Name: spectral reflectometric microscopy on myelinated axons in situ
Uploaded: 2018-07-02T15:00:00.000+00:00
Duration: 9 min 13 s
Description: Watch this Scientific Journal Video about Spectral Reflectometric Microscopy on Myelinated Axons In Situ at JoVE.com

This work was supported by the Institute of Basic Science (IBS-R015-D1) and by Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Education (2017R1A6A1A03015642).

All surgical procedures were approved by the Institutional Animal Care and Use Committee (IACUC) of Sungkyunkwan University.
1. Sample Preparation
Note: Autoclave all surgical instruments before animal handling. Conduct all surgical performance in a room dedicated to surgical procedures. Sterile surgical gowns and gloves should be worn by all personnel in the surgical room at all times.
<ol>
	<li>Tissue Fixation
	<ol>
		<li>Prepare two 10 mL syringes each filled with phosphate-buffered saline (PBS) and 4% paraformaldehyde (PFA) in PBS.</li>
		<li>Anesthetize a 7-12-week-old mouse, (C57BL/6J or any mouse lines with either gender) by administering a mixture of zoletil and xylazine (1:1, 20 mg/kg each) or an alternative suitable anesthesia regime (e.g., mixture of 80 mg/kg ketamine and 10 mg/kg xylazine) with an intraperitoneal injection.</li>
		<li>Once the mouse has reached a surgical plane of anesthesia (use the toe pinch-response method), expose the heart by cutting off the skin and the rib cage with scissors.</li>
		<li>Snip the right atrium with small scissors to drain the blood and perfuse the PBS and PFA solution sequentially through the left ventricle with a needle (perfusion rate = 4 mL/min, volume = 10 mL for each solution). 
		Note: The mouse becomes stiff if the procedure is successfully completed. The use of eye ointment or sterilization is not necessary as the procedure is terminal. The fixation procedure may be skipped. However, this step is recommended to minimize structural deformations including mechanical tissue damage and ischemic axonal swelling. For further details on tissue fixation, refer to the article by Gage, G. J. et al.10</li>
	</ol>
	</li>
	<li>Preparation of the Brain Slice 
	<ol>
		<li>Decapitate the mouse with surgical scissors through the ventral thorax and abdomen.</li>
		<li>Remove the scalp and periosteum using a suitably small scissors until fully exposing the cranium.</li>
		<li>Break the frontal bone and cut the cranium along the sagittal suture from the occipital bone using small scissors with care not to damage the brain.</li>
		<li>Gently remove the cranium and the dura mater sequentially using fine forceps.</li>
		<li>Extract the brain using a soft plastic spoon, taking care not to damage the tissue.</li>
		<li>Rinse and soak the extracted brain in a 4% PFA solution for 30 min at 4 &#176;C.</li>
		<li>Slice the fixed brain into 100 &#8211; 150 &#181;m sections using a vibratome. 
		Note: For further details on the tissue preparation, refer to the article by Segev et al.11. For subsequent procedures, the tissue should be sliced into sections thinner than the thickness of the spacer.</li>
	</ol>
	</li>
	<li>Mounting of the Brain Slice
	<ol>
		<li>Prepare 1 glass slide and 2 square cover glasses (22 &#215; 22 &#215; 0.17 mm) for each tissue slice.</li>
		<li>Cut one of the square cover glasses into half (two rectangular pieces) by using a glass cutter.</li>
		<li>Attach the two of the halved cover glasses using a super glue on the slide glass. 
		Note: The space between the two halved glasses should be slightly larger than the size of the tissue slice.</li>
		<li>Harvest the brain slice using a brush or a pipette and lay the tissue on the slide glass between the spacer. Take care not to fold the tissue.</li>
		<li>Dispense 100 &#956;L of PBS on the tissue surface.</li>
		<li>Place the other square cover glass on top of the tissue. Avoid inclusion of air bubbles during this stage.</li>
		<li>Seal around the cover glass using a nail polish (or alternatively suitable adhesives) to prevent evaporation of PBS and contamination by dust during the subsequent imaging session. 
		Note: The experimenter may pause at this stage.</li>
	</ol>
	</li>
</ol>
2. Calibration
<ol>
	<li>Switch on the microscope at least 1 h before the imaging to allow thermal stabilization of the laser source (see Table of Materials). Then, turn on the software for spectral scanning (see Table of Materials) and click the Acquire button on the top of GUI (Figure 3a).</li>
	<li>Turn on the software shutter for the white-light laser (WLL) and photomultiplier tube (PMT) (Figure 3a).</li>
	<li>Select the xy&#923; mode on the drop-down list in Acquisition Mode, then, the laser input mode is automatically changed from Constant percent to Constant Power. Uncheck the check-box of Automatic SP Movement. And then, set the spectral window of the input laser to 470&#8211;670 nm and the spectral step size to 4 nm on &#923;-Excitation Lambda Scan Settings (Figure 3b).</li>
	<li>SET the spectral range of PMT to 450-690 by double-clicking or moving the adjustment bar for spectral range (Figure 3c). 
	Note: This spectral range should include the complete bandwidth of the input source.</li>
	<li>Select a water-immersion objective lens, suitably with a high numerical aperture (NA &#62; 0.7) and give any value to PMT and laser power to activate AOBS configuration and Live Scan button. Switch the optical path by checking reflection in the AOBS Configuration (Figure 3d).</li>
	<li>Mount a reference mirror (see Table of Materials) on the microscope stage. The mirror surface should be faced against the objective lens. If it is not easy to put the mirror on the microscope stage, bond a mirror onto the flat plate (e.g. slide glass).</li>
	<li>Control the microscope stage to align the focal plane to the mirror surface.</li>
	<li>Adjust the PMT gain and the laser power considering the dynamic range of the detector and then change the laser input mode from Constant Percent to Constant Power. 
	Note: As is typical, a PMT gain of 500 (V) and a relative laser power of 0.1% are used at 570 nm.</li>
	<li>Confirm in a pseudo-color mode to check that there is no saturation throughout the wavelength range. If saturation is observed, lower the laser power (Figure 3a).</li>
	<li>Run the Lambda Scan acquisition.</li>
	<li>Remove the mirror from the stage and repeat the same acquisition without a sample in order to obtain the dark reference (i.e., dark offset).</li>
	<li>Save the data in a Multistacked TIF format.</li>
</ol>
3. SpeRe Image Acquisition
<ol>
	<li>Place the mounted tissue on the microscope stage. To roughly align the tissue to the focal plane of the objective lens, use the wide-field fluorescence mode through an eye-piece.</li>
	<li>With the Live Scan on, control the microscope stage to align the focal plane to the region of interest in the tissue. To avoid the background noise from the cover slip, select the target region of at least 15 &#956;m depth from the glass-tissue interface.</li>
	<li>Acquire the spectral image stack for the target region using the same procedure as described in steps 2.1&#8211;2.10.</li>
	<li>Save the data for the tissue and the dark offset in Multistacked TIF format. 
	Note: The experimenter may pause at this stage.</li>
</ol>
4. Image Processing and Analysis
<ol>
	<li>Open the spectral data (multistacked TIF) for the reference mirror and the brain tissue in ImageJ.</li>
	<li>Select the ROIs (regions of interest) for the opened image stacks&#8212;the central area for the reference mirror and the segment of an axon fiber for the brain tissue.</li>
	<li>Acquire the raw spectra for the selected ROIs by running Image &#8594; Stacks &#8594; Plot Z-axis profile on the ImageJ menu.</li>
	<li>Open the dark offset data, one taken for the reference mirror and the other taken for the brain tissue.</li>
	<li>Acquire the spectra for the dark offsets by running Image &#8594; Stacks &#8594; Plot Z-axis profile on the ImageJ menu.</li>
	<li>Save all the acquired spectra by using the Copy and Paste options. 
	Note: For SpeRe imaging, use of the central imaging field to minimize off-axis optical aberrations is recommended. The axon fibers may be structurally heterogeneous along their length. Hence, selecting the ROI on a small axon segment, typically &#60; 5 &#181;m, to minimize partial-volume artifact is recommended.</li>
</ol>
5. Baseline Correction and SpeRe Signal Analysis
<ol>
	<li>Subtract the offset spectra from the spectra of the reference mirror and the brain tissues.</li>
	<li>Normalize each spectrum by dividing the maximum intensity of the spectrum.</li>
	<li>Divide the normalized spectrum of the axon by the normalized spectrum of the reference mirror&#160;and subtract DC offset from the normalized spectrum.</li>
	<li>Measure the wavenumber frequency&#160;by fitting the acquired spectrum to a sinusoidal curve (see Table of Materials), and then convert the acquired wavenumber to periodicity by taking reciprocal.</li>
	<li>Convert the wavenumber periodicity to the axon diameter using the equation in Figure 4c.</li>
</ol>

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	<li>Fernandez-Moran, H., Finean, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electron+microscope+and+low-angle+x-ray+diffraction+studies+of+the+nerve+myelin+sheath.">Electron microscope and low-angle x-ray diffraction studies of the nerve myelin sheath.</a> Journal of Cell Biology. 3, (1957).</li><li>Blaurock, A. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+spaces+between+membrane+bilayers+within+PNS+myelin+as+characterized+by+X-ray+diffraction.">The spaces between membrane bilayers within PNS myelin as characterized by X-ray diffraction.</a> Brain Research. 210, 383-387 (1981).</li><li>Shim, S. -. 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The authors declare competing financial interests: J. Kwon and M. Choi are inventors of the patent-pending technology described in this Article.

SpeRe is a new label-free imaging modality based on spectral interferometry, which for the first time, offers the nanoscale information in live myelinated axons. In the current acquisition protocol, the spatial resolution for the axon diameter is of the order of 10 nm. Moreover, SpeRe utilizes orders-of-magnitude lower light dose compared to other super-resolution microscopies; thus, it is free from phototoxicity and photobleaching. SpeRe would provide a new avenue for studying nanoscale dynamics of the myelinated axons....

In the mammalian nervous system, myelin provides rapid nerve conduction and axonal integrity by enwrapping the axon fibers with multilayered membranous sheaths. Its multilayered structure is composed of alternating nanoscale thin-films composed of plasma membranes (~5 nm), cytosol (~3 nm), and extracellular spaces (~7 nm)1,2. Optical microscopy, including the recent super-resolution microscopy, are not suitable for observing the nanoscale myelin dynamics due to their insufficient resolution due to optical diffraction3,4,5. Although electron microscopy can provide fine details of the myelin nanostructure, it is not compatible with the living biological systems due to highly invasive sample preparations involving chemical fixation and ultrasectioning6,7. Until recently, there has been no technique applicable to observe nanoscale dynamics of myelinated axons in situ.
Schain et al. previously reported that myelinated axons exhibit colorful light reflectance8. By adopting the spectroscopic analysis on the reflected light, we have devised a new imaging modality for nanoscale imaging of myelinated axons, named spectral reflectometry (SpeRe)9. SpeRe is based on the thin-film interference occurring in the multi-layered structure of the myelin sheath (Figure 1). By optic simulation on various axons, we have revealed that the reflectance spectrum is a periodic function of wavenumber and its periodicity (<img alt="Equation 1" src="/files/ftp_upload/57965/57965eq1.jpg" />) is inversely proportional to the axon diameter (d). This simple relationship (<img alt="Equation 2" src="/files/ftp_upload/57965/57965eq2.jpg" />) offers facile quantification of axon diameter from the SpeRe data. Utilizing this, we revealed the prevalent axon bulging under mild traumatic brain injury in our prior report.
The SpeRe system is based on confocal microscopy and consists of a specialized laser source and filters (Figure 2). The input source is a white-light laser, providing broadband spectral output visible to infrared regions. For the spectral scan, the system is equipped with two acousto-optic devices: an acousto-optic tunable filter (AOTF) for delivering a selected wavelength from the input broadband source and an acousto-optic beam splitter (AOBS) for guiding the selected reflected wavelength to the detector. The software for hyperspectral confocal microscopy (see Table of Materials) provides a customizable spectral scan option to sequentially acquire the reflectance images at various input wavelengths. In addition, chromatic aberration can critically interfere in the spectral measurement; therefore, use of an apochromat objective lens is recommended.
Of note, white-light lasers produce an uneven spectral output and the optical components also affect the spectral profile. Therefore, the acquired spectra need to be calibrated for the subsequent quantitative analysis. A protected silver mirror is typically used as a reference, which provides a nearly constant reflectance (&#62; 97%) over the full visible region. The acquired spectra are then divided by the reference spectra from the mirror.
The spectral step size for the spectral scan determines the acquisition speed; thus, it needs to be optimized. As a larger axon has a higher spectral period, it requires finer spectral sampling. For example, an axon with a diameter of 10 &#181;m, one of the largest physiologic axons, has a spectral period of ~8 nm. By applying Nyquist sampling criteria, we employed the spectral sampling interval of 4 nm to cover all the physiologic axons in the mouse nervous tissues. This approach typically takes over several seconds for a full spectral scan and thus is not suitable for in vivo applications, where physiologic motion (e.g. respiration and heartbeat) interferes stable spectral acquisition. We previously solved this issue by instrumenting a customized upright microscope, designed to acquire the full spectrum for each point using an array spectrometer (acquisition speed &#8776; 30 ms per pixel).
In this report, we describe a detailed protocol on the SpeRe imaging on a fixed brain slice, which can be performed in a commercial hyperspectral microscope (see Table of Materials). Thus, the protocol can be completed by experimenters without expertise in optical instrumentation. We also cover the potential issues and troubleshooting for acquisition and analysis of SpeRe data.

<table><tbody><tr><td>Glass cutter</td><td>-</td><td>-</td><td>Can be purchased in a local convenience store or online stores.</td></tr><tr><td>Nail polish</td><td>-</td><td>-</td><td>Can be purchased in a local convenience store or online stores.</td></tr><tr><td>Apochromat objective 40&amp;times;, NA 1.1</td><td>Leica Microsystems</td><td>15506357</td><td>Water-immersion type</td></tr><tr><td>Fluoromyelin Green</td><td>Thermo Fisher</td><td>F34651</td><td>Alternatively, Fluoromyelin Red (F34652) can be used.</td></tr><tr><td>Leica SP8 TCS microscope</td><td>Leica Microsystems</td><td>SP8</td><td>Refer to the &quot;Configuration of microscope&quot; in Introduction Section for details.</td></tr><tr><td>Imaging software</td><td>Leica Microsystems</td><td>LAS-X</td><td>-</td></tr><tr><td>Matlab</td><td>MathWorks</td><td>-</td><td>-</td></tr><tr><td>Mirror</td><td>Thorlabs</td><td>PF10-03-P01</td><td>Coated with protected silver.</td></tr><tr><td>Phosphate-buffered saline (PBS)</td><td>Life technologies</td><td>14190-136</td><td>-</td></tr><tr><td>Paraformaldehyde</td><td>Biosolution</td><td>BP031a</td><td>4% v/v in PBS</td></tr><tr><td>Cover slip</td><td>Thermo Fisher</td><td>3306</td><td>Thickness: #1 (0.13 to 0.17 mm)</td></tr><tr><td>Slide glass</td><td>Muto Pure Chemicals</td><td>5116-20F</td><td>Thickness: ~1 mm</td></tr><tr><td>Super glue</td><td>Henkel</td><td>Loctite 406</td><td>Use a dispensing equipment to avoid skin or eye contact.</td></tr><tr><td>Syringe pump</td><td>Brainetree Scientific</td><td>BS-8000 DUAL</td><td>-</td></tr><tr><td>Vibratome</td><td>Leica Biosystems</td><td>VT1200S</td><td>-</td></tr><tr><td>White-light laser</td><td>NKT photonics</td><td>EXB-6</td><td>EXB-6 was discontinued and replaced by EXU-6.</td></tr></tbody></table>

spectral reflectometric microscopy on myelinated axons in situ

In a mammalian nervous system, myelin provides an electrical insulation by enwrapping the axon fibers in a multilayered spiral. Inspired by its highly-organized subcellular architecture, we recently developed a new imaging modality, named spectral reflectometry (SpeRe), which enables unprecedented label-free nanoscale imaging of the live myelinated axons in situ. The underlying principle is to obtain nanostructural information by analyzing the reflectance spectrum of the multilayered subcellular structure. In this article, we describe a detailed step-by-step protocol for performing a basic SpeRe imaging of the nervous tissues using a commercial confocal microscopic system, equipped with a white-light laser and a tunable filter. We cover the procedures of sample preparation, acquisition of spectral data, and image processing for obtaining nanostructural information.

According to the protocol, a fixed brain slice was prepared with exogenous staining, a myelin-targeting fluorophore (see Table of Materials). SpeRe imaging was performed on the brain slice using a commercial hyperspectral confocal microscope in conjunction with confocal fluorescence imaging (Figure 4a). For SpeRe, input optical intensity was set to 5 &#181;W/&#181;m2 with a pixel dwell time of ~1 &#181;s. This light dose is over an...

Watch this Scientific Journal Video about Spectral Reflectometric Microscopy on Myelinated Axons In Situ at JoVE.com

Spectral Reflectometric Microscopy on Myelinated Axons In Situ

Here, we present a step-by-step protocol for imaging myelinated axons in a fixed brain slice using a label-free nanoscale imaging technique based on spectral reflectometry.

Spectral Reflectometric Microscopy on Myelinated Axons In Situ

In a mammalian nervous system, myelin provides an electrical insulation by enwrapping the axon fibers in a multilayered spiral. Inspired by its ...

spectral-reflectometric-microscopy-on-myelinated-axons-in-situ

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7.3K Views.  Sungkyunkwan University. Here, we present a step-by-step protocol for imaging myelinated axons in a fixed brain slice using a label-free nanoscale imaging technique based on spectral reflectometry.

Video: Spectral Reflectometric Microscopy on Myelinated Axons In Situ

Acute Dissociation of Lamprey Reticulospinal Axons to Enable Recording from the Release Face Membrane of Individual Functional Presynaptic Terminals

Synaptic transmission is an extremely rapid process. Action potential driven influx of Ca2+&#160;into the presynaptic terminal, through voltage-gated calcium channels (VGCCs) located in the release face membrane, is the trigger for vesicle fusion and neurotransmitter release. Crucial to the rapidity of synaptic transmission is the spatial and temporal synchrony between the arrival of the action potential, VGCCs and the neurotransmitter release machinery. The ability to directly record Ca2+&#160;currents from the release face membrane of individual presynaptic terminals is imperative for a precise understanding of the relationship between presynaptic Ca2+&#160;and neurotransmitter release. Access to the presynaptic release face membrane for electrophysiological recording is not available in most preparations and presynaptic Ca2+&#160;entry has been characterized using imaging techniques and macroscopic current measurements &#8211; techniques that do not have sufficient temporal resolution to visualize Ca2+ entry. The characterization of VGCCs directly at single presynaptic terminals has not been possible in central synapses and has thus far been successfully achieved only in the calyx-type synapse of the chick ciliary ganglion and in rat calyces. We have successfully addressed this problem in the giant reticulospinal synapse of the lamprey spinal cord by developing an acutely dissociated preparation of the spinal cord that yields isolated reticulospinal axons with functional presynaptic terminals devoid of postsynaptic structures. We can fluorescently label and identify individual presynaptic terminals and target them for recording. Using this preparation, we have characterized VGCCs directly at the release face of individual presynaptic terminals using immunohistochemistry and electrophysiology approaches. Ca2+ currents have been recorded directly at the release face membrane of individual presynaptic terminals, the first such recording to be carried out at central synapses.

Recording Ca2+ currents at the presynaptic release face membrane is key to a precise understanding of Ca2+ entry and neurotransmitter release. We present an acute dissociation of the lamprey spinal cord that yields functional isolated reticulospinal axons, permitting recording directly from the release face membrane of individual presynaptic terminals.

Synaptic transmission is an extremely rapid process. Action potential driven influx of Ca2+&#160;into the presynaptic terminal, through voltage-gated ...

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal degeneration. Lack of growth cone formation, regeneration, and loss of additional myelinated axonal projections within the spinal cord greatly limits neurological recovery following injury. To assess how central myelinated axons of the spinal cord respond to injury, we developed an ex vivo living spinal cord model utilizing transgenic mice that express yellow fluorescent protein in axons and a focal and highly reproducible laser-induced spinal cord injury to document the fate of axons and myelin (lipophilic fluorescent dye Nile Red) over time using two-photon excitation time-lapse microscopy. Dynamic processes such as acute axonal injury, axonal retraction, and myelin degeneration are best studied in real-time. However, the non-focal nature of contusion-based injuries and movement artifacts encountered during in vivo spinal cord imaging make differentiating primary and secondary axonal injury responses using high resolution microscopy challenging. The ex vivo spinal cord model described here mimics several aspects of clinically relevant contusion/compression-induced axonal pathologies including axonal swelling, spheroid formation, axonal transection, and peri-axonal swelling providing a useful model to study these dynamic processes in real-time. Major advantages of this model are excellent spatiotemporal resolution that allows differentiation between the primary insult that directly injures axons and secondary injury mechanisms; controlled infusion of reagents directly to the perfusate bathing the cord; precise alterations of the environmental milieu (e.g., calcium, sodium ions, known contributors to axonal injury, but near impossible to manipulate in vivo); and murine models also offer an advantage as they provide an opportunity to visualize and manipulate genetically identified cell populations and subcellular structures. Here, we describe how to isolate and image the living spinal cord from mice to capture dynamics of acute axonal injury.

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

We present a protocol utilizing two-photon excitation time-lapse microscopy to simultaneously visualize the dynamics of axon and myelin injuries in real time. This proposed protocol permits studies of both intrinsic and extrinsic factors which can influence central myelinated axon fate after injury and contribute to permanent clinical disability.

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal ...

Experimental Demyelination and Remyelination of Murine Spinal Cord by Focal Injection of Lysolecithin

Multiple sclerosis is an inflammatory demyelinating disease of the central nervous system characterized by plaque formation containing lost oligodendrocytes, myelin, axons, and neurons. Remyelination is an endogenous repair mechanism whereby new myelin is produced subsequent to proliferation, recruitment, and differentiation of oligodendrocyte precursor cells into myelin-forming oligodendrocytes, and is necessary to protect axons from further damage. Currently, all therapeutics for the treatment of multiple sclerosis target the aberrant immune component of the disease, which reduce inflammatory relapses but do not prevent progression to irreversible neurological decline. It is therefore imperative that remyelination-promoting strategies be developed which may delay disease progression and perhaps reverse neurological symptoms. Several animal models of demyelination exist, including experimental autoimmune encephalomyelitis and curprizone; however, there are limitations in their use for studying remyelination. A more robust approach is the focal injection of toxins into the central nervous system, including the detergent lysolecithin into the spinal cord white matter of rodents. In this protocol, we demonstrate that the surgical procedure involved in injecting lysolecithin into the ventral white matter of mice is fast, cost-effective, and requires no additional materials than those commercially available. This procedure is important not only for studying the normal events involved in the remyelination process, but also as a pre-clinical tool for screening candidate remyelination-promoting therapeutics.

Demyelinating diseases can be modeled in animals by focal application of lysolecithin into the CNS. A single injection of lysolecithin into mouse spinal cord produces a lesion that spontaneously repairs over time. The goal is to study factors involved in de- and remyelination, and to test agents for enhancing repair.

Multiple sclerosis is an inflammatory demyelinating disease of the central nervous system characterized by plaque formation containing lost ...

Visualization of Motor Axon Navigation and Quantification of Axon Arborization In Mouse Embryos Using Light Sheet Fluorescence Microscopy

Spinal motor neurons (MNs) extend their axons to communicate with their innervating targets, thereby controlling movement and complex tasks in vertebrates. Thus, it is critical to uncover the molecular mechanisms of how motor axons navigate to, arborize, and innervate their peripheral muscle targets during development and degeneration. Although transgenic Hb9::GFP mouse lines have long served to visualize motor axon trajectories during embryonic development, detailed descriptions of the full spectrum of axon terminal arborization remain incomplete due to the pattern complexity and limitations of current optical microscopy. Here, we describe an improved protocol that combines light sheet fluorescence microscopy (LSFM) and robust image analysis to qualitatively and quantitatively visualize developing motor axons. This system can be easily adopted to cross genetic mutants or MN disease models with Hb9::GFP lines, revealing novel molecular mechanisms that lead to defects in motor axon navigation and arborization.

Here, we describe a protocol for visualizing motor neuron projection and axon arborization in transgenic Hb9::GFP mouse embryos. After immunostaining for motor neurons, we used light sheet fluorescence microscopy to image embryos for subsequent quantitative analysis. This protocol is applicable to other neuron navigation processes in the central nervous system.

Spinal motor neurons (MNs) extend their axons to communicate with their innervating targets, thereby controlling movement and complex tasks in ...

In vivo Imaging of Biological Tissues with Combined Two-Photon Fluorescence and Stimulated Raman Scattering Microscopy

Stimulated Raman scattering (SRS) microscopy enables label-free imaging of the biological tissues in its natural microenvironment based on intrinsic molecular vibration, thus providing a perfect tool for in vivo study of biological processes at subcellular resolution. By integrating two-photon excited fluorescence (TPEF) imaging into the SRS microscope, the dual-modal in vivo imaging of tissues can acquire critical biochemical and biophysical information from multiple perspectives which helps understand the dynamic processes involved in cellular metabolism, immune response and tissue remodeling, etc. In this video protocol, the setup of a TPEF-SRS microscope system as well as the in vivo imaging method of the animal spinal cord is introduced. The spinal cord, as part of the central nervous system, plays a critical role in the communication between the brain and peripheral nervous system. Myelin sheath, abundant in phospholipids, surrounds and insulates the axon to permit saltatory conduction of action potentials. In vivo imaging of myelin sheaths in the spinal cord is important to study the progression of neurodegenerative diseases and spinal cord injury. The protocol also describes animal preparation and in vivo TPEF-SRS imaging methods to acquire high-resolution biological images.

In vivo Imaging of Biological Tissues with Combined Two-Photon Fluorescence and Stimulated Raman Scattering Microscopy

Stimulated Raman scattering (SRS) microscopy allows label-free imaging of biomolecules based on their intrinsic vibration of specific chemical bonds. In this protocol, the instrumental setup of an integrated SRS and two-photon fluorescence microscope is described to visualize cellular structures in the spinal cord of live mice.

Stimulated Raman scattering (SRS) microscopy enables label-free imaging of the biological tissues in its natural microenvironment based on intrinsic ...

Bioengineering

Preparation and Immunostaining of Myelinating Organotypic Cerebellar Slice Cultures

In the nervous system, myelin is a complex membrane structure generated by myelinating glial cells, which ensheathes axons and facilitates fast electrical conduction. Myelin alteration has been shown to occur in various neurological diseases, where it is associated with functional deficits. Here, we provide a detailed description of an ex vivo model consisting of mouse organotypic cerebellar slices, which can be maintained in culture for several weeks and further be labeled to visualize myelin.

Here we present a method to prepare organotypic slice cultures from mouse cerebellum and myelin sheath staining by immunohistochemistry suitable for investigating mechanisms of myelination and remyelination in the central nervous system.

In the nervous system, myelin is a complex membrane structure generated by myelinating glial cells, which ensheathes axons and facilitates fast electrical ...

Imaging and Analysis of Neurofilament Transport in Excised Mouse Tibial Nerve

Neurofilament protein polymers move along axons in the slow component of axonal transport at average speeds of ~0.35-3.5 mm/day. Until recently the study of this movement in situ was only possible using radioisotopic pulse-labeling, which permits analysis of axonal transport in whole nerves with a temporal resolution of days and a spatial resolution of millimeters. To study neurofilament transport in situ with higher temporal and spatial resolution, we developed a hThy1-paGFP-NFM transgenic mouse that expresses neurofilament protein M tagged with photoactivatable GFP in neurons. Here we describe fluorescence photoactivation pulse-escape and pulse-spread methods to analyze neurofilament transport in single myelinated axons of tibial nerves from these mice ex vivo. Isolated nerve segments are maintained on the microscope stage by perfusion with oxygenated saline and imaged by spinning disk confocal fluorescence microscopy. Violet light is used to activate the fluorescence in a short axonal window. The fluorescence in the activated and flanking regions is analyzed over time, permitting the study of neurofilament transport with temporal and spatial resolution on the order of minutes&#160;and microns, respectively. Mathematical modeling can be used to extract kinetic parameters of neurofilament transport including the velocity, directional bias and pausing behavior from the resulting data. The pulse-escape and pulse-spread methods can also be adapted to visualize neurofilament transport in other nerves. With the development of additional transgenic mice, these methods could also be used to image and analyze the axonal transport of other cytoskeletal and cytosolic proteins in axons.

We describe fluorescence photoactivation methods to analyze the axonal transport of neurofilaments in single myelinated axons of peripheral nerves from transgenic mice that express a photoactivatable neurofilament protein.

Neurofilament protein polymers move along axons in the slow component of axonal transport at average speeds of ~0.35-3.5 mm/day. Until recently the study ...

Measuring Properties of the Membrane Periodic Skeleton of the Axon Initial Segment using 3D-Structured Illumination Microscopy (3D-SIM)

The axon initial segment (AIS) is the site at which action potentials initiate and constitutes a transport filter and diffusion barrier that contribute to the maintenance of neuronal polarity by sorting somato-dendritic cargo. A membrane periodic skeleton (MPS) comprising periodic actin rings provides a scaffold for anchoring various AIS proteins, including structural proteins and different ion channels. Although recent proteomic approaches have identified a considerable number of novel AIS components, details of the structure of the MPS and the roles of its individual components are lacking. The distance between individual actin rings in the MPS (~190 nm) necessitates the employment of super-resolution microscopy techniques to resolve the structural details of the MPS. This protocol describes a method for using cultured rat hippocampal neurons to examine the precise localization of an AIS protein in the MPS relative to sub-membranous actin rings using 3D-structured illumination microscopy (3D-SIM). In addition, an analytical approach to quantitively assess the periodicity of individual components and their position relative to actin rings is also described.

Measuring Properties of the Membrane Periodic Skeleton of the Axon Initial Segment using 3D-Structured Illumination Microscopy 3D-SIM

The present protocol describes a method to visualize and measure actin rings and other components of the membrane periodic skeleton of the axon initial segment using cultured rat hippocampal neurons and 3D-structured illumination microscopy (3D-SIM).

The axon initial segment (AIS) is the site at which action potentials initiate and constitutes a transport filter and diffusion barrier that contribute to ...

Coherent Anti-Stokes Raman Spectroscopy (CARS) Application for Imaging Myelination in Brain Slices

Coherent anti-Stokes Raman spectroscopy (CARS) is a technique classically employed by chemists and physicists to produce a coherent signal of signature vibrations of molecules. However, these vibrational signatures are also characteristic of molecules within anatomical tissue such as the brain, making it increasingly useful and applicable for Neuroscience applications. For example, CARS can measure lipids by specifically exciting chemical bonds within these molecules, allowing for quantification of different aspects of tissue, such as myelin involved in neurotransmission. In addition, compared to other techniques typically used to quantify myelin, CARS can also be set up to be compatible with immunofluorescent techniques, allowing for co-labeling with other markers such as sodium channels or other components of synaptic transmission. Myelination changes are an inherently important mechanism in demyelinating diseases such as multiple sclerosis or other neurological conditions such as Fragile X Syndrome or autism spectrum disorders is an emerging area of research. In conclusion, CARS can be utilized in innovative ways to answer pressing questions in Neuroscience and provide evidence for underlying mechanisms related to many different neurological conditions.

Coherent Anti-Stokes Raman Spectroscopy CARS Application for Imaging Myelination in Brain Slices

Visualizing myelination is an important goal for many researchers studying the nervous system. CARS is a technique that is compatible with immunofluorescence that can natively image lipids within tissue such as the brain illuminating specialized structures such as myelin.

Coherent anti-Stokes Raman spectroscopy (CARS) is a technique classically employed by chemists and physicists to produce a coherent signal of signature ...

Spectral Reflectometric Microscopy on Myelinated Axons In Situ

Summary

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Chapters in this video

Acute Dissociation of Lamprey Reticulospinal Axons to Enable Recording from the Release Face Membrane of Individual Functional Presynaptic Terminals

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

Experimental Demyelination and Remyelination of Murine Spinal Cord by Focal Injection of Lysolecithin

Visualization of Motor Axon Navigation and Quantification of Axon Arborization In Mouse Embryos Using Light Sheet Fluorescence Microscopy

In vivo Imaging of Biological Tissues with Combined Two-Photon Fluorescence and Stimulated Raman Scattering Microscopy

Preparation and Immunostaining of Myelinating Organotypic Cerebellar Slice Cultures

Imaging and Analysis of Neurofilament Transport in Excised Mouse Tibial Nerve

Measuring Properties of the Membrane Periodic Skeleton of the Axon Initial Segment using 3D-Structured Illumination Microscopy (3D-SIM)

Coherent Anti-Stokes Raman Spectroscopy (CARS) Application for Imaging Myelination in Brain Slices