This study was supported by the German Research Foundation (HA 7728/2-1 and EXC2145 Project ID 390857198) and the Max Planck Society.

All animal experiments were performed following the relevant guidelines and regulations of the Government of Upper Bavaria. The primary neurons were prepared from E16.5 C57BL/6 wild-type mouse embryos of both sexes following standard methods as previously described6.
1. Assembly of the microfluidic device
<ol>
	<li>Coat one six-well glass-bottom tissue culture plate with a final concentration of 20 &#181;g/mL of Poly-D-Lysine and 3.4 &#181;g/mL of Laminin in PBS (phosphate-buffered saline) (see Table of Materials).</li>
	<li>Incubate the coated plate overnight in the dark at room temperature. 
	NOTE: Coating can also be performed at 37 &#176;C for 1-2 h. The choice of the coating depends on the cell type used.</li>
	<li>After coating, bring the six-well plates to the sterile hood and wash them twice with sterile ddH2O. 
	NOTE: Do not wash with salt-containing buffers such as PBS, as salt crystals will interfere with seal formation.</li>
	<li>Allow the plate to dry in a tilted position for 3-5 min. Remove excess water by vacuum suction or pipetting.</li>
	<li>Soak the microfluidic chamber in 80% ethanol. 
	NOTE: The microfluidic chamber was obtained from commercial sources (see Table of Materials).</li>
	<li>Allow the microfluidic chamber to dry for 3-5 min in a tilted position. Remove excess ethanol by vacuum, suction, or pipetting.</li>
	<li>When completely dry, place the microfluidic chamber in the center of the well. 
	NOTE: The formation of the seal can be observed as a change in reflective properties upon exclusion of air from the interface between the plate and the silicone device. Both the plate and microfluidic chamber must be completely dry before assembly to create a good seal. However, drying time needs to be minimized as much as possible to ensure proper adherence to the cells. Therefore, the wells and chambers must be visually inspected for remaining liquid droplets. If no droplets are visible, immediately assemble the chambers.</li>
	<li>Gently tap the microfluidic chamber at its borders and microgroove section in the middle for proper attachment to the glass plate.</li>
</ol>
2. Seeding and maintaining of neurons
<ol>
	<li>Collect the desired number of dissociated hippocampal neurons per chamber in a 1.5 mL reaction tube. 
	NOTE: For the present study, 1.5 x 105 neurons were seeded per device for imaging-based applications.</li>
	<li>Centrifuge neurons at 1000 x g for 4 min at room temperature.</li>
	<li>Discard the supernatant with a pipette and resuspend the pellet in 8 &#181;L of B27-Neurobasal media (see Table of Materials).</li>
	<li>Pipette the cell solution into the channel entrance of the microfluidic chamber (b in Figure 1A).</li>
	<li>Tap on the back of the plate to assist flow through the channel. 
	NOTE: Tapping can be done quite forcefully without worry that the seal will break.</li>
	<li>Aspirate with a pipette any remaining cell suspension at the exit of the channel (g in Figure 1A) to decrease the number of cells outside the channel.</li>
	<li>Incubate the microfluidic chamber with cells for 15-20 min at 37 &#176;C and 5% CO2.</li>
	<li>Fill the top axonal well with 50 &#181;L of B27-Neurobasal media in (c in Figure 1A).</li>
	<li>Tap on the back of the plate to assist flow through the channel.</li>
	<li>Fill wells on the soma side (a and h in Figure 1A) with 150 &#181;L of B27-Neurobasal media each, creating a tension bubble on top. The volume of the bubble is about 20 &#181;L. 
	NOTE: Creating a tension bubble is not necessary to achieve fluidic isolation, but it provides a clear visual of the higher volume on the soma side.</li>
	<li>Fill both wells of the axonal side (c and i in Figure 1A) with 100 &#181;L of B27-Neurobasal media each. 
	NOTE: To promote axonal growth through the microgrooves, it is recommended that the wells on the soma side contain more media than the wells on the axon side. However, axons will by chance grow through the microgrooves even without volume differences.</li>
	<li>Incubate the microfluidic chamber at 37 &#176;C and 5% CO2 for 7-8 days. 
	NOTE: Growth of axons into the axonal chamber can usually be observed starting at day in vitro (DIV) 5. Longer culturing times are possible if more mature cultures are desired.</li>
	<li>Feed neurons every 2-3 days by removing the medium from the two upper wells (a and c) and replacing it with fresh B27-Neurobasal medium until a tension bubble is formed.</li>
</ol>
3. Staining with mitochondrial membrane potential sensitive dye
<ol>
	<li>At DIV 7-8, assess that the axons have grown through the microgrooves and extend into the axonal compartment underneath a light microscope. 
	NOTE: Different DIV stages are also possible depending on the desired age.</li>
	<li>Perform two washes with pre-warmed imaging medium (37 &#176;C) by removing the B27-Neurobasal medium from all wells and adding about 100 &#181;L of the imaging medium to the top wells of both the axonal and soma channels (a and c). Let the medium flow through to the lower wells (h and i).
	<ol>
		<li>Remove the medium from the lower wells and any leftover medium in the top wells, and repeat with fresh medium, leaving the wells empty at the end of the wash. 
		NOTE: Due to the high capillary force in the channels (d and f), there will be a remaining wash medium in the channels that should not be removed to prevent the neurons from drying. Any phenol-red-free imaging medium can be used here, for instance, Hibernate E (see Table of Materials). If the microscope is not set up with a CO2 supply, ensure that the imaging medium uses an alternative buffering system to carbonate/CO2 (e.g., HEPES)8.</li>
	</ol>
	</li>
	<li>Dilute 1 mM tetramethylrhodamine ethyl ester (TMRE, red, see Table of Materials) stock to 5 nM in the imaging medium. 
	NOTE: Do not exceed 1% DMSO in the final dilution to avoid toxicity.</li>
	<li>Add 100 &#181;L of the 5 nM TMRE dilution to both the somatic and axonal top wells (a and c), and let the medium flow through both the somatic and axonal compartments until there is an equal volume in all wells. Fill up with the TMRE-containing medium.</li>
	<li>Place the neurons back into a controlled environment chamber (37 &#176;C and 5% CO2) for 25 min.</li>
	<li>Perform two washes with the pre-warmed imaging medium (37 &#176;C) as described before (step 3.2). On the last wash, fill up with 100 &#181;L in each well and create a tension bubble of the medium on the somatic wells (a and c).</li>
</ol>
4. Live-cell imaging
NOTE: Still images shown were acquired on a spinning disk confocal microscope, using a 40x NA 1.25 immersion objective (see Table of Materials). 200 ms exposure time and 10% laser power for the red channel and 500 ms exposure time for brightfield were chosen. However, regular confocal or widefield inverted microscopes can also be used to study TMRE intensity.
<ol>
	<li>Image the cells with an inverted microscope.</li>
	<li>Ensure that the microscope is equipped with a stage incubator to maintain the neuronal culture at 37 &#176;C throughout the experiment.</li>
	<li>Select a region of interest in the somatic compartment, and follow it through the microgrooves into the axonal compartment. Ensure that the microgrooves imaged match those in the somatic and the axonal compartments (for easier baseline and post-treatment comparison).</li>
	<li>Capture fluorescence images using excitation at 561 nm and emission at 625 nm for red fluorescence signal.</li>
	<li>Acquire the images.</li>
	<li>To induce mitochondrial depolarization, add 20 &#181;M Antimycin A (see Table of Materials) in the imaging medium to the axonal compartment. To ensure a volume difference between the soma and axonal chambers, check the presence of a tension bubble on the somatic side (equals volume &#62;110 &#181;L).
	<ol>
		<li>Remove all imaging medium from the lower well of the axonal side (i), except for the medium inside the channel (f). Add 160 &#181;L of 20 &#181;M Antimycin A to the top axonal well (c) and let it flow through the channel until there is an equal volume in both axonal wells. Volumes in axonal wells are roughly 80 &#181;L per well. 
		NOTE: Ensure that the volume in the axonal wells is smaller than in the somatic wells to ensure proper fluid pressure. A difference of at least 10 &#181;L (10% of the well volume) is recommended, but also higher differences are possible.</li>
	</ol>
	</li>
	<li>Incubate neurons in the controlled environment chamber (37 &#176;C) for 20-30 min.</li>
	<li>Repeat imaging as described above (steps 4.3-4.5). Ensure that the same positions are imaged (easily identified by the microgrooves) as during the baseline acquisition.</li>
</ol>

<ol>
	<li>Murali Mahadevan, H., Hashemiaghdam, A., Ashrafi, G., Harbauer, A. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondria+in+neuronal+health:+from+energy+metabolism+to+Parkinson's+disease.">Mitochondria in neuronal health: from energy metabolism to Parkinson's disease.</a> Advanced Biology. 5 (9), 2100663 (2021).</li><li>Dauer, W., Przedborski, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Parkinson's+disease:+mechanisms+and+models.">Parkinson's disease: mechanisms and models.</a> Neuron. 39 (6), 889-909 (2003).</li><li>Moratalla, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+vulnerability+of+primate+caudate-putamen+and+striosome-matrix+dopamine+systems+to+the+neurotoxic+effects+of+1-methyl-4-phenyl-1,2,3,6-+tetrahydropyridine.">Differential vulnerability of primate caudate-putamen and striosome-matrix dopamine systems to the neurotoxic effects of 1-methyl-4-phenyl-1,2,3,6- tetrahydropyridine.</a> Proceedings of the National Academy of Sciences. 89 (9), 3859-3863 (1992).</li><li>Cheng, H. -. C., Ulane, C. M., Burke, R. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Clinical+progression+in+Parkinson+disease+and+the+neurobiology+of+axons.">Clinical progression in Parkinson disease and the neurobiology of axons.</a> Annals of Neurology. 67 (6), 715-725 (2010).</li><li>Taylor, A. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+microfluidic+culture+platform+for+CNS+axonal+injury,+regeneration+and+transport.">A microfluidic culture platform for CNS axonal injury, regeneration and transport.</a> Nature Methods. 2 (8), 599-605 (2005).</li><li>Harbauer, A. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuronal+mitochondria+transport+Pink1+mRNA+via+synaptojanin+2+to+support+local+mitophagy.">Neuronal mitochondria transport Pink1 mRNA via synaptojanin 2 to support local mitophagy.</a> Neuron. 110 (9), 1516-1531 (2022).</li><li>Ashrafi, G., Schlehe, J. S., LaVoie, M. J., Schwarz, T. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitophagy+of+damaged+mitochondria+occurs+locally+in+distal+neuronal+axons+and+requires+PINK1+and+Parkin.">Mitophagy of damaged mitochondria occurs locally in distal neuronal axons and requires PINK1 and Parkin.</a> Journal of Cell Biology. 206 (5), 655-670 (2014).</li><li>Shipman, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+4-(2-hydroxyethyl)-1-piperazine&#235;thanesulfonic+acid+(HEPES)+as+a+tissue+culture+buffer.">Evaluation of 4-(2-hydroxyethyl)-1-piperazine&#235;thanesulfonic acid (HEPES) as a tissue culture buffer.</a> Proceedings of the Society for Experimental Biology and Medicine. 130 (1), 305-310 (1969).</li><li>Harbauer, A. B., Schneider, A., Wohlleber, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+mitochondria+by+single-organelle+resolution.">Analysis of mitochondria by single-organelle resolution.</a> Annual Review of Analytical Chemistry. 15, 1-16 (2022).</li><li>Taylor, A. M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Axonal+mRNA+in+uninjured+and+regenerating+cortical+mammalian+axons.">Axonal mRNA in uninjured and regenerating cortical mammalian axons.</a> The Journal of Neuroscience. 29 (15), 4697-4707 (2009).</li><li>Altman, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Axonal+TDP-43+condensates+drive+neuromuscular+junction+disruption+through+inhibition+of+local+synthesis+of+nuclear+encoded+mitochondrial+proteins.">Axonal TDP-43 condensates drive neuromuscular junction disruption through inhibition of local synthesis of nuclear encoded mitochondrial proteins.</a> Nature Communications. 12 (1), 1-17 (2021).</li></ol>

The authors declare no competing interests.

The present protocol describes a method to seed and culture dissociated hippocampal neurons in a microfluidic device to treat axonal mitochondria separately. The utility of this approach with the membrane-sensitive dye TMRE and the complex III inhibitor Antimycin A (as previously demonstrated7) is demonstrated here, but this method can be easily adapted to other mitochondrial dyes or genetically encoded sensors of mitochondrial functions that allow local, microscopy-based readouts<sup class='xref'...

Mitochondria are the main suppliers of ATP (adenosine triphosphate) in neurons. As neuronal health is intimately linked to mitochondrial function, it is not surprising that dysfunctional regulation of these organelles has been associated with the onset of various neurodegenerative diseases, including Parkinson&#39;s disease1. Furthermore, mitochondrial intoxication has successfully been used to model Parkinsonian symptoms in animals2. In both animal models and human disease, the demise of neurons starts at the distal parts3,4, hinting that axonal mitochondria might be more susceptible to insults. However, the biology of mitochondria in axons is not well understood due to the difficulties associated with targeted treatment and analysis of axonal mitochondria without simultaneous disturbance of cell body processes.
Recent advances in culturing techniques of dissociated neurons in vitro now allow the fluidic separation of axons and cell bodies through microfluidic devices5. As depicted in Figure 1A, these devices feature four access wells (a/h and c/i), with two channels connecting each pair (d and f). The large channels are connected with each other by a series of 450 &#181;m long microchannels (e). Intentional differences in the fill levels between the two chambers create a fluid pressure gradient (Figure 1B) that prevents the diffusion of small molecules from the channel with a lower fluid level to the other side (Figure 1C, illustrated with Trypan blue dye).
We recently used microfluidic devices to study local translation requirements in axonal mitophagy, the selective removal of damaged mitochondria6. In the present protocol, different steps are presented to induce local mitochondrial damage through selective treatment of axons using the mitochondrial complex III inhibitor Antimycin A6,7.

<table><tbody><tr><td>6-well Glass bottom plate</td><td>Cellvis</td><td>P06.1.5H-N</td><td>Silicone device</td></tr><tr><td>Antimycin A</td><td>Sigma</td><td>A8674</td><td></td></tr><tr><td>B27</td><td>Gibco</td><td>17504044</td><td></td></tr><tr><td>EVOS M5000 widefield microscope</td><td>Thermofischer Scientific</td><td>EVOS M5000</td><td>fully integrated digital widefield microscope</td></tr><tr><td>Hibernate E</td><td>BrainBits</td><td>HE500</td><td></td></tr><tr><td>Inverted spinning disk confocal</td><td>Nikon</td><td>TI2-E + CSU-W1</td><td>With incubator chamber</td></tr><tr><td>Laminin</td><td>Invitrogen</td><td>L2020</td><td></td></tr><tr><td>Microfluidic devices</td><td>XONA microfluidics</td><td>RD450</td><td></td></tr><tr><td>Neurobasal medium</td><td>Gibco</td><td>21103049</td><td></td></tr><tr><td>Poly-D-Lysine</td><td>Sigma</td><td>P2636</td><td></td></tr><tr><td>TMRE</td><td>Sigma</td><td>87917</td><td></td></tr></tbody></table>

microfluidics-assisted selective depolarization of axonal mitochondria

Mitochondria are the primary suppliers of ATP (adenosine triphosphate) in neurons. Mitochondrial dysfunction is a common phenotype in many neurodegenerative diseases. Given some axons' elaborate architecture and extreme length, it is not surprising that mitochondria in axons can experience different environments compared to their cell body counterparts. Interestingly, dysfunction of axonal mitochondria often precedes effects on the cell body. To model axonal mitochondrial dysfunction in vitro, microfluidic devices allow treatment of axonal mitochondria without affecting the somal mitochondria. The fluidic pressure gradient in these chambers prevents diffusion of molecules against the gradient, thus allowing for analysis of mitochondrial properties in response to local pharmacological challenges within axons. The current protocol describes the seeding of dissociated hippocampal neurons in microfluidic devices, staining with a membrane-potential sensitive dye, treatment with a mitochondrial toxin, and the subsequent microscopic analysis. This versatile method to study axonal biology can be applied to many pharmacological perturbations and imaging readouts, and is suitable for several neuronal subtypes.

Primary hippocampal neurons were grown in microfluidic devices for 7-8 days before mitochondria were stained with the membrane-sensitive dye (TMRE) for 25 min in both the channels. As shown in Figure 2A, this yielded homogenous staining of mitochondria on both sides of the microgrooves, yet it was insufficient to equilibrate the staining into the middle of the microgrooves. Upon addition of Antimycin A to the axonal side, somal mitochondria retained the TMRE signal (Figu...

Watch this Scientific Journal Video about Microfluidics-Assisted Selective Depolarization of Axonal Mitochondria at JoVE.com

Microfluidics-Assisted Selective Depolarization of Axonal Mitochondria

The present protocol describes the seeding and staining of neuronal mitochondria in microfluidic chambers. The fluidic pressure gradient in these chambers allows for the selective treatment of mitochondria in axons to analyze their properties in response to pharmacological challenges without affecting the cell body compartment.

Mitochondria are the primary suppliers of ATP (adenosine triphosphate) in neurons. Mitochondrial dysfunction is a common phenotype in many ...

microfluidics-assisted-selective-depolarization-of-axonal-mitochondria

Research

JoVE Journal

Neuroscience

1.7K Views.  Technical University of Munich. The present protocol describes the seeding and staining of neuronal mitochondria in microfluidic chambers. The fluidic pressure gradient in these chambers allows for the selective treatment of mitochondria in axons to analyze their properties in response to pharmacological challenges without affecting the cell body compartment.

Video: Microfluidics-Assisted Selective Depolarization of Axonal Mitochondria

Assessing Mitochondrial Function in Sciatic Nerve by High-Resolution Respirometry

Mitochondrial dysfunction in peripheral nerves accompanies several diseases associated with peripheral neuropathy, which can be triggered by multiple causes, including autoimmune diseases, diabetes, infections, inherited disorders, and tumors. Assessing mitochondrial function in mouse peripheral nerves can be challenging due to the small sample size, a limited number of mitochondria present in the tissue, and the presence of a myelin sheath. The technique described in this work minimizes these challenges by using a unique permeabilization protocol adapted from one used for muscle fibers, to assess sciatic nerve mitochondrial function instead of isolating the mitochondria from the tissue. By measuring fluorimetric reactive species production with Amplex Red/Peroxidase and comparing different mitochondrial substrates and inhibitors in saponin-permeabilized nerves, it was possible to detect mitochondrial respiratory states, reactive oxygen species (ROS), and the activity of mitochondrial complexes simultaneously. Therefore, the method presented here offers advantages compared to the assessment of mitochondrial function by other techniques.

High-resolution respirometry coupled to fluorescence sensors determines mitochondrial oxygen consumption and reactive oxygen species (ROS) generation. The present protocol describes a technique to assess mitochondrial respiratory rates and ROS production in the permeabilized sciatic nerve.

Mitochondrial dysfunction in peripheral nerves accompanies several diseases associated with peripheral neuropathy, which can be triggered by multiple ...

Biochemistry

Isolation of Mitochondrial Contact Sites

An Improved Method to Isolate Mitochondrial Contact Sites

Mitochondria are present in virtually all eukaryotic cells and perform essential functions that go far beyond energy production, for instance, the synthesis of iron-sulfur clusters, lipids, or proteins, Ca2+ buffering, and the induction of apoptosis. Likewise, mitochondrial dysfunction results in severe human diseases such as cancer, diabetes, and neurodegeneration. In order to perform these functions, mitochondria have to communicate with the rest of the cell across their envelope, which consists of two membranes. Therefore, these two membranes have to interact constantly. Proteinaceous contact sites between the mitochondrial inner and outer membranes are essential in this respect. So far, several contact sites have been identified. In the method described here, Saccharomyces cerevisiae mitochondria are used to isolate contact sites and, thus, identify candidates that qualify for contact site proteins. We used this method to identify the mitochondrial contact site and cristae organizing system (MICOS) complex, one of the major contact site-forming complexes in the mitochondrial inner membrane, which is conserved from yeast to humans. Recently, we further improved this method to identify a novel contact site consisting of Cqd1 and the Por1-Om14 complex.

Author Spotlight: Unveiling Mitochondrial Contact Sites and Architectural Insights 

Mitochondrial contact sites are protein complexes that interact with mitochondrial inner and outer membrane proteins. These sites are essential for the communication between the mitochondrial membranes and, thus, between the cytosol and the mitochondrial matrix. Here, we describe a method to identify candidates qualifying for this specific class of proteins.

Mitochondria are present in virtually all eukaryotic cells and perform essential functions that go far beyond energy production, for instance, the ...

Interfacing Microfluidics with Microelectrode Arrays for Studying Neuronal Communication and Axonal Signal Propagation

Microelectrode arrays (MEAs) are widely used to study neuronal function in vitro. These devices allow concurrent non-invasive recording/stimulation of electrophysiological activity for long periods. However, the property of sensing signals from all sources around every microelectrode can become unfavorable when trying to understand communication and signal propagation in neuronal circuits. In a neuronal network, several neurons can be simultaneously activated and can generate overlapping action potentials, making it difficult to discriminate and track signal propagation. Considering this limitation, we have established an in vitro setup focused on assessing electrophysiological communication, which is able to isolate and amplify axonal signals with high spatial and temporal resolution. By interfacing microfluidic devices and MEAs, we are able to compartmentalize neuronal cultures with a well-controlled alignment of the axons and microelectrodes. This setup allows recordings of spike propagation with a high signal-to-noise ratio over the course of several weeks. Combined with specialized data analysis algorithms, it provides detailed quantification of several communication related properties such as propagation velocity, conduction failure, firing rate, anterograde spikes, and coding mechanisms.
This protocol demonstrates how to create a compartmentalized neuronal culture setup over substrate-integrated MEAs, how to culture neurons in this setup, and how to successfully record, analyze and interpret the results from such experiments. Here, we show how the established setup simplifies the understanding of neuronal communication and axonal signal propagation. These platforms pave the way for new in vitro models with engineered and controllable neuronal network topographies. They can be used in the context of homogeneous neuronal cultures, or with co-culture configurations where, for example, communication between sensory neurons and other cell types is monitored and assessed. This setup provides very interesting conditions to study, for example, neurodevelopment, neuronal circuits, information coding, neurodegeneration and neuroregeneration approaches.

This protocol aims to demonstrate how to combine in vitro microelectrode arrays with microfluidic devices for studying action potential transmission in neuronal cultures. Data analysis, namely the detection and characterization of propagating action potentials, is performed using a new advanced, yet user-friendly and freely available, computational tool.

Microelectrode arrays (MEAs) are widely used to study neuronal function in vitro. These devices allow concurrent non-invasive recording/stimulation of ...

Bioengineering

Analyzing Mitochondrial Transport and Morphology in Human Induced Pluripotent Stem Cell-Derived Neurons in Hereditary Spastic Paraplegia

Neurons have intense demands for high energy in order to support their functions. Impaired mitochondrial transport along axons has been observed in human neurons, which may contribute to neurodegeneration in various disease states. Although it is challenging to examine mitochondrial dynamics in live human nerves, such paradigms are critical for studying the role of mitochondria in neurodegeneration. Described here is a protocol for analyzing mitochondrial transport and mitochondrial morphology in forebrain neuron axons derived from human induced pluripotent stem cells (iPSCs). The iPSCs are differentiated into telencephalic glutamatergic neurons using well-established methods. Mitochondria of the neurons are stained with MitoTracker CMXRos, and mitochondrial movement within the axons are captured using a live-cell imaging microscope equipped with an incubator for cell culture. Time-lapse images are analyzed using software with &#34;MultiKymograph&#34;, &#34;Bioformat importer&#34;, and &#34;Macros&#34; plugins. Kymographs of mitochondrial transport are generated, and average mitochondrial velocity in the anterograde and retrograde directions is read from the kymograph. Regarding mitochondrial morphology analysis, mitochondrial length, area, and aspect ratio are obtained using the ImageJ. In summary, this protocol allows characterization of mitochondrial trafficking along axons and analysis of their morphology to facilitate studies of neurodegenerative diseases.

Impaired mitochondrial transport and morphology are involved in various neurodegenerative diseases. The presented protocol uses induced pluripotent stem cell-derived forebrain neurons to assess mitochondrial transport and morphology in hereditary spastic paraplegia. This protocol allows characterization of mitochondrial trafficking along axons and analysis of their morphology, which will facilitate the study of neurodegenerative disease.

Neurons have intense demands for high energy in order to support their functions. Impaired mitochondrial transport along axons has been observed in human ...

Axonal Transport of Organelles in Motor Neuron Cultures using Microfluidic Chambers System

Motor neurons (MNs) are highly polarized cells with very long axons. Axonal transport is a crucial mechanism for MN health, contributing to neuronal growth, development, and survival. We describe a detailed method for the use of microfluidic chambers (MFCs) for tracking axonal transport of fluorescently labeled organelles in MN axons. This method is rapid, relatively inexpensive, and allows for the monitoring of intracellular cues in space and time. We describe a step by step protocol for: 1) Fabrication of polydimethylsiloxane (PDMS) MFCs; 2) Plating of ventral spinal cord explants and MN dissociated culture in MFCs; 3) Labeling of mitochondria and acidic compartments followed by live confocal imagining; 4) Manual and semiautomated axonal transport analysis. Lastly, we demonstrate a difference in the transport of mitochondria and acidic compartments of HB9::GFP ventral spinal cord explant axons as a proof of the system validity. Altogether, this protocol provides an efficient tool for studying the axonal transport of various axonal components, as well as a simplified manual for MFC usage to help discover spatial experimental possibilities.

Axonal transport is a crucial mechanism for motor neuron health. In this protocol we provide a detailed method for tracking the axonal transport of acidic compartments and mitochondria in motor neuron axons using microfluidic chambers.

Motor neurons (MNs) are highly polarized cells with very long axons. Axonal transport is a crucial mechanism for MN health, contributing to neuronal ...

Immunodetection of Outer Membrane Proteins by Flow Cytometry of Isolated Mitochondria

Methods to detect and monitor mitochondrial outer membrane protein components in animal tissues are vital to study mitochondrial physiology and pathophysiology. This protocol describes a technique where mitochondria isolated from rodent tissue are immunolabeled and analyzed by flow cytometry. Mitochondria are isolated from rodent spinal cords and subjected to a rapid enrichment step so as to remove myelin, a major contaminant of mitochondrial fractions prepared from nervous tissue. Isolated mitochondria are then labeled with an antibody of choice and a fluorescently conjugated secondary antibody. Analysis by flow cytometry verifies the relative purity of mitochondrial preparations by staining with a mitochondrial specific dye, followed by detection and quantification of immunolabeled protein. This technique is rapid, quantifiable and high-throughput, allowing for the analysis of hundreds of thousands of mitochondria per sample. It is applicable to assess novel proteins at the mitochondrial surface under normal physiological conditions as well as the proteins that may become mislocalized to this organelle during pathology. Importantly, this method can be coupled to fluorescent indicator dyes to report on certain activities of mitochondrial subpopulations and is feasible for mitochondria from the central nervous system (brain and spinal cord) as well as liver.

Described here is a method to detect and quantify mitochondrial outer membrane proteins by immunolabeling of mitochondria isolated from rodent tissue and analysis by flow cytometry. This method can be extended to assess functional aspects of mitochondrial subpopulations.

Methods to detect and monitor mitochondrial outer membrane protein components in animal tissues are vital to study mitochondrial physiology and ...

Biology

Isolation and Functional Analysis of Mitochondria from Cultured Cells and Mouse Tissue

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of cell heath due to their role in both cell metabolism and energy production as well as control of apoptosis. Therefore, direct evaluation of isolated mitochondria and mitochondrial perturbation offers the ability to determine if organelle-specific (dys)function is occurring. The methods described in this protocol include isolation of intact, functional mitochondria from HEK cultured cells and mouse liver and spinal cord, but can be easily adapted for use with other cultured cells or animal tissues. Mitochondrial function assessed by TMRE and the use of common mitochondrial uncouplers and inhibitors in conjunction with a fluorescent plate reader allow this protocol not only to be versatile and accessible to most research laboratories, but also offers high throughput.

Comparison of mitochondrial membrane potential between samples yields valuable information about cellular status. Detailed steps for isolating mitochondria and assessing response to inhibitors and uncouplers using fluorescence are described. The method and utility of this protocol are illustrated by use of a cell culture and animal model of cellular stress.

Comparison between two or more distinct groups, such as healthy vs. disease, is necessary to determine cellular status. Mitochondria are at the nexus of ...

MATLAB Optimization for Analyzing Mitochondrial Networks in Neuroscience

Optimized Automated Analysis of Live Neuronal Mitochondria Homeostasis Modulation by Isoform-Specific Retinoic Acid Receptors

The complex mitochondrial network makes it very challenging to segment, follow, and analyze live cells. MATLAB tools allow the analysis of mitochondria in timelapse files, considerably simplifying and speeding up the process of image processing. Nonetheless, existing tools produce a large output volume, requiring individual manual attention, and basic experimental setups have an output of thousands of files, each requiring extensive and time-consuming handling. 
To address these issues, a routine optimization was developed, in both MATLAB code and live-script forms, allowing for swift file analysis and significantly reducing document reading and data processing. With a speed of 100 files/min, the optimization allows an overall rapid analysis. The optimization achieves the results output by averaging frame-specific data for individual mitochondria throughout time frames, analyzing data in a defined manner, consistent with those output from existing tools. Live confocal imaging was performed using the dye tetramethylrhodamine methyl ester, and the routine optimization was validated by treating neuronal cells with retinoic acid receptor (RAR) agonists, whose effects on neuronal mitochondria are established in the literature. The results were consistent with the literature and allowed further characterization of mitochondrial network behavior in response to isoform-specific RAR modulation. 
This new methodology allowed rapid and validated characterization of whole-neuron mitochondria network, but it also allows for differentiation between axon and cell body mitochondria, an essential feature to apply in the neuroscience field. Moreover, this protocol can be applied to experiments using fast-acting treatments, allowing the imaging of the same cells before and after treatments, transcending the field of neuroscience.

Author Spotlight: An Optimized Automated Method for Investigating Retinoic Acid Receptors in Neuronal Mitochondria

The mitochondrial network is extremely complex, making it very challenging to analyze. A novel MATLAB tool analyzes live confocal imaged mitochondria in timelapse images but results in a large output volume requiring individual manual attention. To address this issue, a routine optimization was developed, allowing for speedy file analysis.

The complex mitochondrial network makes it very challenging to segment, follow, and analyze live cells. MATLAB tools allow the analysis of mitochondria in ...

Assessment of Open Probability of the Mitochondrial Permeability Transition Pore in the Setting of Coenzyme Q Excess

The mitochondrial permeability transition pore (mPTP) is a voltage-gated, nonselective, inner mitochondrial membrane (IMM) mega-channel important in health and disease. The mPTP mediates leakage of protons across the IMM during low-conductance opening and is specifically inhibited by cyclosporine A (CsA). Coenzyme Q (CoQ) is a regulator of the mPTP, and tissue-specific differences have been found in CoQ content and open probability of the mPTP in forebrain and heart mitochondria in a newborn mouse model of fragile X syndrome (FXS, Fmr1 knockout). We developed a technique to determine the voltage threshold for mPTP opening in this mutant strain, exploiting the role of the mPTP as a proton leak channel.
To do so, oxygen consumption and membrane potential (&#916;&#936;) were simultaneously measured in isolated mitochondria using polarography and a tetraphenylphosphonium (TPP+) ion-selective electrode during leak respiration. The threshold for mPTP opening was determined by the onset of CsA-mediated inhibition of proton leak at specific membrane potentials. Using this approach, differences in voltage gating of the mPTP were precisely defined in the context of CoQ excess. This novel technique will permit future investigation for enhancing the understanding of physiological and pathological regulation of low-conductance opening of the mPTP.

This method exploits the contribution of the mitochondrial permeability transition pore to low-conductance proton leak to determine the voltage threshold for pore opening in neonatal fragile X syndrome mice with increased cardiomyocyte mitochondrial coenzyme Q content compared to wildtype control.

The mitochondrial permeability transition pore (mPTP) is a voltage-gated, nonselective, inner mitochondrial membrane (IMM) mega-channel important in ...

Improved Protein Quantification Using Two-Step Mitochondrial Isolation

Two-Step Tag-Free Isolation of Mitochondria for Improved Protein Discovery and Quantification

Most physiological and disease processes, from central metabolism to immune response to neurodegeneration, involve mitochondria. The mitochondrial proteome is composed of more than 1,000 proteins, and the abundance of each can vary dynamically in response to external stimuli or during disease progression. Here, we describe a protocol for isolating high-quality mitochondria from primary cells and tissues. The two-step procedure comprises (1) mechanical homogenization and differential centrifugation to isolate crude mitochondria, and (2) tag-free immune capture of mitochondria to isolate pure organelles and eliminate contaminants. Mitochondrial proteins from each purification stage are analyzed by quantitative mass spectrometry, and enrichment yields are calculated, allowing the discovery of novel mitochondrial proteins by subtractive proteomics. Our protocol provides a sensitive and comprehensive approach to studying mitochondrial content in cell lines, primary cells, and tissues.

Author Spotlight: Two-Step Tag-Free Isolation of Mitochondria for Improved Protein Discovery and Quantification  

We present a two-step protocol for high-quality mitochondria isolation that is compatible with protein discovery and quantification at a proteome scale. Our protocol does not require genetic engineering and is thus suitable for studying mitochondria from any primary cells and tissues.