A combination of a sensitive blue light-gated channelrhodopsin actuator and red-shifted Arch-based voltage sensors allows all-optical electrophysiology without cross-talk in cultured neurons or brain slices. (...) - by Hochbaum DR et al., Nature Methods 11, 825–833, 22 June 2014
Neuroscience education can be promoted by the availability of low cost and engaging teaching materials. To address this issue, we developed an open-source lipid bilayer amplifier, the OpenPicoAmp, which is appropriate for use in introductory courses in biophysics or neurosciences dealing with the electrical properties of the cell membrane. The amplifier is designed using the common lithographic printed circuit board fabrication process and off-the-shelf electronic components. In addition, we propose a specific design for experimental chambers allowing the insertion of a commercially available polytetrafluoroethylene film. This experimental setup can be used in simple experiments in which students monitor the bilayer formation by capacitance measurement and record unitary currents produced by ionic channels like gramicidin A. Used in combination with a low-cost data acquisition board this system provides a complete solution for hands-on lessons, therefore improving the effectiveness in teaching basic neurosciences or biophysics. - by Shlyonsky V et al., arXiv:1403.7439 2014
The field of optogenetics uses channelrhodopsins (ChRs) for light-induced neuronal activation. However, optimized tools for cellular inhibition at moderate light levels are lacking. We found that replacement of E90 in the central gate of ChR with positively charged residues produces chloride-conducting ChRs (ChloCs) with only negligible cation conductance. Molecular dynamics modeling unveiled that a high-affinity Cl–-binding site had been generated near the gate. Stabilizing the open state dramatically increased the operational light sensitivity of expressing cells (slow ChloC). In CA1 pyramidal cells, ChloCs completely inhibited action potentials triggered by depolarizing current injections or synaptic stimulation. Thus, by inverting the charge of the selectivity filter, we have created a class of directly light-gated anion channels that can be used to block neuronal output in a fully reversible fashion. - by Wietek J. et al., Science 25 April 2014: Vol. 344 no. 6182 pp. 409-412
Julien Hering, PhD's insight:
Silencing Neurons with Light
Neural networks control the activity of living individuals as central processing units control the functions of modern computers. In a neuronal circuit, information is transmitted through neurons in the form of an action potential, which is the electric potential difference between the inside and the outside of a neuron. Ion channel proteins in the neuronal membrane act as molecular devices that create and regulate action potentials. A technology called optogenetics allows neuronal circuits to be manipulated by a combination of optics and genetically targeted incorporation of microbial retinal binding proteins, called opsins, into neurons. On pages 409 and 420 of this Science issue, Wietek et al. use structure-based molecular engineering to invert the charge selectivity of different opsins, channelrhodopsins from algae, resulting in much improved neuron silencers for use in optogenetics.
The advent of scanning two-photon microscopy (2PM) has created a fertile new avenue for noninvasive investigation of brain activity in depth. One principal weakness of this method, however, lies with the limit of scanning speed, which makes optical interrogation of action potential-like activity in a neuronal network problematic. Encoded multisite two-photon microscopy (eMS2PM), a scanless method that allows simultaneous imaging of multiple targets in depth with high temporal resolution, addresses this drawback. eMS2PM uses a liquid crystal spatial light modulator to split a high-power femto-laser beam into multiple subbeams. To distinguish them, a digital micromirror device encodes each subbeam with a specific binary amplitude modulation sequence. Fluorescence signals from all independently targeted sites are then collected simultaneously onto a single photodetector and site-specifically decoded. We demonstrate that eMS2PM can be used to image spike-like voltage transients in cultured cells and fluorescence transients (calcium signals in neurons and red blood cells in capillaries from the cortex) in depth in vivo. These results establish eMS2PM as a unique method for simultaneous acquisition of neuronal network activity. (...) - by Ducros Met al., PNASAugust 6, 2013 vol. 110 no. 32 13138-13143
Freehand injection of recombinant AAV into the neonatal mouse brain offers a fast and easy way to attain widespread genetic manipulation of neurons throughout the brain. Rapid onset and year-long persistence of viral expression permits study of both critical periods and aging. Viral titer can be used to control mosaicism, and multiple viruses can be co-injected for bigenic expression. The technique’s simplicity and the availability of viral reagents should facilitate a range of experiments. (...) - by Ji-Yoen Kim et al., European Journal of Neuroscience, Volume 37, Issue 8, pages 1203–1220, April 2013
Cell culture systems are widely used for molecular, genetic and biochemical studies. Primary cell cultures of animal tissues offer the advantage that specific cell types can be studied in vitro outside of their normal environment. We provide a detailed protocol for generating primary neural cell cultures derived from larval brains of Drosophila melanogaster. The developing larval brain contains stem cells such as neural precursors and intermediate neural progenitors, as well as fully differentiated and functional neurons and glia cells. We describe how to analyze these cell types in vitro by immunofluorescent staining and scanning confocal microscopy. Cell type–specific fluorescent reporter lines and genetically encoded calcium sensors allow the monitoring of developmental, cellular processes and neuronal activity in living cells in vitro. The protocol provides a basis for functional studies of wild-type or genetically manipulated primary neural cells in culture, both in fixed and living samples. The entire procedure takes ∼3 weeks. - by Egger Bet al., Nature Protocols8,958–965(2013)
G protein–coupled receptors (GPCRs), the largest family of membrane signaling proteins, respond to neurotransmitters, hormones and small environmental molecules. The neuronal function of many GPCRs has been difficult to resolve because of an inability to gate them with subtype specificity, spatial precision, speed and reversibility. To address this, we developed an approach for opto-chemical engineering of native GPCRs. We applied this to the metabotropic glutamate receptors (mGluRs) to generate light-agonized and light-antagonized mGluRs (LimGluRs). The light-agonized LimGluR2, on which we focused, was fast, bistable and supported multiple rounds of on/off switching. Light gated two of the primary neuronal functions of mGluR2: suppression of excitability and inhibition of neurotransmitter release. We found that the light-antagonized tool LimGluR2-block was able to manipulate negative feedback of synaptically released glutamate on transmitter release. We generalized the optical control to two additional family members: mGluR3 and mGluR6. This system worked in rodent brain slices and in zebrafish in vivo, where we found that mGluR2 modulated the threshold for escape behavior. These light-gated mGluRs pave the way for determining the roles of mGluRs in synaptic plasticity, memory and disease. - by Levitz J et al., Nature Neuroscience16,507–516(2013)
It is now possible to map the activity of nearly all the neurons in a vertebrate brain at cellular resolution. What does this mean for neuroscience research and projects like the Brain Activity Map proposal?
In an Article that just went live in Nature Methods, Misha Ahrens and Philipp Keller from HHMI’s Janelia Farm Research Campus used high-speed light sheet microscopy to image the activity of 80% of the neurons in the brain of a fish larva at speeds of a whole brain every 1.3 seconds. This represents—to our knowledge—the first technology that achieves whole brain imaging of a vertebrate brain at cellular resolution with speeds that approximate neural activity patterns and behavior. (...) - by erika pastrana, Nature Methods, 18 Mar 2013
Northwestern researchers have developed a novel push-pen patch clamp electrode system that integrates a linear hydraulic actuator in the pipette holder. The actuator moves the metal Ag/AgCl electrode within the pipette to a position where it protrudes from the pipette orifice. This mechanism has multiple benefits in conventional whole-cell experiments. For example, it lowers the series resistance since the resistivity of the electrode is less than that of the pipette solution. The reduced series resistance permits the recording of higher bandwidth signals. Further, the push-pen operation serves as a physical structure to help remove the commonly found cellular debris clog in the pipette tip by pushing it out and clearing it. Lastly, the push-pen operation also reduces the leakage of cytosol into the pipette which results in the ability to conduct longer experiments. (...) - flintbox , Dec 14, 2012
Cellular and subcellular structures in thick biological samples typically are visualized either by genetically encoded fluorescent proteins or by antibody staining against proteins of interest. However, both approaches have drawbacks. Fluorescent proteins do not survive treatments for tissue preservation well, are available in only a few colors, and often emit weak signals. Antibody stainings are slow, do not penetrate thick samples well, and often result in considerable background staining. We have overcome these limitations by using genetically encoded chemical tags that result in rapid, even staining of thick biological samples with high-signal and low-background labeling. We introduce tools for flies and mice that drastically improve the speed and specificity for labeling genetically marked cells in biological tissues.(...) - by Kohl J et al., PNAS, vol. 111 no. 36, E3805–E3814, doi: 10.1073/pnas.1411087111
In this technical report, St-Pierre and colleagues introduce a new genetically encoded voltage sensor called Accelerated Sensor of Action Potentials 1 (ASAP1), which consists of a circularly permuted GFP inserted in the extracellular voltage-sensing domain of a phosphatase. ASAP1 surpasses existing sensors in reliably detecting single action potentials and tracking subthreshold potentials and high-frequency spike trains. (...) - by St-Pierre F. et al., Nature Neuroscience 17, 884–889 (2014)
An understanding of the molecular mechanisms of axon regeneration after injury is key for the development of potential therapies. Single-cell axotomy of dissociated neurons enables the study of the intrinsic regenerative capacities of injured axons. This protocol describes how to perform single-cell axotomy on dissociated hippocampal neurons containing synapses. Furthermore, to axotomize hippocampal neurons integrated in neuronal circuits, we describe how to set up coculture with a few fluorescently labeled neurons. This approach allows axotomy of single cells in a complex neuronal network and the observation of morphological and molecular changes during axon regeneration. Thus, single-cell axotomy of mature neurons is a valuable tool for gaining insights into cell intrinsic axon regeneration and the plasticity of neuronal polarity of mature neurons. Dissociation of the hippocampus and plating of hippocampal neurons takes ∼2 h. Neurons are then left to grow for 2 weeks, during which time they integrate into neuronal circuits. Subsequent axotomy takes 10 min per neuron and further imaging takes 10 min per neuron. - by Gomis-Rüth S et al., Nature Protocols 9, 1028–1037 (2014)
Direct electrical access to presynaptic ion channels has hitherto been limited to large specialized terminals such as the calyx of Held or hippocampal mossy fiber bouton. The electrophysiology and ion-channel complement of far more abundant small synaptic terminals (≤1 μm) remain poorly understood. Here we report a method based on superresolution scanning ion conductance imaging of small synapses in culture at approximately 100–150 nm 3D resolution, which allows presynaptic patch-clamp recordings in all four configurations (cell-attached, inside-out, outside-out, and whole-cell). Using this technique, we report presynaptic recordings of K+, Na+, Cl−, and Ca2+ channels. This semiautomated approach allows direct investigation of the distribution and properties of presynaptic ion channels at small central synapses. - by Novak P.et al., NeuronVolume 79, Issue 6, 18 September 2013, Pages 1067–1077
Julien Hering, PhD's insight:
This semiautomated method seems to be robust and accesible to low experienced scientists in electrophysiology. The nanoscale resolution opens a window on the physiology of small presynaptic boutons and the ion channels distribution and properties in these neuronal terminals. - Article in open access !
Light microscopy can be applied in vivo and can sample large tissue volumes, features crucial for the study of single neurons and neural circuits. However, light microscopy per se is diffraction-limited in resolution, and the substructure of core signaling compartments of neuronal circuits—axons, presynaptic active zones, postsynaptic densities and dendritic spines—can be only insufficiently characterized by standard light microscopy. Recently, several forms of super-resolution light microscopy breaking the diffraction-imposed resolution limit have started to allow highly resolved, dynamic imaging in the cell-biologically highly relevant 10–100 nanometer range ('mesoscale'). New, sometimes surprising answers concerning how protein mobility and protein architectures shape neuronal communication have already emerged. Here we start by briefly introducing super-resolution microscopy techniques, before we describe their use in the analysis of neuronal compartments. We conclude with long-term prospects for super-resolution light microscopy in the molecular and cellular neurosciences.(...) - by Marta Maglione & Stephan J Sigrist, Nature Neuroscience 16, 790–797, 25 June 2013
Synaptic vesicles (SVs) are essential organelles that participate in the release of neurotransmitters from a neuron. Biochemical analysis of purified SVs was instrumental in the identification of proteins involved in exocytotic membrane fusion and neurotransmitter uptake. Although numerous protocols have been published detailing the isolation of SVs from the brain, those that give the highest-purity vesicles often have low yields. Here we describe a protocol for the small-scale isolation of SVs from mouse and rat brain. The procedure relies on standard fractionation techniques, including differential centrifugation, rate-zonal centrifugation and size-exclusion chromatography, but it has been optimized for minimal vesicle loss while maintaining a high degree of purity. The protocol can be completed in less than 1 d and allows the recovery of ∼150 μg of vesicle protein from a single mouse brain, thus allowing vesicle isolation from transgenic mice. (...) - by Ahmed Set al., Nature Protocols8,998–1009(2013)
A study with low statistical power has a reduced chance of detecting a true effect, but it is less well appreciated that low power also reduces the likelihood that a statistically significant result reflects a true effect. Here, we show that the average statistical power of studies in the neurosciences is very low. The consequences of this include overestimates of effect size and low reproducibility of results. There are also ethical dimensions to this problem, as unreliable research is inefficient and wasteful. Improving reproducibility in neuroscience is a key priority and requires attention to well-established but often ignored methodological principles. - By Button KSet al., Nature Reviews Neuroscience14, 365-376(May 2013)
Technique to make tissue transparent offers three-dimensional view of neural networks.
A chemical treatment that turns whole organs transparent offers a big boost to the field of ‘connectomics’ — the push to map the brain’s fiendishly complicated wiring. Scientists could use the technique to view large networks of neurons with unprecedented ease and accuracy. The technology also opens up new research avenues for old brains that were saved from patients and healthy donors. (...) - by Helen Shen, Nature News, 10 April 2013
A new way to link artificial arms and hands to the nervous system could allow the brain to control prostheses as smoothly as if they were natural limbs (...) - By D. Kacy Cullen and Douglas H. Smith, Scientific American, January 14, 2013
by Henneberger C Rusakov DA, Nature Protocols 7, 2171–2179 (2012)
Julien Hering, PhD's insight:
[Abstract] Rapid signal exchange between astroglia and neurons has emerged as a key player in neural communication in the brain. To understand the mechanisms involved, it is often important to have access to individual astrocytes while monitoring the activity of nearby synapses. Achieving this with standard electrophysiological tools is not always feasible. The protocol presented here enables the monitoring of synaptic activity using whole-cell current-clamp recordings from a local astrocyte. This approach takes advantage of the fact that the low input resistance of electrically passive astroglia allows extracellular currents to pass through the astrocytic membrane with relatively little attenuation. Once the slice preparation is ready, it takes ∼30 min to several hours to implement this protocol, depending on the experimental design, which is similar to other patch-clamp techniques. The technique presented here can be used to directly access the intracellular medium of individual astrocytes while examining synapses functioning in their immediate proximity.
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