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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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

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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

Mechanical Manipulation of Neurons to Control Axonal Development

Cell manipulations and extension of neuronal axons can be accomplished with calibrated glass micro-fibers capable of measuring and applying forces in the 10-1000 &mu;dyne range1,2. Force measurements are obtained through observation of the Hookean bending of the glass needles, which are calibrated by a direct and empirical method3. Equipment requirements and procedures for fabricating, calibrating, treating, and using the needles on cells are fully described. The force regimes previously used and different cell types to which these techniques have been applied demonstrate the flexibility of the methodology and are given as examples for future investigation4-6. The technical advantages are the continuous 'visualization' of the forces produced by the manipulations and the ability to directly intervene in a variety of cellular events. These include direct stimulation and regulation of axonal growth and retraction7; as well as detachment and mechanical measurements on any type of cultured cell8.

Application and direct measurements of forces on neurons in the 2-1000 microdyne range are achieved with high precision using calibrated glass needles. This methodology can be used to control and measure several aspects of axonal development, including axonal initiation, axonal tension, velocity of axonal elongation, and force vectors.

Cell manipulations and extension of neuronal axons can be accomplished with calibrated glass micro-fibers capable of measuring and applying forces in the ...

Chromatin Immunoprecipitation from Dorsal Root Ganglia Tissue following Axonal Injury

Axons in the central nervous system (CNS) do not regenerate while those in the peripheral nervous system (PNS) do regenerate 
to a limited extent after injury (Teng et al., 2006). It is recognized that transcriptional programs essential for neurite and axonal outgrowth are 
reactivated upon injury in the PNS (Makwana et al., 2005). However the tools available to analyze neuronal gene regulation in vivo are limited and 
often challenging.
The dorsal root ganglia (DRG) offer an excellent injury model system because both the CNS and PNS are innervated by a 
bifurcated axon originating from the same soma. The ganglia represent a discrete collection of cell bodies where all transcriptional events occur, 
and thus provide a clearly defined region of transcriptional activity that can be easily and reproducibly removed from the animal. Injury of nerve 
fibers in the PNS (e.g. sciatic nerve), where axonal regeneration does occur, should reveal a set of transcriptional programs that are distinct from 
those responding to a similar injury in the CNS, where regeneration does not take place (e.g. spinal cord). Sites for transcription factor binding, 
histone and DNA modification resulting from injury to either PNS or CNS can be characterized using chromatin immunoprecipitation (ChIP). 
Here, we describe a ChIP protocol using fixed mouse DRG tissue following axonal injury. This powerful combination provides a means for characterizing the pro-regeneration chromatin environment necessary for promoting axonal regeneration.

We present a method for chromatin immunoprecipitation from dorsal root ganglia tissue following axonal injury. The approach can be used to identify specific transcription factor binding sites and epigenetic modification of histone and DNA important for the regeneration of injured axons in both the peripheral and central nervous system.

Axons in the central nervous system (CNS) do not regenerate while those in the peripheral nervous system (PNS) do regenerate 
to a limited extent after ...

Subtype-selective Electroporation of Cortical Interneurons

The study of central nervous system (CNS) maturation relies on genetic targeting of neuronal populations. However, the task of restricting the expression of genes of interest to specific neuronal subtypes has proven remarkably challenging due to the relative scarcity of specific promoter elements. GABAergic interneurons constitute a neuronal population with extensive genetic and morphological diversity. Indeed, more than 11 different subtypes of GABAergic interneurons have been characterized in the mouse cortex1. Here we present an adapted protocol for selective targeting of GABAergic populations. We achieved subtype selective targeting of GABAergic interneurons by using the enhancer element of the homeobox transcription factors Dlx5 and Dlx6, homologues of the Drosophila distal-less (Dll) gene2,3, to drive the expression of specific genes through in utero electroporation.

This procedure shows how to target interneurons in the developing mouse forebrain by means of in utero electroporation. This technique was particularly efficient to achieve selective gene expression in interneuron subtypes destined to the superficial layers of the cortex.

The study of central nervous system (CNS) maturation relies on genetic targeting of neuronal populations. However, the task of restricting the expression ...

Real-time Imaging of Axonal Transport of Quantum Dot-labeled BDNF in Primary Neurons

BDNF plays an important role in several facets of neuronal survival, differentiation, and function. Structural and functional deficits in axons are increasingly viewed as an early feature of neurodegenerative diseases, including Alzheimer&#8217;s disease (AD) and Huntington&#8217;s disease (HD). As yet unclear is the mechanism(s) by which axonal injury is induced. We reported the development of a novel technique to produce biologically active, monobiotinylated BDNF (mBtBDNF) that can be used to trace axonal transport of BDNF. Quantum dot-labeled BDNF (QD-BDNF) was produced by conjugating quantum dot 655 to mBtBDNF. A microfluidic device was used to isolate axons from neuron cell bodies. Addition of QD-BDNF to the axonal compartment allowed live imaging of BDNF transport in axons. We demonstrated that QD-BDNF moved essentially exclusively retrogradely, with very few pauses, at a moving velocity of around 1.06 &#956;m/sec. This system can be used to investigate mechanisms of disrupted axonal function in AD or HD, as well as other degenerative disorders.

Axonal transport of BDNF, a neurotrophic factor, is critical for the survival and function of several neuronal populations. Some degenerative disorders are marked by disruption of axonal structure and function. We demonstrated the techniques used to examine live trafficking of QD-BDNF in microfluidic chambers using primary neurons.

BDNF plays an important role in several facets of neuronal survival, differentiation, and function. Structural and functional deficits in axons are ...

Characterizing the Composition of Molecular Motors on Moving Axonal Cargo Using "Cargo Mapping" Analysis

Understanding the mechanisms by which molecular motors coordinate their activities to transport vesicular cargoes within neurons requires the quantitative analysis of motor/cargo associations at the single vesicle level. The goal of this protocol is to use quantitative fluorescence microscopy to correlate (&#8220;map&#8221;) the position and directionality of movement of live cargo to the composition and relative amounts of motors associated with the same cargo. &#8220;Cargo mapping&#8221; consists of live imaging of fluorescently labeled cargoes moving in axons cultured on microfluidic devices, followed by chemical fixation during recording of live movement, and subsequent immunofluorescence (IF) staining of the exact same axonal regions with antibodies against motors. Colocalization between cargoes and their associated motors is assessed by assigning sub-pixel position coordinates to motor and cargo channels, by fitting Gaussian functions to the diffraction-limited point spread functions representing individual fluorescent point sources. Fixed cargo and motor images are subsequently superimposed to plots of cargo movement, to &#8220;map&#8221; them to their tracked trajectories. The strength of this protocol is the combination of live and IF data to record both the transport of vesicular cargoes in live cells and to determine the motors associated to these exact same vesicles. This technique overcomes previous challenges that use biochemical methods to determine the average motor composition of purified heterogeneous bulk vesicle populations, as these methods do not reveal compositions on single moving cargoes. Furthermore, this protocol can be adapted for the analysis of other transport and/or trafficking pathways in other cell types to correlate the movement of individual intracellular structures with their protein composition. Limitations of this protocol are the relatively low throughput due to low transfection efficiencies of cultured primary neurons and a limited field of view available for high-resolution imaging. Future applications could include methods to increase the number of neurons expressing fluorescently labeled cargoes.

Characterizing the Composition of Molecular Motors on Moving Axonal Cargo Using &quot;Cargo Mapping&quot; Analysis

Intracellular transport of cargoes, such as vesicles or organelles, is carried out by molecular motor proteins that track on polarized microtubules. This protocol describes the correlation of the directionality of transport of individual cargo particles moving inside neurons, to the relative amount and type of associated motor proteins.

Understanding the mechanisms by which molecular motors coordinate their activities to transport vesicular cargoes within neurons requires the quantitative ...

Three-dimensional Imaging and Analysis of Mitochondria within Human Intraepidermal Nerve Fibers

The goal of this protocol is to study mitochondria within intraepidermal nerve fibers. Therefore, 3D imaging and analysis techniques were developed to isolate nerve-specific mitochondria and evaluate disease-induced alterations of mitochondria in the distal tip of sensory nerves. The protocol combines fluorescence immunohistochemistry, confocal microscopy and 3D image analysis techniques to visualize and quantify nerve-specific mitochondria. Detailed parameters are defined throughout the procedures in order to provide a concrete example of how to use these techniques to isolate nerve-specific mitochondria. Antibodies were used to label nerve and mitochondrial signals within tissue sections of skin punch biopsies, which was followed by indirect immunofluorescence to visualize nerves and mitochondria with a green and red fluorescent signal respectively. Z-series images were acquired with confocal microscopy and 3D analysis software was used to process and analyze the signals. It is not necessary to follow the exact parameters described within, but it is important to be consistent with the ones chosen throughout the staining, acquisition and analysis steps. The strength of this protocol is that it is applicable to a wide variety of circumstances where one fluorescent signal is used to isolate other signals that would otherwise be impossible to study alone.

This protocol uses three-dimensional (3D) imaging and analysis techniques to visualize and quantify nerve-specific mitochondria. The techniques are applicable to other situations where one fluorescent signal is used to isolate a subset of data from another fluorescent signal.

The goal of this protocol is to study mitochondria within intraepidermal nerve fibers. Therefore, 3D imaging and analysis techniques were developed to ...

Analysis of Brain Mitochondria Using Serial Block-Face Scanning Electron Microscopy

Human brain is a high energy consuming organ that mainly relies on glucose as a fuel source. Glucose is catabolized by brain mitochondria via glycolysis, tri-carboxylic acid (TCA) cycle and oxidative phosphorylation (OXPHOS) pathways to produce cellular energy in the form of adenosine triphosphate (ATP). Impairment of mitochondrial ATP production causes mitochondrial disorders, which present clinically with prominent neurological and myopathic symptoms. Mitochondrial defects are also present in neurodevelopmental disorders (e.g. autism spectrum disorder) and neurodegenerative disorders (e.g. amyotrophic lateral sclerosis, Alzheimer&#39;s and Parkinson&#39;s diseases). Thus, there is an increased interest in the field for performing 3D analysis of mitochondrial morphology, structure and distribution under both healthy and disease states. The brain mitochondrial morphology is extremely diverse, with some mitochondria especially those in the synaptic region being in the range of &#60;200 nm diameter, which is below the resolution limit of traditional light microscopy. Expressing a mitochondrially-targeted green fluorescent protein (GFP) in the brain significantly enhances the organellar detection by confocal microscopy. However, it does not overcome the constraints on the sensitivity of detection of relatively small sized mitochondria without oversaturating the images of large sized mitochondria. While serial transmission electron microscopy has been successfully used to characterize mitochondria at the neuronal synapse, this technique is extremely time-consuming especially when comparing multiple samples. The serial block-face scanning electron microscopy (SBFSEM) technique involves an automated process of sectioning, imaging blocks of tissue and data acquisition. Here, we provide a protocol to perform SBFSEM of a defined region from rodent brain to rapidly reconstruct and visualize mitochondrial morphology. This technique could also be used to provide accurate information on mitochondrial number, volume, size and distribution in a defined brain region. Since the obtained image resolution is high (typically under 10 nm) any gross mitochondrial morphological defects may also be detected.

Mitochondrial visualization and analysis from mammalian brain tissue is a challenging task. Here, we describe how three dimensional (3D) reconstruction analysis from the serial block-face scanning electron microscopy (SBFSEM) can be used to gain insights on the morphological and volumetric analysis of this critical energy generating organelle.

Human brain is a high energy consuming organ that mainly relies on glucose as a fuel source. Glucose is catabolized by brain mitochondria via glycolysis, ...

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons extend growth cones to navigate along specific pathways by responding to a large number of axon guidance cues that emanate from the surrounding extracellular environment. Growth cone target recognition also plays a critical role in neuromuscular specificity. This work presents a standard immunohistochemistry protocol to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. This protocol includes a few key steps, including a genotyping procedure, to sort the desired mutant embryos; an immunostaining procedure, to tag embryos with fasciclin II (FasII) antibody; and a dissection procedure, to generate filleted preparations from fixed embryos. Motor axon projections and muscle patterns in the periphery are much better visualized in flat preparations of filleted embryos than in whole-mount embryos. Therefore, the filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition, and it can also be applied to both loss-of-function and gain-of-function genetic screens.

Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila

This work details a standard immunohistochemistry method to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. The filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition during neural development.

The establishment of functional neuromuscular circuits relies on precise connections between developing motor axons and target muscles. Motor neurons ...

In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing

Electrophysiology enables the objective assessment of peripheral nerve function in vivo. Traditional nerve conduction measures such as amplitude and latency detect chronic axon loss and demyelination, respectively. Axonal excitability techniques &#34;by threshold tracking&#34; expand upon these measures by providing information regarding the activity of ion channels, pumps and exchangers that relate to acute function and may precede degenerative events. As such, the use of axonal excitability in animal models of neurological disorders may provide a useful in vivo measure to assess novel therapeutic interventions. Here we describe an experimental setup for multiple measures of motor axonal excitability techniques in the rat ulnar nerve.
The animals are anesthetized with isoflurane and carefully monitored to ensure constant and adequate depth of anesthesia. Body temperature, respiration rate, heart rate and saturation of oxygen in the blood are continuously monitored. Axonal excitability studies are performed using percutaneous stimulation of the ulnar nerve and recording from the hypothenar muscles of the forelimb paw. With correct electrode placement, a clear compound muscle action potential that increases in amplitude with increasing stimulus intensity is recorded. An automated program is then utilized to deliver a series of electrical pulses which generate 5 specific excitability measures in the following sequence: stimulus response behavior, strength duration time constant, threshold electrotonus, current-threshold relationship and the recovery cycle.
Data presented here indicate that these measures are repeatable and show similarity between left and right ulnar nerves when assessed on the same day. A limitation of these techniques in this setting is the effect of dose and time under anesthesia. Careful monitoring and recording of these variables should be undertaken for consideration at the time of analysis.

In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing

Axonal excitability techniques provide a powerful tool to examine pathophysiology and biophysical changes that precede irreversible degenerative events. This manuscript demonstrates the use of these techniques on the ulnar nerve of anesthetized rats.

Electrophysiology enables the objective assessment of peripheral nerve function in vivo. Traditional nerve conduction measures such as amplitude and ...

FM Dye Cycling at the Synapse: Comparing High Potassium Depolarization, Electrical and Channelrhodopsin Stimulation

FM dyes are used to study the synaptic vesicle (SV) cycle. These amphipathic probes have a hydrophilic head and hydrophobic tail, making them water-soluble with the ability to reversibly enter and exit membrane lipid bilayers. These styryl dyes are relatively non-fluorescent in aqueous medium, but insertion into the outer leaflet of the plasma membrane causes a &#62;40X increase in fluorescence. In neuronal synapses, FM dyes are internalized during SV endocytosis, trafficked both within and between SV pools, and released with SV exocytosis, providing a powerful tool to visualize presynaptic stages of neurotransmission. A primary genetic model of glutamatergic synapse development and function is the Drosophila neuromuscular junction (NMJ), where FM dye imaging has been used extensively to quantify SV dynamics in a wide range of mutant conditions. The NMJ synaptic terminal is easily accessible, with a beautiful array of large synaptic boutons ideal for imaging applications. Here, we compare and contrast the three ways to stimulate the Drosophila NMJ to drive activity-dependent FM1-43 dye uptake/release: 1) bath application of high [K+] to depolarize neuromuscular tissues, 2) suction electrode motor nerve stimulation to depolarize the presynaptic nerve terminal, and 3) targeted transgenic expression of channelrhodopsin variants for light-stimulated, spatial control of depolarization. Each of these methods has benefits and disadvantages for the study of genetic mutation effects on the SV cycle at the Drosophila NMJ. We will discuss these advantages and disadvantages to assist the selection of the stimulation approach, together with the methodologies specific to each strategy. In addition to fluorescent imaging, FM dyes can be photoconverted to electron-dense signals visualized using transmission electron microscopy (TEM) to study SV cycle mechanisms at an ultrastructural level. We provide the comparisons of confocal and electron microscopy imaging from the different methods of Drosophila NMJ stimulation, to help guide the selection of future experimental paradigms.

Synaptic vesicle (SV) cycling is the core mechanism of intercellular communication at neuronal synapses. FM dye uptake and release are the primary means of quantitatively assaying SV endo- and exocytosis. Here, we compare all the stimulation methods to drive FM1-43 cycling at the Drosophila neuromuscular junction (NMJ) model synapse.