorylation status, which places importance on understanding the responsible kinases and phosphatases. Great progress has been made in identifying kinases regulating autophagy, but much less is known about the phosphatases counteracting their function. Genetic screens and modern proteomic approaches provide powerful tools to identify candidate phosphatases, but further experiments are required to assign direct roles for candidates. We have devised a novel protocol to test the role of purified phosphatases in dephosphorylating specific targets in situ. This approach has the potential to visualize context-specific differences in target dephosphorylation that are not easily detected by lysate-based approaches such as Western blots.

Graphical abstract:

G-protein coupled signaling pathways are organized into multi-protein complexes called signalosomes that are located within and on cellular membranes. We describe the use of silica nanoparticles coated with a unilamellar phospholipid bilayer (lipobeads) to reconstitute the activated photoreceptor G-protein α-subunit (Gtα*) with its cognate effector (phosphodiesterase-6; PDE6) for biochemical and structural studies of the activation mechanism regulating this GPCR signaling pathway. Lipobeads are prepared by resuspending dried-down phospholipid mixtures with monodisperse 70 nm silica particles, followed by extrusion through a 100 nm membrane filter. This uniform and supported liposomal preparation is easily sedimented, permitting the separation of soluble from membrane-associated proteins. Upon loading lipobeads with Gtα* and PDE6, we find that activation of PDE6 catalysis by Gtα* occurs much more efficiently than in the absence of membranes. Chemical cross-linking of membrane-confined proteins allows detection of changes in protein-protein interactions, resulting from G-protein activation of PDE6. The advantages of using lipobeads over partially purified membrane preparations or traditional liposomal preparations are generally applicable to the study of other membrane-confined signal transduction pathways.

The functional performance of a cell depends on how macromolecules, in particular proteins, come together in a precise orientation, how they assemble into protein complexes and interact with each other. In order to study protein-protein interactions at a molecular level, a variety of methods to investigate these binding processes yield affinity constants and/or the identification of binding regions. There are several well-established biophysical techniques for biomolecular interaction studies, such as fluorescence spectroscopy and surface plasmon resonance. Although these techniques have been proven to be efficient, they either need labeling or immobilization of one interaction partner. Backscattering interferometry (BSI) is a label-free detection method, which allows label- and immobilization-free interaction analysis under physiologically relevant conditions with high sensitivity and in small volumes. We used BSI to measure the interaction of the neuronal calcium sensor recoverin with its target G protein-coupled receptor kinase 1 (GRK1) as a model system. Increasing concentrations of purified recoverin were mixed with a specific concentration of a GRK1 fusion protein. In this protocol, we provide a full description of the instrumental setup, data acquisition, and evaluation. Equilibrium dissociation constants of recoverin-GRK1 interaction determined by the BSI instrumental setup are in full agreement with affinity constants obtained by different methods as described in the literature.

Calcium ions trigger many cellular events, including the release of neurotransmitters at the synaptic terminal and excitotoxic cell death. Recently, we have discovered that a transient increase in the level of cytoplasmic Ca²⁺ triggers the exposure of phosphatidylserine (PS) on the surfaces of necrotic cells in the nematode Caenorhabditis elegans. PS serves as an “eat me” signal that attracts engulfing cells to engulf and degrade necrotic cells. During the above study, we developed a microscopic imaging protocol for real-time monitoring the levels of cytoplasmic Ca²⁺ and cell surface PS in Caenorhabditis elegans touch neurons. Previously, Ca²⁺ dynamics was monitored in neurons in Caenorhabditis elegans larvae in time periods ranging from milliseconds to seconds. Methods for monitoring Ca²⁺ dynamics for a relatively long period of time during embryonic development were not available, let alone for simultaneous monitoring Ca²⁺ and PS dynamics. The protocol reported here utilizes a deconvolution imaging system with an optimized experimental setting that reduces photo-damage and allows the proper development of embryos during the real-time imaging process. This protocol enables the simultaneous measurement of cytosolic Ca²⁺ and cell surface PS levels in necrotic touch neurons during embryonic development in a period longer than six hours. Our method provides an easy and sensitive approach to perform long-time Ca²⁺ and PS recording in living animals, simultaneously or individually. This protocol can be applied to study various cellular and developmental events that involve the dynamic regulation of Ca²⁺ and/or PS.

Computational neuroscience aims to model, reproduce, and predict network dynamics for different neuronal ensembles by distilling knowledge derived from electrophysiological and morphological evidence. However, analyses and simulations often remain critically limited by the sparsity of direct experimental constraints on essential parameters, such as electron microscopy and electrophysiology pair/multiple recording evidence of connectivity statistics. Notably, available data are particularly scarce regarding quantitative information on synaptic connections among identified neuronal types. Here, we present a user-friendly data-driven pipeline to estimate connection probabilities, number of contacts per connected pair, and distances from the pre- and postsynaptic somas along the axonal and dendritic paths from commonly available two-dimensional tracings and other broadly accessible measurements. The described procedure does not require any computational background and is accessible to all neuroscientists. This protocol therefore fills the important gap from neuronal morphology to circuit organization and can be applied to many different neural systems, brain regions, animal species, and data sources.

Graphic abstract:

The processing protocol from 2D reconstructions to quantitated synaptic connections

Protein N-glycosylation plays a vital role in diverse cellular processes, and dysregulated N-glycosylation is implicated in a variety of human diseases including neurodegenerative disorders and cancer. With recent advances in high-resolution mass spectrometry-based glycoproteomics technologies enabling large-scale N-glycoproteome profiling of disease and control samples, analysis of the large datasets has become a challenge. Here, we provide a protocol for the systems-level analysis of in vivo N-glycosylation sites on N-glycosylated proteins and their changes in human disease, such as Alzheimer's disease. The protocol includes quantitation and differential analysis of N-glycopeptide abundance, in addition to integrative N-glycoproteome and proteome data analyses, to determine disease-associated changes in N-glycosylation site occupancy and identify differentially N-glycosylated proteins in human disease versus control samples. This protocol can be modified and applied to study proteome-wide N-glycosylation alterations in response to different cellular stresses or pathophysiological states in other organisms or model systems.

Over the last decade, lipids have emerged as possessing an ever-increasing number of key functions, especially in membrane trafficking. For instance, phosphatidic acid (PA) has been proposed to play a critical role in different steps along the secretory pathway or during phagocytosis. To further investigate in detail the precise nature of PA activities, we need to identify the organelles in which PA is synthesized and the PA subspecies involved in these biological functions. Indeed, PA, like all phospholipids, has a large variety based on its fatty acid composition. The recent development of PA sensors has helped us to follow intracellular PA dynamics but has failed to provide information on individual PA species. Here, we describe a method for the subcellular fractionation of RAW264.7 macrophages that allows us to obtain membrane fractions enriched in specific organelles based on their density. Lipids from these membrane fractions are precipitated and subsequently processed by advanced mass spectrometry-based lipidomics analysis to measure the levels of different PA species based on their fatty acyl chain composition. This approach revealed the presence of up to 50 different species of PA in cellular membranes, opening up the possibility that a single class of phospholipid could play multiple functions in any given organelle. This protocol can be adapted or modified and used for the evaluation of other intracellular membrane compartments or cell types of interest.