live&#45;cell imaging of primary hippocampal neurons for the measurement of  the effects of pathological proteins on axonal transport

<meta charset="utf8"></head><body>Visualisering av proteiner i primära hippocampusneuroner genom avbildning av levande celler

Neurons depend on the bidirectional transport of cargo along the axon to maintain functional synapses and neural connectivity. Axonal transport deficits are thought to be critical contributors to the pathogenesis of several neurodegenerative diseases, including Alzheimer&#39;s disease (AD) and other tauopathies, Parkinson&#39;s disease, amyotrophic lateral sclerosis, and Huntington&#39;s disease1,2,3. Indeed, pathological modifications to several disease-related proteins negatively affect transport (reviewed in 4). Developing methods to investigate the mechanisms by which pathological proteins exert toxicity in disease is necessary to understand neurodegenerative disorders and identify potential targets for therapeutic intervention.
Several important insights into axon transport, including the discovery of conventional kinesin, the kinase- and phosphatase-dependent pathways regulating motor proteins, and mechanisms by which pathological proteins disrupt the regulation of axon transport, were made using the isolated squid axoplasm model4,5. Perfusion of squid axoplasm with pathological forms of tau protein inhibits anterograde fast axonal transport (FAT), an effect dependent on the exposure of the phosphatase-activating domain of tau, which activates protein phosphatase 1 (PP1)6,7,8. PP1 activates glycogen synthase kinase 3 (GSK3), which in turn phosphorylates kinesin light chains causing the release of cargo. Another pathological protein in AD is amyloid-&#946;. Oligomeric forms of amyloid-&#946; inhibit bidirectional FAT through casein kinase 2, which phosphorylates kinesin light chains9. Furthermore, pathological huntingtin protein harboring a polyglutamine expansion and a familial ALS-linked SOD1 mutant each disrupt axonal transport in the squid axoplasm through c-Jun N-terminal kinase and p38 mitogen-activated protein kinase activity, respectively10,11.
While the squid axoplasm model continues to be a valuable tool in understanding the effects of pathological proteins on axonal transport, limited access to the equipment and specimens prevents it from being more widely used. We developed a transport assay using live-cell confocal microscopy imaging of rodent (mouse and rat) primary neurons. This model represents an easily adaptable and manipulatable mammalian neuron-based approach using widely available cell sources and microscopy systems. For example, a variety of pathological proteins (e.g., harboring disease-related modifications) are expressed to identify how specific modifications of these proteins affect transport in axons. Similarly, a variety of fluorescently-tagged cargo proteins can be used to examine cargo-specific changes. Moreover, the underlying molecular mechanisms are studied relatively easily by targeting expression (i.e., knockdown or overexpression) of selected proteins that may mediate these effects. This method also is easily adapted for primary neurons derived from a wide variety of animal models.
We present a detailed protocol describing the live-cell axon transport assay that was previously used in primary hippocampal neurons to show that mutant tau proteins associated with frontotemporal lobar dementias (FTLD; P301L or R5L tau) increase the pausing frequency of fluorescently-labeled cargo proteins&#160;bidirectionally12. Furthermore, the knockdown of the PP1&#947; isoform rescued the pausing effects12. This provides support for the model of pathological tau-induced disruption that is mediated through aberrant activation of a signaling pathway initiated by PP1 as described above6,7,12. In a separate study, we showed that pseudophosphorylation of tau at S199/S202/T205 (the pathogenic AT8 phosphoepitope relevant to tauopathies) increased cargo pause frequency and anterograde segment velocity. These effects were dependent on the N-terminal phosphatase activating domain of tau13. These examples highlight the utility of this model for identifying the mechanisms of how pathological proteins disrupt axon transport in mammalian neurons.
This paper provides a detailed description of the method beginning with the harvest, dissociation, and culturing of primary mouse hippocampal neurons, followed by the transfection of the neurons with cargo proteins fused with a fluorescent protein, and finally, the live-cell imaging and image analysis approach. We demonstrate how this method is used to study the effects of modified tau on the bidirectional transport of the vesicle-associated protein, synaptophysin, as an example. However, there is flexibility in the pathogenic protein and transport cargo protein of interest, which makes this a versatile approach to study axonal transport.

<meta charset="utf8"></head><body>Författare i fokus: Förstå effekten av patologiska proteiner på axonal transport vid neurodegenerativa sjukdomar

Användning av avbildning av levande celler för att mäta effekterna av patologiska proteiner på axonal transport i primära hippocampusneuroner

Neurons rely on bidirectional transport of cargo along the axon to maintain functional synapsis and neural connectivity. Deficits in transport are thought to be critical contributors to the pathogenesis of several neurodegenerative diseases. We aim to identify mechanisms by which disease-associated modifications to tau protein disrupt axonal transporting.We established that multiple pathological forms of tau protein disrupt normal axonal transport in mammalian neurons. This effect depends on changes to phosphorylation-based signaling pathways that regulate axonal transport. These disruptions represent a potential mechanism of tau-induced neurotoxicity in disease.Modified forms of specific proteins can disrupt normal fast axonal transport across several neurodegenerative diseases. However, we don't fully understand the underlying mechanisms of these effects. This provides a reproducible and simple protocol to identify how expression of disease associated proteins affect fast axonal transport in mammalian neurons.This protocol provides a reproducible assay for axonal transport in mammalian primary neurons, that is easily modifiable to include various cargo proteins along with the expression of proteins of interest with specific pathological modifications. It also allows for the targeted knockdown of other pathway components to test particular mechanistic details.

Avbildning av levande celler av primära hippocampusneuroner för mätning av effekterna av patologiska proteiner på axonal transport

livecell-imaging-primary-hippocampal-neurons-for-measurement-effects

Research

JoVE Journal

Neuroscience

Live&#45;Cell Imaging of Primary Hippocampal Neurons for the Measurement of  the Effects of Pathological Proteins on Axonal Transport

Using Live&#45;Cell Imaging to Measure the Effects of Pathological Proteins on Axonal Transport in Primary Hippocampal Neurons

Bidirectional transport of cargos along the axon is critical for maintaining functional synapses, neural connectivity, and healthy neurons. Axonal transport is disrupted in multiple neurodegenerative diseases, and projection neurons are particularly vulnerable because of the need to transport cellular materials over long distances and sustain substantial axonal mass. Pathological modifications of several disease-related proteins negatively affect transport, including tau, amyloid-&#946;, &#945;-synuclein, superoxide dismutase, and huntingtin, providing a potential common mechanism by which pathological proteins exert toxicity in disease. Methods to study these toxic mechanisms are necessary to understand neurodegenerative disorders and identify potential therapeutic interventions.
Here, cultured primary rodent hippocampal neurons are co-transfected with multiple plasmids to study the effects of pathological proteins on fast axonal transport using live-cell confocal imaging of fluorescently-tagged cargo proteins. We begin with the harvest, dissociation, and culturing of primary hippocampal neurons from rodents. Then, we co-transfect the neurons with plasmid DNA constructs to express fluorescent-tagged cargo protein and wild-type or mutant tau (used as an exemplar of pathological proteins). Axons are identified in live cells using an antibody that binds an extracellular&#160;domain of neurofascin, an axon initial segment protein, and an axonal region of interest is imaged to measure fluorescent cargo transport.
Using KymoAnalyzer, a freely available ImageJ macro, we extensively characterize the velocity, pause frequency, and directional cargo density of axonal transport, all of which may be affected by the presence of pathological proteins. Through this method, we identify a phenotype of increased cargo pause frequency associated with the expression of pathological tau protein. Additionally, gene-silencing shRNA constructs can be added to the transfection mix to test the role of other proteins in mediating transport disruption. This protocol is easily adaptable for use with other neurodegenerative disease-related proteins and is a reproducible method to study the mechanisms of how those proteins disrupt axonal transport.

Here, we demonstrate how to combine transfection of primary hippocampal rodent neurons with live-cell confocal imaging to analyze pathological protein-induced effects on axonal transport and identify mechanistic pathways mediating these effects.

Bidirectional transport of cargos along the axon is critical for maintaining functional synapses, neural connectivity, and healthy neurons. Axonal ...

Harvesting of Primary Hippocampal Neurons for Live&#45;Cell Imaging

Primary Hippocampal Neuron Dissociation and Plating for the Measurement of  the Effects of Pathological Proteins on Axonal Transport

Begin by dissecting the hippocampus from an embryonic mouse brain and place the dissected tissue in ice cold calcium and magnesium-free buffer or CMF. Next, remove the CMF and rinse the tissue four times with fresh cold CMF, swirling the tube gently in between rinses. Then add filtered 0.125%trypsin solution and incubate for 15 minutes at 37 degrees Celsius, swirling every five minutes.After incubation, remove the trypsin solution and wash two times with ice cold CMF. Then add three milliliters of trypsin inactivation solution. Next, to dissociate the cells, titrate 30 times using two second draws with a 14-gauge needle attached to a three milliliter syringe.Continue titration with two second draws 30 times using a 15-gauge needle, then 20 times using a 16-gauge needle, followed by 20 times with an 18-gauge needle. Then perform three second draws using a 21-gauge needle 15 times to complete cell dissociation. Check for cell clumps by placing a droplet of the cell suspension on a microscope slide.Next, filter five milliliters of FBS using a 0.22 micron syringe filter. Gently layer the cell suspension onto the FBS in a 15 milliliter conical tube. Centrifuge the cells at 200 x g at four degrees Celsius for five minutes.Remove FBS and resuspend the pellet in one milliliter of warm antibiotic antibiotic-free neurobasal media. Dilute the cell suspension in 0.4%trypan blue and obtain a cell count. Plate the cells in 750 microliters of antibiotic-free neurobasal media in a prewarmed poly-D-lysine coated 4-well chamber slide.Incubate the cells in a humidity controlled chamber at 37 degrees Celsius and 5%CO2.

Primär dissociation och plätering av hippocampusneuroner för mätning av effekterna av patologiska proteiner på axonal transport

Prepare for imaging using a confocal microscope two days after transfection of primary hippocampal neurons. Warm the live cell chamber attachment 60x subjective and lens oil to 37 degrees Celsius and equilibrate the chamber to 5%CO2. Using the eye pieces, identify transfected neurons in the channel containing the cargo protein.Next, click on scan to image the neurons and identify the axon initial segment of the SCI-5 channel. Set the field of view to a rectangular box of 32 by 128 pixels and move it to image a region of the axon, 50 to 150 microns distal to the axon initial segment. Record the directionality of the cargo transport and the orientation of the ROI, then press save as to save it as file.Note that the dots on the box will appear at the top and right in subsequent images. Record the side of the image closest to the cell body and the side closest to the axon terminus to ensure correct orientation of the comagraph Polyline. Set the 561 laser power to 1.40, the pinhole to 7.8, the size to 128 pixels square, and the speed to 32 frames per second.Adjust the gain to visualize the transport of individual cargo vesicles without being obscured by background signal. Select draw rectangular ROI from the ROI dropdown menu and draw the ROI so that it covers the entire field of view. Then right click and select use as stimulation ROI S1.Then in the ND setup tab, click on acquire image to collect the pre stimulation reference image.Then set stimulation to ROI S1, interval to no delay, duration of 1.64 seconds, and loops to seven. Then click on acquire image to collect post stimulation reference image. Select waiting, no acquisition, two minutes to provide a recovery period for fluorescent cargo to repopulate the bleached ROI.Select acquisition interval, no delay, duration five minutes, loops, 8, 615. Click the apply stimulation settings button on the ND setup tab, followed by run now. Also collect transport videos from at two neurons for analysis.Reset the field of view to the entire image and then collect an image of the full neuron in the 561 and 647 channels with the ROI box included. Record the XY coordinates where the image was taken for post fixation confirmation of protein expression. Live cell imaging of a transfected neuron expressing the cargo protein mAPL's Synaptophysin was performed using this protocol.The external domain of neuro fasin in the Axon initial segment was also labeled. Imaging of cargo transport was performed in the axonal region of interest. Further post imaging immunofluorescence analysis confirmed the co-expression of tau protein and mAPL Synaptophysin.