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>

<ol>
	<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. -. H., 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=Super-resolution+fluorescence+imaging+of+organelles+in+live+cells+with+photoswitchable+membrane+probes.">Super-resolution fluorescence imaging of organelles in live cells with photoswitchable membrane probes.</a> Proceedings of the National Academy of Sciences. , 13978-13983 (2012).</li><li>Urban, N. T., Willig, K. I., Hell, S. W., N&#228;gerl, U. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=STED+nanoscopy+of+actin+dynamics+in+synapses+deep+inside+living+brain+slices.">STED nanoscopy of actin dynamics in synapses deep inside living brain slices.</a> Biophysics Journal. 101, 1277-1284 (2011).</li><li>Manley, S., 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=High-density+mapping+of+single-molecule+trajectories+with+photoactivated+localization+microscopy.">High-density mapping of single-molecule trajectories with photoactivated localization microscopy.</a> Nature Methods. 5, 155-157 (2008).</li><li>Peters, A., Sethares, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Is+there+remyelination+during+aging+of+the+primate+central+nervous+system?.">Is there remyelination during aging of the primate central nervous system?.</a> Journal of Comparative Neurology. 460, 238-254 (2003).</li><li>De Campos Vidal, B., Silveira Mello, M. L., Caseiro-Filho, A. C., Godo, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anisotropic+properties+of+the+myelin+sheath.">Anisotropic properties of the myelin sheath.</a> Acta Histochemica. 66, 32-39 (1980).</li><li>Schain, A. J., Hill, R. A., Grutzendler, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Label-free+in+vivo+imaging+of+myelinated+axons+in+health+and+disease+with+spectral+confocal+reflectance+microscopy.">Label-free in vivo imaging of myelinated axons in health and disease with spectral confocal reflectance microscopy.</a> Nature Medicine. 20, 443-449 (2014).</li><li>Kwon, J., 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=Label-free+nanoscale+optical+metrology+on+myelinated+axons+in+vivo.">Label-free nanoscale optical metrology on myelinated axons in vivo.</a> Nature Communication. 8, 1832 (2017).</li><li>Gage, G. J., Kipke, D. R., Shain, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whole+Animal+Perfusion+Fixation+for+Rodents.">Whole Animal Perfusion Fixation for Rodents.</a> JoVE. (65), e3564 (2012).</li><li>Segev, A., Garcia-Oscos, F., Kourrich, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whole-cell+Patch-clamp+Recordings+in+Brain+Slices.">Whole-cell Patch-clamp Recordings in Brain Slices.</a> JoVE. (112), e54024 (2016).</li><li>Waldchen, S., Lehmann, J., Klein, T., Van De Linde, S., Sauer, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Light-induced+cell+damage+in+live-cell+super-resolution+microscopy.">Light-induced cell damage in live-cell super-resolution microscopy.</a> Scientific Reports. 5, 15348 (2015).</li><li>Chang, J. 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=Iterative+expansion+microscopy.">Iterative expansion microscopy.</a> Nature Methods. 14, 593-599 (2017).</li><li>Polishchuk, R. S., 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=Correlative+light-electron+microscopy+reveals+the+tubular-saccular+ultrastructure+of+carriers+operating+between+Golgi+apparatus+and+plasma+membrane.">Correlative light-electron microscopy reveals the tubular-saccular ultrastructure of carriers operating between Golgi apparatus and plasma membrane.</a> Journal of Cell Biology. 148, 45-58 (2000).</li></ol>

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&#215;, 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 &#34;Configuration of microscope&#34; 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

Research

JoVE Journal

Neuroscience

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

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different neurological diseases. Defects in this essential process can have detrimental effects on neuronal functioning and development. We have developed a dissection protocol that is designed to expose the Drosophila larval segmental nerves to view axonal transport in real time. We have adapted this protocol for live imaging from the one published by Hurd and Saxton (1996) used for immunolocalizatin of larval segmental nerves. Careful dissection and proper buffer conditions are critical for maximizing the lifespan of the dissected larvae. When properly done, dissected larvae have shown robust vesicle transport for 2-3 hours under physiological conditions. We use the UAS-GAL4 method 1 to express GFP-tagged APP or synaptotagmin vesicles within a single axon or many axons in larval segmental nerves by using different neuronal GAL4 drivers. Other fluorescently tagged markers, for example mitochrondria (MitoTracker) or lysosomes (LysoTracker), can be also applied to the larvae before viewing. GFP-vesicle movement and particle movement can be viewed simultaneously using separate wavelengths.

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

This protocol discusses the live dissection of Drosophila larvae for the purpose of imaging the movement of GFP tagged axonal vesicles on microtubule tracks.

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different ...

Live Imaging of Dorsal Root Axons after Rhizotomy

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. Regeneration of dorsal root (DR) axons into spinal cord is prevented at the dorsal root entry zone (DREZ), the interface between the CNS and PNS. Our understanding of the molecular and cellular events that prevent regeneration at DREZ is incomplete, in part because complex changes associated with nerve injury have been deduced from postmortem analyses. Dynamic cellular processes, such as axon regeneration, are best studied with techniques that capture real-time events with multiple observations of each living animal. Our ability to monitor neurons serially in vivo has increased dramatically owing to revolutionary innovations in optics and mouse transgenics. Several lines of thy1-GFP transgenic mice, in which subsets of neurons are genetically labeled in distinct fluorescent colors, permit individual neurons to be imaged in vivo1. These mice have been used extensively for in vivo imaging of muscle2-4 and brain5-7, and have provided novel insights into physiological mechanisms that static analyses could not have resolved. Imaging studies of neurons in living spinal cord have only recently begun. Lichtman and his colleagues first demonstrated their feasibility by tracking injured dorsal column (DC) axons with wide-field microscopy8,9. Multi-photon in vivo imaging of deeply positioned DC axons, microglia and blood vessels has also been accomplished10. Over the last few years, we have pioneered in applying in vivo imaging to monitor regeneration of DR axons using wide-field microscopy and H line of thy1-YFP mice. These studies have led us to a novel hypothesis about why DR axons are prevented from regenerating within the spinal cord11.
In H line of thy1-YFP mice, distinct YFP+ axons are superficially positioned, which allows several axons to be monitored simultaneously. We have learned that DR axons arriving at DREZ are better imaged in lumbar than in cervical spinal cord. In the present report we describe several strategies that we have found useful to assure successful long-term and repeated imaging of regenerating DR axons. These include methods that eliminate repeated intubation and respiratory interruption, minimize surgery-associated stress and scar formation, and acquire stable images at high resolution without phototoxicity.

An in vivo imaging protocol to monitor primary sensory axons following dorsal root crush is described. The procedures utilize wide-field fluorescence microscopy and thy1-YFP transgenic mice, and permit repeated imaging of axon regeneration over 4 cm in the PNS and axon interactions with the interface of the CNS.

The primary sensory axons injured by spinal root injuries fail to regenerate into the spinal cord, leading to chronic pain and permanent sensory loss. ...

Spectral Confocal Imaging of Fluorescently tagged Nicotinic Receptors in Knock-in Mice with Chronic Nicotine Administration

Ligand-gated ion channels in the central nervous system (CNS) are implicated in numerous conditions with serious medical and social consequences. For instance, addiction to nicotine via tobacco smoking is a leading cause of premature death worldwide (World Health Organization) and is likely caused by an alteration of ion channel distribution in the brain1. Chronic nicotine exposure in both rodents and humans results in increased numbers of nicotinic acetylcholine receptors (nAChRs) in brain tissue1-3. Similarly, alterations in the glutamatergic GluN1 or GluA1 channels have been implicated in triggering sensitization to other addictive drugs such as cocaine, amphetamines and opiates4-6.

Consequently, the ability to map and quantify distribution and expression patterns of specific ion channels is critically important to understanding the mechanisms of addiction. The study of brain region-specific effects of individual drugs was advanced by the advent of techniques such as radioactive ligands. However, the low spatial resolution of radioactive ligand binding prevents the ability to quantify ligand-gated ion channels in specific subtypes of neurons. 

Genetically encoded fluorescent reporters, such as green fluorescent protein (GFP) and its many color variants, have revolutionized the field of biology7.By genetically tagging a fluorescent reporter to an endogenous protein one can visualize proteins in vivo7-10. One advantage of fluorescently tagging proteins with a probe is the elimination of antibody use, which have issues of nonspecificity and accessibility to the target protein. We have used this strategy to fluorescently label nAChRs, which enabled the study of receptor assembly using F&ouml;rster Resonance Energy Transfer (FRET) in transfected cultured cells11.More recently, we have used the knock-in approach to engineer mice with yellow fluorescent protein tagged &alpha;4 nAChR subunits (&alpha;4YFP), enabling precise quantification of the receptor ex vivo at submicrometer resolution in CNS neurons via spectral confocal microscopy12. The targeted fluorescent knock-in mutation is incorporated in the endogenous locus and under control of its native promoter, producing normal levels of expression and regulation of the receptor when compared to untagged receptors in wildtype mice. This knock-in approach can be extended to fluorescently tag other ion channels and offers a powerful approach of visualizing and quantifying receptors in the CNS.

In this paper we describe a methodology to quantify changes in nAChR expression in specific CNS neurons after exposure to chronic nicotine. Our methods include mini-osmotic pump implantation, intracardiac perfusion fixation, imaging and analysis of fluorescently tagged nicotinic receptor subunits from &alpha;4YFP knock-in mice (Fig. 1). We have optimized the fixation technique to minimize autofluorescence from fixed brain tissue.We describe in detail our imaging methodology using a spectral confocal microscope in conjunction with a linear spectral unmixing algorithm to subtract autofluoresent signal in order to accurately obtain &alpha;4YFP fluorescence signal. Finally, we show results of chronic nicotine-induced upregulation of &alpha;4YFP receptors in the medial perforant path of the hippocampus.

We have developed a novel technique of quantifying nicotinic acetylcholine receptor changes within subcellular regions of specific subtypes of CNS neurons to better understand the mechanisms of nicotine addiction by using a combination of approaches including fluorescent protein tagging of the receptor using the knock-in approach and spectral confocal imaging.

Ligand-gated ion channels in the central nervous system (CNS) are implicated in numerous conditions with serious medical and social consequences.  For ...

Functional Neuroimaging Using Ultrasonic Blood-brain Barrier Disruption and Manganese-enhanced MRI

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains technically challenging. One approach, Activation-Induced Manganese-enhanced MRI (AIM MRI), has been used successfully to map neuronal activity in rodents 1-5. In AIM MRI, Mn2+ acts a calcium analog and accumulates in depolarized neurons 6,7. Because Mn2+ shortens the T1 tissue property, regions of elevated neuronal activity will enhance in MRI. Furthermore, Mn2+ clears slowly from the activated regions; therefore, stimulation can be performed outside the magnet prior to imaging, enabling greater experimental flexibility. However, because Mn2+ does not readily cross the blood-brain barrier (BBB), the need to open the BBB has limited the use of AIM MRI, especially in mice.
One tool for opening the BBB is ultrasound. Though potentially damaging, if ultrasound is administered in combination with gas-filled microbubbles (i.e., ultrasound contrast agents), the acoustic pressure required for BBB opening is considerably lower. This combination of ultrasound and microbubbles can be used to reliably open the BBB without causing tissue damage 8-11.
Here, a method is presented for performing AIM MRI by using microbubbles and ultrasound to open the BBB. After an intravenous injection of perflutren microbubbles, an unfocused pulsed ultrasound beam is applied to the shaved mouse head for 3 minutes. For simplicity, we refer to this technique of BBB Opening with Microbubbles and UltraSound as BOMUS 12. Using BOMUS to open the BBB throughout both cerebral hemispheres, manganese is administered to the whole mouse brain. After experimental stimulation of the lightly sedated mice, AIM MRI is used to map the neuronal response.
To demonstrate this approach, herein BOMUS and AIM MRI are used to map unilateral mechanical stimulation of the vibrissae in lightly sedated mice 13. Because BOMUS can open the BBB throughout both hemispheres, the unstimulated side of the brain is used to control for nonspecific background stimulation. The resultant 3D activation map agrees well with published representations of the vibrissae regions of the barrel field cortex 14. The ultrasonic opening of the BBB is fast, noninvasive, and reversible; and thus this approach is suitable for high-throughput and/or longitudinal studies in awake mice.

A technique is described for broadly opening the blood-brain barrier in the mouse using microbubbles and ultrasound. Using this technique, manganese can be administered to the mouse brain. Because manganese is an MRI contrast agent that accumulates in depolarized neurons, this approach enables imaging of neuronal activity.

Although mice are the dominant model system for studying the genetic and molecular underpinnings of neuroscience, functional neuroimaging in mice remains ...

Analyzing Murine Schwann Cell Development Along Growing Axons

The development of peripheral nerves is an intriguing process. Neurons send out axons to innervate specific targets, which in humans are often more than 100 cm away from the soma of the neuron. Neuronal survival during development depends on target-derived growth factors but also on the support of Schwann cells (SCs). To this end SC ensheath axons from the region of the neuronal soma (or the transition from central to peripheral nervous system) to the synapse or neuromuscular junction. Schwann cells are derivatives of the neural crest and migrate as precursors along emerging axons until the entire axon is covered with SCs. This shows the importance of SC migration for the development of the peripheral nervous system and underlines the necessity to investigate this process. In order to analyze SC development, a setup is needed which next to the SCs also includes their physiological substrate for migration, the axon. Due to intrauterine development in vivo time-lapse imaging, however, is not feasible in placental vertebrates like mouse (mus musculus). To circumvent this, we adapted the superior cervical ganglion (SCG) explant technique. Upon treatment with nerve growth factor (NGF) SCG explants extend axons, followed by SC precursors migrating along the axons from the ganglion to the periphery. The beauty of this system is that the SC are derived from a pool of endogenous SC and that they migrate along their own physiological axons which are growing at the same time. This system is especially intriguing, because the SC development along axons can be analyzed by time-lapse imaging, opening further possibilities to gain insights into SC migration.

Here we describe a Schwann cell (SC) migration assay in which SCs are able to develop along extending axons.

The  development of peripheral nerves is an intriguing process. Neurons send out  axons to innervate specific targets, which in humans are often more than ...

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety of brain disorders1,2. To better understand the mechanisms by which brain circuits react to an animal's experience requires the ability to monitor the experience-dependent molecular changes in a given set of neurons, over a prolonged period of time, in the live animal. While experience and associated neural activity is known to trigger gene expression changes in neurons1,2, most of the methods to detect such changes do not allow repeated observation of the same neurons over multiple days or do not have sufficient resolution to observe individual neurons3,4. Here, we describe a method that combines in vivo two-photon microscopy with a genetically encoded fluorescent reporter to track experience-dependent gene expression changes in individual cortical neurons over the course of day-to-day experience. 
One of the well-established experience-dependent genes is Activity-regulated cytoskeletal associated protein (Arc)5,6. The transcription of Arc is rapidly and highly induced by intensified neuronal activity3, and its protein product regulates the endocytosis of glutamate receptors and long-term synaptic plasticity7. The expression of Arc has been widely used as a molecular marker to map neuronal circuits involved in specific behaviors3. In most of those studies, Arc expression was detected by in situ hybridization or immunohistochemistry in fixed brain sections. Although those methods revealed that the expression of Arc was localized to a subset of excitatory neurons after behavioral experience, how the cellular patterns of Arc expression might change with multiple episodes of repeated or distinctive experiences over days was not investigated. 
In vivo two-photon microscopy offers a powerful way to examine experience-dependent cellular changes in the living brain8,9. To enable the examination of Arc expression in live neurons by two-photon microscopy, we previously generated a knock-in mouse line in which a GFP reporter is placed under the control of the endogenous Arc promoter10. This protocol describes the surgical preparations and imaging procedures for tracking experience-dependent Arc-GFP expression patterns in neuronal ensembles in the live animal. In this method, chronic cranial windows were first implanted in Arc-GFP mice over the cortical regions of interest. Those animals were then repeatedly imaged by two-photon microscopy after desired behavioral paradigms over the course of several days. This method may be generally applicable to animals carrying other fluorescent reporters of experience-dependent molecular changes4.

In Vivo Two-photon Imaging Of Experience-dependent Molecular Changes In Cortical Neurons

Experience-dependent molecular changes in neurons are essential for the brain's ability to adapt in response to behavioral challenges. An in vivo two-photon imaging method is described here that allows the tracking of such molecular changes in individual cortical neurons through genetically encoded reporters.

The brain's ability to change in response to experience is essential for healthy brain function, and abnormalities in this process contribute to a variety ...

Laser Nanosurgery of Cerebellar Axons In Vivo

Only a few neuronal populations in the central nervous system (CNS) of adult mammals show local regrowth upon dissection of their axon. In order to understand the mechanism that promotes neuronal regeneration, an in-depth analysis of the neuronal types that can remodel after injury is needed. Several studies showed that damaged climbing fibers are capable of regrowing also in adult animals1,2. The investigation of the time-lapse dynamics of degeneration and regeneration of these axons within their complex environment can be performed by time-lapse two-photon fluorescence (TPF) imaging in vivo3,4. This technique is here combined with laser surgery, which proved to be a highly selective tool to disrupt fluorescent structures in the intact mouse cortex5-9.
This protocol describes how to perform TPF time-lapse imaging and laser nanosurgery of single axonal branches in the cerebellum in vivo. Olivocerebellar neurons are labeled by anterograde tracing with a dextran-conjugated dye and then monitored by TPF imaging through a cranial window. The terminal portion of their axons are then dissected by irradiation with a Ti:Sapphire laser at high power. The degeneration and potential regrowth of the damaged neuron are monitored by TPF in vivo imaging during the days following the injury.

Laser Nanosurgery of Cerebellar Axons In Vivo

Two-photon imaging, coupled to laser nanodissection, are useful tools to study degenerative and regenerative processes in the central nervous system with subcellular resolution. This protocol shows how to label, image, and dissect single climbing fibers in the cerebellar cortex in vivo.

Only a few neuronal populations in the central nervous system (CNS) of adult mammals show local regrowth upon dissection of their axon. In order to ...

Live Imaging of Nicotine Induced Calcium Signaling and Neurotransmitter Release Along Ventral Hippocampal Axons

Sustained enhancement of axonal signaling and increased neurotransmitter release by the activation of pre-synaptic nicotinic acetylcholine receptors (nAChRs) is an important mechanism for neuromodulation by acetylcholine (ACh). The difficulty with access to probing the signaling mechanisms within intact axons and at nerve terminals both in vitro and in vivo has limited progress in the study of the pre-synaptic components of synaptic plasticity. Here we introduce a gene-chimeric preparation of ventral hippocampal (vHipp)&#8211;accumbens (nAcc) circuit in vitro that allows direct live imaging to analyze both the pre- and post-synaptic components of transmission while selectively varying the genetic profile of the pre- vs post-synaptic neurons. We demonstrate that projections from vHipp microslices, as pre-synaptic axonal input, form multiple, reliable glutamatergic synapses with post-synaptic targets, the dispersed neurons from nAcc. The pre-synaptic localization of various subtypes of nAChRs are detected and the pre-synaptic nicotinic signaling mediated synaptic transmission are monitored by concurrent electrophysiological recording and live cell imaging. This preparation also provides an informative approach to study the pre- and post-synaptic mechanisms of glutamatergic synaptic plasticity in vitro.

We developed a gene-chimeric preparation of ventral hippocampal &#8211; accumbens circuit in vitro that allows direct live imaging to analyze presynaptic mechanisms of nicotinic acetylcholine receptors (nAChRs) mediated synaptic transmission. This preparation also provides an informative approach to study the pre- and post-synaptic mechanisms of synaptic plasticity.

Sustained enhancement of axonal signaling and increased neurotransmitter release by the activation of pre-synaptic nicotinic acetylcholine receptors ...

Rapid In Situ Hybridization using Oligonucleotide Probes on Paraformaldehyde-prefixed Brain of Rats with Serotonin Syndrome

3,4-Methylenedioxymethamphetamine (MDMA; ecstasy) toxicity may cause region-specific changes in serotonergic mRNA expression due to acute serotonin (5-hydroxytryptamine; 5-HT) syndrome. This hypothesis can be tested using in situ hybridization to detect the serotonin 5-HT2A receptor gene htr2a. In the past, such procedures, utilizing radioactive riboprobe, were difficult because of the complicated workflow that needs several days to perform and the added difficulty that the technique required the use of fresh frozen tissues maintained in an RNase-free environment. Recently, the development of short oligonucleotide probes has simplified in situ hybridization procedures and allowed the use of paraformaldehyde-prefixed brain sections, which are more widely available in laboratories. Here, we describe a detailed protocol using non-radioactive oligonucleotide probes on the prefixed brain tissues. Hybridization probes used for this study include dapB (a bacterial gene coding for dihydrodipicolinate reductase), ppiB (a housekeeping gene coding for peptidylprolyl isomerase B), and htr2a (a serotonin gene coding for 5-HT2A receptors). This method is relatively simply, cheap, reproducible and requires less than two days to complete.

Rapid In Situ Hybridization using Oligonucleotide Probes on Paraformaldehyde-prefixed Brain of Rats with Serotonin Syndrome

This protocol describes a rapid and simplified in situ hybridization method ideal forparaformaldehyde-prefixed brain, thus reducing the need for prolonged complex steps while using fresh frozen tissues. The method is validated using the identification of the serotonin 5-HT2A receptor gene htr2a in rats.

3,4-Methylenedioxymethamphetamine (MDMA; ecstasy) toxicity may cause region-specific changes in serotonergic mRNA expression due to acute serotonin ...

Determination of Photoreceptor Cell Spectral Sensitivity in an Insect Model from In Vivo Intracellular Recordings

Intracellular recording is a powerful technique used to determine how a single cell may respond to a given stimulus. In vision research, intracellular recording has historically been a common technique used to study sensitivities of individual photoreceptor cells to different light stimuli that is still being used today. However, there remains a dearth of detailed methodology in the literature for researchers wishing to replicate intracellular recording experiments in the eye. Here we present the insect as a model for examining eye physiology more generally. Insect photoreceptor cells are located near the surface of the eye and are therefore easy to reach, and many of the mechanisms involved in vision are conserved across animal phyla. We describe the basic procedure for in vivo intracellular recording of photoreceptor cells in the eye of a butterfly, with the goal of making this technique more accessible to researchers with little prior experience in electrophysiology. We introduce the basic equipment needed, how to prepare a live butterfly for recording, how to insert a glass microelectrode into a single cell, and finally the recording procedure itself. We also explain the basic analysis of raw response data for determining spectral sensitivity of individual cell types. Although our protocol focuses on determining spectral sensitivity, other stimuli (e.g., polarized light) and variations of the method are applicable to this setup.

Determination of Photoreceptor Cell Spectral Sensitivity in an Insect Model from In Vivo Intracellular Recordings

The electrophysiological technique of intracellular recording is demonstrated and used to determine spectral sensitivities of single photoreceptor cells in the compound eye of a butterfly.

Intracellular recording is a powerful technique used to determine how a single cell may respond to a given stimulus. In vision research, intracellular ...