The authors would like to thank Paula Monsma for instruction and assistance with confocal microscopy and tibial nerve dissection and Dr. Atsuko Uchida, Chloe Duger and Sana Chahande for assistance with mouse husbandry. This work was supported in part by collaborative National Science Foundation Grants IOS1656784 to A.B. and IOS1656765 to P.J., and National Institutes of Health Grants R01 NS038526, P30 NS104177 and S10 OD010383 to A.B. N.P.B. was supported by a fellowship from the Ohio State University President&#8217;s Postdoctoral Scholars Program.

<ol>
	<li>Hoffman, P. N., Lasek, R. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+slow+component+of+axonal+transport.+Identification+of+major+structural+polypeptides+of+the+axon+and+their+generality+among+mammalian+neurons.">The slow component of axonal transport. Identification of major structural polypeptides of the axon and their generality among mammalian neurons.</a> Journal of Cell Biology. 66 (2), 351-366 (1975).</li><li>Brown, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Slow+Axonal+Transport.">Slow Axonal Transport.</a> Reference Module in Biomedical Sciences. , (2014).</li><li>Wang, L., Ho, C. -. L., Sun, D., Liem, R. K. H., Brown, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rapid+movement+of+axonal+neurofilaments+interrupted+by+prolonged+pauses.">Rapid movement of axonal neurofilaments interrupted by prolonged pauses.</a> Nature Cell Biology. 2 (3), 137-141 (2000).</li><li>Uchida, A., Monsma, P. C., Fenn, J. D., Brown, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live-cell+imaging+of+neurofilament+transport+in+cultured+neurons.">Live-cell imaging of neurofilament transport in cultured neurons.</a> Methods in Cell Biology. 131, 21-90 (2016).</li><li>Trivedi, N., Jung, P., Brown, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neurofilaments+switch+between+distinct+mobile+and+stationary+states+during+their+transport+along+axons.">Neurofilaments switch between distinct mobile and stationary states during their transport along axons.</a> Journal of Neuroscience. 27 (3), 507-516 (2007).</li><li>Monsma, P. C., Li, Y., Fenn, J. D., Jung, P., Brown, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Local+regulation+of+neurofilament+transport+by+myelinating+cells.">Local regulation of neurofilament transport by myelinating cells.</a> Journal of Neuroscience. 34 (8), 2979-2988 (2014).</li><li>Walker, C. L., 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=Local+Acceleration+of+Neurofilament+Transport+at+Nodes+of+Ranvier.">Local Acceleration of Neurofilament Transport at Nodes of Ranvier.</a> Journal of Neuroscience. 39 (4), 663-677 (2019).</li><li>Li, Y., Brown, A., Jung, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Deciphering+the+axonal+transport+kinetics+of+neurofilaments+using+the+fluorescence+photoactivation+pulse-escape+method.">Deciphering the axonal transport kinetics of neurofilaments using the fluorescence photoactivation pulse-escape method.</a> Physical Biology. 11 (2), 026001 (2014).</li><li>Schindelin, 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=Fiji:+an+open-source+platform+for+biological-image+analysis.">Fiji: an open-source platform for biological-image analysis.</a> Nature Methods. 9, 676-682 (2012).</li><li>Breuer, A. C., 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=Fast+axonal+transport+in+amyotrophic+lateral+sclerosis:+an+intra-axonal+organelle+traffic+analysis.">Fast axonal transport in amyotrophic lateral sclerosis: an intra-axonal organelle traffic analysis.</a> Neurology. 37 (5), 738-748 (1987).</li><li>Bancaud, A., Huet, S., Rabut, G., Ellenberg, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescence+perturbation+techniques+to+study+mobility+and+molecular+dynamics+of+proteins+in+live+cells:+FRAP,+photoactivation,+photoconversion,+and+FLIP.">Fluorescence perturbation techniques to study mobility and molecular dynamics of proteins in live cells: FRAP, photoactivation, photoconversion, and FLIP.</a> Cold Spring Harbor Protocols. 2010 (12), (2010).</li><li>Model, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Intensity+calibration+and+flat-field+correction+for+fluorescence+microscopes.">Intensity calibration and flat-field correction for fluorescence microscopes.</a> Current Protocols in Cytometry. 68, (2014).</li><li>Schmidt, M. M., Dringen, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+effects+of+iodoacetamide+and+iodoacetate+on+glycolysis+and+glutathione+metabolism+of+cultured+astrocytes.">Differential effects of iodoacetamide and iodoacetate on glycolysis and glutathione metabolism of cultured astrocytes.</a> Frontiers in Neuroenergetics. 1, 1-10 (2009).</li><li>Surre, 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=Strong+increase+in+the+autofluorescence+of+cells+signals+struggle+for+survival.">Strong increase in the autofluorescence of cells signals struggle for survival.</a> Scientific Reports. 8 (1), 12088 (2018).</li><li>Cox, M., Lucey, S., Sridharan, S., Cohn, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Least+Squares+Congealing+for+Unsupervised+Alignment+of+Images.">Least Squares Congealing for Unsupervised Alignment of Images.</a> Proceedings of the IEEE Computer Society Conference on Computer Vision and Pattern Recognition. , (2008).</li><li>Huang, S., Heikal, A. A., Webb, W. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two-photon+fluorescence+spectroscopy+and+microscopy+of+NAD(P)H+and+flavoprotein.">Two-photon fluorescence spectroscopy and microscopy of NAD(P)H and flavoprotein.</a> Biophysical Journal. 82 (5), 2811-2825 (2002).</li><li>Fenn, J. D., Johnson, C. M., Peng, J., Jung, P., Brown, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kymograph+analysis+with+high+temporal+resolution+reveals+new+features+of+neurofilament+transport+kinetics.">Kymograph analysis with high temporal resolution reveals new features of neurofilament transport kinetics.</a> Cytoskeleton. 75 (1), 22-41 (2018).</li><li>Alami, N. H., Jung, P., Brown, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Myosin+Va+increases+the+efficiency+of+neurofilament+transport+by+decreasing+the+duration+of+long-term+pauses.">Myosin Va increases the efficiency of neurofilament transport by decreasing the duration of long-term pauses.</a> Journal of Neuroscience. 29 (20), 6625-6634 (2009).</li><li>Xu, Z., Tung, V. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temporal+and+Spatial+Variations+in+Slow+Axonal+Transport+Velocity+Along+Peripheral+Motoneuron+Axons.">Temporal and Spatial Variations in Slow Axonal Transport Velocity Along Peripheral Motoneuron Axons.</a> Neuroscience. 102 (1), 193-200 (2001).</li><li>Jung, P., Brown, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modeling+the+Slowing+of+Neurofilament+Transport+Along+the+Mouse+Sciatic+Nerve.">Modeling the Slowing of Neurofilament Transport Along the Mouse Sciatic Nerve.</a> Physical Biology. 6 (4), 046002 (2009).</li><li>Cohen, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+effect+size+index:+d.">The effect size index: d.</a> Statistical Power Analysis for the Behavioral Sciences 2nd ed. , 20-26 (1988).</li><li>Misgeld, T., Kerschensteiner, M., Bareyre, F. M., Burgess, R. W., Lichtman, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+axonal+transport+of+mitochondria+in+vivo.">Imaging axonal transport of mitochondria in vivo.</a> Nature Methods. 4 (7), 559-561 (2007).</li><li>Gilley, 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=Age-dependent+axonal+transport+and+locomotor+changes+and+tau+hypophosphorylation+in+a+"P301L"+tau+knockin+mouse.">Age-dependent axonal transport and locomotor changes and tau hypophosphorylation in a "P301L" tau knockin mouse.</a> Neurobiology of Aging. 33 (3), 1-15 (2012).</li><li>Marinkovic, P., 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+transport+deficits+and+degeneration+can+evolve+independently+in+mouse+models+of+amyotrophic+lateral+sclerosis.">Axonal transport deficits and degeneration can evolve independently in mouse models of amyotrophic lateral sclerosis.</a> Proceedings of the National Academy of Science of the United States of America. 109 (11), 4296-4301 (2012).</li><li>Milde, S., Adalbert, R., Elaman, M. H., Coleman, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Axonal+transport+declines+with+age+in+two+distinct+phases+separated+by+a+period+of+relative+stability.">Axonal transport declines with age in two distinct phases separated by a period of relative stability.</a> Neurobiology of Aging. 36 (2), 971-981 (2015).</li><li>Gibbs, K. L., Kalmar, B., Sleigh, J. N., Greensmith, L., Schiavo, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+imaging+of+axonal+transport+in+murine+motor+and+sensory+neurons.">In vivo imaging of axonal transport in murine motor and sensory neurons.</a> Journal of Neuroscience Methods. 257, 26-33 (2016).</li></ol>

The authors have nothing to disclose.

Care must be taken in the analysis of pulse-escape and pulse-spread experiments because there is significant potential for the introduction of error during the post-processing, principally during the flat-field correction, image alignment and bleach correction. Flat-field correction is necessary to correct for non-uniformity in the illumination, which results in a fall-off in intensity across the field of view from center to periphery. The extent of non-uniformity is wavelength-dependent and thus, should always be performed at the wavelength that is to be used for acquiring the experimental data. It is important to ensure that the non-uniformity in the flat-field image is truly representative of the non-uniformity in the images to be corrected. Pulse-spread experiments are particularly vulnerable to error from improper flat-field correction because the measurement ROIs extend towards the periphery of the image where the intensity fall-off is greatest.
The optimal paGFP activation settings will vary by activation method and laser parameters, and must be determined empirically before the first imaging session. Too little activation will result in a low signal-to-noise ratio for the activated fluorescence, whereas too much will result in bleaching of the activated fluorescence because both the non-activated and activated paGFP can be excited by violet light. To selectively illuminate a region of interest in the image, we use an Andor FRAPPA laser galvo scanner, which generates a clearly defined region of fluorescence with sharp boundaries, as shown in the Representative Results. We have also had success using an Andor Mosaic Digital Diaphragm, which is a digital micromirror device, with a mercury arc lamp as the illumination source7. Other options are commercially available. A challenge when working with paGFP is the delayed increase in fluorescence intensity that happens within 1 minute after the photoactivation step. This increase occurs because illumination with violet light causes a proportion of the activated paGFP molecules to enter a &#8220;dark state&#8221; from which they can relax back on a time course of tens of seconds11. In our experience, the increase is usually less than 5% with widefield excitation but can exceed 20% with laser excitation, which can lead to a significant underestimation of the activated fluorescence. We account for this by acquiring a second &#8220;post-activation&#8221; image of the paGFP fluorescence 1 minute after photoactivation, and then using this image as the reference point for the subsequent timelapse. However, it is important to recognize that this does introduce another potential source of error in determining the initial fluorescence and the decay kinetics at early times.
Additional attention must be paid to the alignment of the image stacks. The principal source of misalignment is drift or other movement of the specimen during timelapse image acquisition. This can be minimized by using inner gaskets of the appropriate thickness and allowing the preparation to &#8220;settle in&#8221; before imaging. However, any such delay will reduce the time available for image acquisition since the preparation has a limited window of viability. It is also important to avoid alignment algorithms that warp the image or introduce sub-pixel shifts, as these procedures would resample the pixel intensities in the image. Pulse-spread experiments are particularly vulnerable to errors arising from improper alignment because the borders of the region of activated fluorescence are relatively sharp, and fluorescence in this region is significantly higher than the flanking unactivated regions. Even a slight misalignment of the image planes in the stack can result in large jumps in the fluorescence values in the distal and proximal measurement windows. Thus, it is imperative that image sets are aligned properly and inspected carefully before proceeding with the analysis.
Correction for photobleaching is necessary to ensure that changes in fluorescence intensity over time accurately reflect changes in the amount of the fluorescent protein. Such corrections can be a significant source of error in estimating the absolute fluorescence intensities in the central and flanking windows. As photobleaching kinetics depend on the intensity of illumination and the environment of the fluorophore, actual photobleaching rates can vary between sessions and from axon to axon. Thus, it is necessary to measure multiple axons and average the resulting data, which itself introduces some error. The approach described above is to treat the nerve with glycolytic inhibitors and then activate the fluorescence in a region of interest and track the loss of fluorescence intensity over time. The glycolytic inhibitors deplete ATP and thus inhibit neurofilament transport so that the loss of fluorescence is due entirely to photobleaching and not movement of neurofilaments out of the activated region. The average bleaching kinetics determined in this way are then used to correct the experimental data. Since it is not practical to perform a separate bleaching calibration in each imaging session, a single calibration must be applied to multiple sessions spread over many weeks. A disadvantage of this approach is that it does not account for variations in laser power/illumination intensity from day to day, which should therefore be monitored. An alternative approach, which we refer to as &#8220;intrinsic bleaching correction&#8221;, is to estimate the photobleaching by quantifying the fluorescence in the center of the activated region in the same timelapse movies that are used for the transport measurements. If this measurement region is centrally located and much shorter than the length of the activated region, and then at short times any fluorescent neurofilaments that move out of the measurement region will be replaced by fluorescent neurofilaments that move into it. However, with time the probability of non-fluorescent neurofilaments moving into the measurement region from flanking non-fluorescent regions of the axon increases, leading to an overestimation of the loss of fluorescence due to bleaching. An advantage of this approach is that it corrects for the bleaching characteristics within that same field, but a disadvantage is that the duration of the time window must be determined empirically and will depend on the rate of the transport as well as the length of the activated region. All this having been said, it should be noted that the estimation of the directionality of neurofilament transport using our method is robust to bleaching errors (as it is for flanking window size) because it is given by the ratio of the slopes in the flanking windows and any bleaching correction is a multiplier applied to both numerator and denominator in that calculation.
Due to the noise inherent in a fluorescent system and the potential for additional error added during the image post-processing steps described above, it is important to use large sample sizes for sufficient statistical power. While it is not possible to accurately determine population means, deviations and effect sizes before experimentation, we recommend using the Cohen method21 assuming a medium effect size and an alpha of 0.05. This results in a Cohen&#8217;s d, which is the difference in means divided by the pooled standard deviation of both populations, of 0.5, which suggests acquiring at least 105 samples per group. Once this threshold has been reached, one may perform a post hoc power analysis to reassess statistical power in light of actual population measures. Under optimal experimental conditions, it should be possible to obtain from 1-7 analyzable axons (out of 9-20 total axons) per activation, and 5-8 activations per nerve assuming the acquisition of a 10-minute timelapse per activation.
Transgenic mice expressing fluorescent fusion proteins targeted to mitochondria or vesicles have also been used to study axonal transport of membranous organelles in peripheral nerves ex vivo22,23,24,25. In addition, the Schiavo lab has developed approaches to image the axonal transport of retrogradely moving membranous organelles in axons in vivo using fluorescently tagged tetanus toxin fragments, which can be injected into the muscle and are taken up by motor nerve terminals26. However, since membranous organelles can be resolved in these axons by fluorescence microscopy, it is possible to analyze the velocity and frequency of their movement directly using timelapse or kymograph analysis. The pulse-escape and pulse-spread methods described here were developed with the specific goal of analyzing the kinetics of neurofilament transport in these axons, in which single neurofilaments cannot be resolved. This requires a population-level analysis over a period of minutes or tens of minutes. We use a transgenic mouse for this purpose, but it should also be possible to express the photoactivatable protein using methods such as viral transduction or in utero electroporation. While the focus has been neurofilament transport, the pulse-escape and pulse-spread methods may also be adapted to study the movement of other cytoskeletal and cytosolic proteins that are transported along axons in the slow components of axonal transport. We confine our analyses to myelinated axons because their size and myelin sheath allows us to resolve them from their neighbors. Unmyelinated axons cannot be resolved due to their small caliber and tendency to cluster in Remak bundles. We describe the use of tibial nerves ex vivo but the methods should also be applicable to other peripheral nerves that are sufficiently long (&#62;5 mm) and unbranched. In principle, it should also be possible to adapt these methods to image neurofilament transport in vivo (e.g., by surgically exposing a nerve in a sedated mouse and then placing the mouse on a microscope stage).

The axonal transport of neurofilaments was first demonstrated in the 1970s by radioisotopic pulse-labeling1. This approach has yielded a wealth of information about neurofilament transport in vivo, but it has relatively low spatial and temporal resolution, typically on the order of millimeters and days at best2. Moreover, radioisotopic pulse-labeling is an indirect approach that requires the injection and sacrifice of multiple animals to generate a single time course. With the discovery of fluorescent proteins and advances in fluorescence microscopy in the 1990s, it subsequently became possible to image neurofilament transport directly in cultured neurons on a time scale of seconds or minutes and with sub-micrometer spatial resolution, affording much greater insight into the mechanism of movement3. These studies have revealed that neurofilament polymers in axons move rapidly and intermittently in both anterograde and retrograde directions along microtubule tracks, propelled by microtubule motor proteins. However, neurofilaments are diffraction-limited structures just 10 nm in diameter that are typically spaced apart from their neighbors by only tens of nanometers; therefore, the polymers can only be tracked in cultured neurons that contain sparsely distributed neurofilaments so that the moving polymers can be resolved from their neighbors4. Thus, it is not presently possible to track single neurofilaments in axons that contain abundant neurofilament polymers, such as myelinated axons.
To analyze the axonal transport of neurofilaments in neurofilament-rich axons using fluorescence microscopy, we use a fluorescence photoactivation pulse-escape method that we developed to study the long-term pausing behavior of neurofilaments in cultured nerve cells4,5. Neurofilaments tagged with a photoactivatable fluorescent neurofilament fusion protein are activated in a short segment of axon, and then the rate of departure of those filaments from the activated region is quantified by measuring the fluorescence decay over time. The advantage of this approach is that it is a population-level analysis of neurofilament transport that can be applied on a time-scale of minutes or hours without the need to track the movement of individual neurofilament polymers. For example, we have used this method to analyze the kinetics of neurofilament transport in myelinating cultures6.
Recently, we described the development of an hThy1-paGFP-NFM transgenic mouse that expresses low levels of a paGFP-tagged neurofilament protein M (paGFP-NFM) in neurons under the control of the human neuron-specific Thy1 promoter7. This mouse permits the analysis of neurofilament transport in situ using fluorescence microscopy. In this article, we describe the experimental approaches for analyzing neurofilament transport in myelinated axons of tibial nerves from these mice using two approaches. The first of these approaches is the pulse-escape method described above. This method can generate information about the pausing behavior of the neurofilaments, but is blind to the direction in which the filaments depart the activated region, and therefore does not permit measurement of the net directionality and transport velocity8. The second of these approaches is a new pulse-spread method in which we analyze not just the loss of fluorescence from the activated region, but also the transient increase in fluorescence in two flanking windows through which the fluorescent filaments move as they depart the activated region in both anterograde and retrograde directions. In both approaches, parameters of neurofilament transport such as the average velocity, net directionality and pausing behavior can be obtained by using mathematical analysis and modeling of the changes in fluorescence in the measurement windows. Figure 3&#160;illustrates these two approaches.
This protocol demonstrates&#160;dissection and preparation of the nerve, activation and imaging of the paGFP fluorescence, and quantification of neurofilament transport from the acquired images using the FIJI distribution package of ImageJ9. We use the tibial nerve because it is long (several cm) and does not branch; however, in principle any nerve expressing paGFP-NFM is appropriate for use with this technique if it can be dissected and de-sheathed without damaging the axons.

<table>
	<tbody>
		<tr>
			<td>14 x 22 Rectangle Gasket 0.1mm</td>
			<td>Bioptechs</td>
			<td>1907-1422-100</td>
			<td>inner gasket</td>
		</tr>
		<tr>
			<td>2-deoxy-D-glucose</td>
			<td>Sigma</td>
			<td>D6134</td>
			<td></td>
		</tr>
		<tr>
			<td>30mm Round Gasket w/ Holes</td>
			<td>Bioptechs</td>
			<td>1907-08-750</td>
			<td>outer gasket</td>
		</tr>
		<tr>
			<td>35 x 10mm dish</td>
			<td>Thermo Fisher</td>
			<td>153066</td>
			<td>dissection dishes</td>
		</tr>
		<tr>
			<td>40mm round coverslips</td>
			<td>Bioptechs</td>
			<td>40-1313-0319</td>
			<td></td>
		</tr>
		<tr>
			<td>60mL syringe - Luer-lock tip</td>
			<td>BD</td>
			<td>309653</td>
			<td></td>
		</tr>
		<tr>
			<td>Andor Revolution WD spinning-disk confocal system</td>
			<td>Andor</td>
			<td></td>
			<td>outfitted with Perfect Focus and FRAPPA systems</td>
		</tr>
		<tr>
			<td>Calcium chloride</td>
			<td>Fisher</td>
			<td>C79</td>
			<td></td>
		</tr>
		<tr>
			<td>Coverslips</td>
			<td>Fisher</td>
			<td>12-541-B</td>
			<td>for fluorescein slide</td>
		</tr>
		<tr>
			<td>D-(+)-glucose solution</td>
			<td>Sigma</td>
			<td>G8769</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting pins</td>
			<td>Fine Science Tools</td>
			<td>26001-70</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection forceps</td>
			<td>Fine Science Tools</td>
			<td>11251-30</td>
			<td>fine tipped forceps</td>
		</tr>
		<tr>
			<td>Dissection microscope</td>
			<td>Zeiss</td>
			<td>47 50 03</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection pan with wax</td>
			<td>Ginsberg Scientific</td>
			<td>568859</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection scissors</td>
			<td>Fine Science Tools</td>
			<td>14061-09</td>
			<td>initial dissection scissors</td>
		</tr>
		<tr>
			<td>FCS2 perfusion chamber</td>
			<td>Bioptechs</td>
			<td>060319-2-03</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluorescein sodium</td>
			<td>Fluka</td>
			<td>46960</td>
			<td></td>
		</tr>
		<tr>
			<td>Inline solution heater</td>
			<td>Warner Instruments</td>
			<td>SH27-B</td>
			<td></td>
		</tr>
		<tr>
			<td>Laminectomy forceps</td>
			<td>Fine Science Tools</td>
			<td>11223-20</td>
			<td>initial dissection forceps</td>
		</tr>
		<tr>
			<td>Magnesium sulfate</td>
			<td>Sigma-Aldrich</td>
			<td>M7506</td>
			<td></td>
		</tr>
		<tr>
			<td>Microaqueduct slide</td>
			<td>Bioptechs</td>
			<td>130119-5</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope slides</td>
			<td>Fisher</td>
			<td>12-544-3</td>
			<td>for fluorescein slide</td>
		</tr>
		<tr>
			<td>Microscope stage insert</td>
			<td>Applied Scientific Instrumentation</td>
			<td>I-3017</td>
			<td></td>
		</tr>
		<tr>
			<td>Objective heater system</td>
			<td>Okolab</td>
			<td></td>
			<td>Oko Touch with objective collar</td>
		</tr>
		<tr>
			<td>Objective oil - type A</td>
			<td>Nikon</td>
			<td>discontinued</td>
			<td></td>
		</tr>
		<tr>
			<td>Plan Apo VC 100x 1.40 NA objective</td>
			<td>Nikon</td>
			<td>MRD01901</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium chloride</td>
			<td>Fisher</td>
			<td>P217</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium phosphate</td>
			<td>Sigma-Aldrich</td>
			<td>P0662</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>Sigma-Aldrich</td>
			<td>S6297</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Sigma-Aldrich</td>
			<td>S7653</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium iodoacetate</td>
			<td>Sigma-Aldrich</td>
			<td>I2512</td>
			<td></td>
		</tr>
		<tr>
			<td>Syringe pump</td>
			<td>Sage Instruments</td>
			<td>Model 355</td>
			<td></td>
		</tr>
		<tr>
			<td>Tubing adapter - female</td>
			<td>Small Parts Inc.</td>
			<td>1005109</td>
			<td></td>
		</tr>
		<tr>
			<td>Tubing adapter - male</td>
			<td>Small Parts Inc.</td>
			<td>1005012</td>
			<td></td>
		</tr>
		<tr>
			<td>Tygon tubing</td>
			<td>Bioptechs</td>
			<td></td>
			<td>1/16&#34; ID, 1/32&#34; wall thickness</td>
		</tr>
		<tr>
			<td>Vannas spring scissors</td>
			<td>Fine Science Tools</td>
			<td>15018-10</td>
			<td>fine scissors</td>
		</tr>
	</tbody>
</table>

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.

Watch this Scientific Journal Video about Imaging and Analysis of Neurofilament Transport in Excised Mouse Tibial Nerve at JoVE.com

Imaging and Analysis of Neurofilament Transport in Excised Mouse Tibial Nerve

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 ...

imaging-analysis-neurofilament-transport-excised-mouse-tibial

Nanyang Technological University

Research

JoVE Journal

Neuroscience

6.0K Views.  The Ohio State University. 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.

Video: Imaging and Analysis of Neurofilament Transport in Excised Mouse Tibial Nerve

Organotypic Slice Culture of GFP-expressing Mouse Embryos for Real-time Imaging of Peripheral Nerve Outgrowth

For many purposes, the cultivation of mouse embryos ex vivo as organotypic slices is desirable. For example, we employ a transgenic mouse line (tauGFP) in which the enhanced version of the green fluorescent protein (EGFP) is exclusively expressed in all neurons of the developing central and peripheral nervous system1, allowing the possibility to both film the innervation of the forelimb and to manipulate this process with pharmacological and genetic techniques2. The most critical parameter in the successful cultivation of such slice cultures is the method by which the slices are prepared. After extensive testing of a variety of methods, we have found that a vibratome is the best possible device to slice the embryos such that they routinely result in a culture that demonstrates viability over a period of several days, and most importantly, develops in an age-specific manner. For mid-gestation embryos, this includes the normal outgrowth of spinal nerves from the spinal cord and the dorsal root ganglia to their targets in the periphery and the proper determination of skeletal and muscle tissue. 
In this work, we present a method for processing whole embryos of embryonic day (E) E10 to E12 into 300 - 400 micrometer slices for cultivation in a standard tissue culture incubator, which can be studied for up to two days after slice preparation. Critical for the success of this approach is the use of a vibratome to slice each agarose-embedded embryo. This is followed by the cultivation of the slices upon Millicell culture membrane inserts placed upon a small volume of medium, resulting in an interface culture technique. One litter with an average of 7 embryos routinely produces at least 14 slices (2-3 slices of the forelimb region per embryo), which varies slightly due to the age of the embryos as well as to the thickness of the slices. About 80% of the cultured slices show nerve outgrowth, which can be measured througout the culturing period2. Representative results using the tauGFP mouse line are demonstrated.

We present a method to prepare organotypic slices of mid-gestation mouse embryos for the cultivation and time-lapse imaging of peripheral nerve outgrowth.

For many purposes, the cultivation of mouse embryos ex vivo as organotypic slices is desirable. For example, we employ a transgenic mouse line (tauGFP) in ...

Multiple-mouse Neuroanatomical Magnetic Resonance Imaging

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and evaluating mouse models of human disease. MRI is an excellent modality for investigating genetically altered animals. It is capable of whole brain coverage, can be used in vivo, and provides multiple contrast mechanisms for investigating different aspects of neuranatomy and physiology. The advent of high-field scanners along with the ability to scan multiple mice simultaneously allows for rapid phenotyping of novel mutations.
Effective mouse MRI studies require attention to many aspects of experiment design. In this article, we will describe general methods to acquire quality images for mouse phenotyping using a system that images mice concurrently in shielded transmit/receive radio frequency (RF) coils in a common magnet (Bock et al., 2003). We focus particularly on anatomical phenotyping, an important and accessible application that has shown a high potential for impact in many mouse models at our imaging centre. Before we can provide the detailed steps to acquire such images, there are important practical considerations for both in vivo brain imaging (Dazai et al., 2004) and ex vivo brain imaging (Spring et al., 2007) that should be noted. These are discussed below.

Magnetic resonance imaging (MRI) has become an increasingly popular tool for examining the phenotype of genetically altered mice. This article illustrates the methods necessary to achieve high-throughput phenotyping of genetically altered mice using multiple-mouse MRI.

The field of mouse phenotyping with magnetic resonance imaging (MRI) is rapidly growing, motivated by the need for improved tools for characterizing and ...

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured neurons for extended durations1-3. This is now possible through the use of vital dyes and fluorescent proteins with which cytoskeletal components, organelles, and other structures in living cells can be labeled and then visualized via dynamic fluorescence microscopy. For example, in embryonic chicken sympathetic neurons, mitochondrial movement was characterized using the vital dye rhodamine 1234. In another study, mitochondria were visualized in rat forebrain neurons by transfection of mitochondrially targeted eYFP5. However, imaging of primary neurons over minutes, hours, or even days presents a number of issues. Foremost among these are: 1) maintenance of culture conditions such as temperature, humidity, and pH during long imaging sessions; 2) a strong, stable fluorescent signal to assure both the quality of acquired images and accurate measurement of signal intensity during image analysis; and 3) limiting exposure times during image acquisition to minimize photobleaching and avoid phototoxicity.
Here, we describe a protocol that permits the observation, visualization, and analysis of mitochondrial movement in cultured hippocampal neurons with high temporal resolution and under optimal life support conditions. We have constructed an affordable stage-top incubator that provides good temperature regulation and atmospheric gas flow, and also limits the degree of media evaporation, assuring stable pH and osmolarity. This incubator is connected, via inlet and outlet hoses, to a standard tissue culture incubator, which provides constant humidity levels and an atmosphere of 5-10% CO2/air. This design offers a cost-effective alternative to significantly more expensive microscope incubators that don't necessarily assure the viability of cells over many hours or even days. To visualize mitochondria, we infect cells with a lentivirus encoding a red fluorescent protein that is targeted to the mitochondrion. This assures a strong and persistent signal, which, in conjunction with the use of a stable xenon light source, allows us to limit exposure times during image acquisition and all but precludes photobleaching and phototoxicity. Two injection ports on the top of the stage-top incubator allow the acute administration of neurotransmitters and other reagents intended to modulate mitochondrial movement. In sum, lentivirus-mediated expression of an organelle-targeted red fluorescent protein and the combination of our stage-top incubator, a conventional inverted fluorescence microscope, CCD camera, and xenon light source allow us to acquire time-lapse images of mitochondrial transport in living neurons over longer durations than those possible in studies deploying conventional vital dyes and off-the-shelf life support systems.

We describe a protocol that allows imaging of mitochondria in living neurons via fluorescence microscopy over long durations. Imaging over extended periods is accomplished through lentivirus-mediated expression of a mitochondrially targeted fluorescent protein and use of an inexpensive stage-top incubator that was designed and built in our laboratory.

To understand the relationship between mitochondrial transport and neuronal function, it is critical to observe mitochondrial behavior in live cultured ...

Reproducible Mouse Sciatic Nerve Crush and Subsequent Assessment of Regeneration by Whole Mount Muscle Analysis

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative response mounted by PNS neurons thereby possibly illuminating the failures of CNS regeneration1. Sciatic nerve crush (axonotmesis) is one of the most common models of peripheral nerve injury in rodents2. Crushing interrupts all axons but Schwann cell basal laminae are preserved so that regeneration is optimal3,4. This allows the investigator to study precisely the ability of a growing axon to interact with both the Schwann cell and basal laminae4. Rats have generally been the preferred animal models for experimental nerve crush. They are widely available and their lesioned sciatic nerve provides a reasonable approximation of human nerve lesions5,4. Though smaller in size than rat nerve, the mouse nerve has many similar qualities. Most importantly though, mouse models are increasingly valuable because of the wide availability of transgenic lines now allows for a detailed dissection of the individual molecules critical for nerve regeneration6, 7. Prior investigators have used multiple methods to produce a nerve crush or injury including simple angled forceps, chilled forceps, hemostatic forceps, vascular clamps, and investigator-designed clamps8,9,10,11,12. Investigators have also used various methods of marking the injury site including suture, carbon particles and fluorescent beads13,14,1. We describe our method to obtain a reproducibly complete sciatic nerve crush with accurate and persistent marking of the crush-site using a fine hemostatic forceps and subsequent carbon crush-site marking. As part of our description of the sciatic nerve crush procedure we have also included a relatively simple method of muscle whole mount we use to subsequently quantify regeneration.

In this report we describe a method to crush mouse sciatic nerve. This method uses readily available hemostatic forceps and easily and reproducibly produces complete sciatic nerve crush. In addition, we describe a method to prepare muscle whole mounts suitable for analysis of nerve regeneration after sciatic nerve crush.

Regeneration in the peripheral nervous system (PNS) is widely studied both for its relevance to human disease and to understand the robust regenerative ...

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date this tool has been widely used to study the regulation of cellular proliferation, differentiation and neuronal migration especially in the developing cerebral cortex 6-8. Here we detail our protocol to electroporate in utero the cerebral cortex and the hippocampus and provide evidence that this approach can be used to study dendrites and spines in these two cerebral regions.
Visualization and manipulation of neurons in primary cultures have contributed to a better understanding of the processes involved in dendrite, spine and synapse development. However neurons growing in vitro are not exposed to all the physiological cues that can affect dendrite and/or spine formation and maintenance during normal development. Our knowledge of dendrite and spine structures in vivo in wild-type or mutant mice comes mostly from observations using the Golgi-Cox method 9. However, Golgi staining is considered to be unpredictable. Indeed, groups of nerve cells and fiber tracts are labeled randomly, with particular areas often appearing completely stained while adjacent areas are devoid of staining. Recent studies have shown that IUE of fluorescent constructs represents an attractive alternative method to study dendrites, spines as well as synapses in mutant / wild-type mice 10-11 (Figure 1A). Moreover in comparison to the generation of mouse knockouts, IUE represents a rapid approach to perform gain and loss of function studies in specific population of cells during a specific time window. In addition, IUE has been successfully used with inducible gene expression or inducible RNAi approaches to refine the temporal control over the expression of a gene or shRNA 12. These advantages of IUE have thus opened new dimensions to study the effect of gene expression/suppression on dendrites and spines not only in specific cerebral structures (Figure 1B) but also at a specific time point of development (Figure 1C).
Finally, IUE provides a useful tool to identify functional interactions between genes involved in dendrite, spine and/or synapse development. Indeed, in contrast to other gene transfer methods such as virus, it is straightforward to combine multiple RNAi or transgenes in the same population of cells. 
In summary, IUE is a powerful method that has already contributed to the characterization of molecular mechanisms underlying brain function and disease and it should also be useful in the study of dendrites and spines.

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

This article describes in detail a protocol to electroporate in utero the cerebral cortex and the hippocampus at E14.5 in mice. We also show that this is a valuable method to study dendrites and spines in these two cerebral regions.

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date ...

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 ...

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 ...

In Vivo Multimodal Imaging and Analysis of Mouse Laser-Induced Choroidal Neovascularization Model

Laser-induced choroidal neovascularization (CNV) is a well-established model to mimic the wet form of age-related macular degeneration (AMD). In this protocol, we aim to guide the reader not simply through the technical considerations of generating laser-induced lesions to trigger neovascular processes, but rather focus on the powerful information that can be obtained from multimodal longitudinal in vivo imaging throughout the follow-up period.
The laser-induced mouse CNV model was generated by a diode laser administration. Multimodal in vivo imaging techniques were used to monitor CNV induction, progression and regression. First, spectral domain optical coherence tomography (SD-OCT) was performed immediately after the lasering to verify a break of Bruch&#39;s membrane. Subsequent in vivo imaging using fluorescein angiography (FA) confirmed successful damage of Bruch&#39;s membrane from serial images acquired at the choroidal level. Longitudinal follow-up of CNV proliferation and regression on days 5, 10, and 14 after the lasering was performed using both SD-OCT and FA. Simple and reliable grading of leaky CNV leasions from FA images is&#160;presented. Automated segmentation for measurement of total retinal thickness, combined with manual caliber application for measurement of retinal thickness at CNV sites, allow unbiased evaluation of the presence of edema. Finally, histological verification of CNV is performed using isolectin GS-IB4 staining on choroidal flatmounts. The staining is thresholded, and the isolectin-positive area is calculated with ImageJ.
This protocol is especially useful in therapeutics studies requiring high-throughput-like screening of CNV pathology as it allows fast, multimodal, and reliable classification of CNV pathology and retinal edema. In addition, high resolution SD-OCT enables the recording of other pathological hallmarks, such as the accumulation of subretinal or intraretinal fluid. However, this method does not provide a possibility to automate CNV volume analysis from SD-OCT images, which has to be performed manually.

In Vivo Multimodal Imaging and Analysis of Mouse Laser-Induced Choroidal Neovascularization Model

Here, we present the usefulness of longitudinal in vivo imaging in the follow-up of morphological changes of laser-induced choroidal neovascularization in mice.

Laser-induced choroidal neovascularization (CNV) is a well-established model to mimic the wet form of age-related macular degeneration (AMD). In this ...

In Vivo Imaging of Cerebrospinal Fluid Transport through the Intact Mouse Skull using Fluorescence Macroscopy

Cerebrospinal fluid (CSF) flow in rodents has largely been studied using ex vivo quantification of tracers. Techniques such as two-photon microscopy and magnetic resonance imaging (MRI) have enabled in vivo quantification of CSF flow but they are limited by reduced imaging volumes and low spatial resolution, respectively. Recent work has found that CSF enters the brain parenchyma through a network of perivascular spaces surrounding the pial and penetrating arteries of the rodent cortex. This perivascular entry of CSF is a primary driver of the glymphatic system, a pathway implicated in the clearance of toxic metabolic solutes (e.g., amyloid-&#946;). Here, we illustrate a new macroscopic imaging technique that allows real-time, mesoscopic imaging of fluorescent CSF tracers through the intact skull of live mice. This minimally-invasive method facilitates a multitude of experimental designs and enables single or repeated testing of CSF dynamics. Macroscopes have high spatial and temporal resolution and their large gantry and working distance allow for imaging while performing tasks on behavioral devices. This imaging approach has been validated using two-photon imaging and fluorescence measurements obtained from this technique strongly correlate with ex vivo fluorescence and quantification of radio-labeled tracers. In this protocol, we describe how transcranial macroscopic imaging can be used to evaluate glymphatic transport in live mice, offering an accessible alternative to more costly imaging modalities.

Transcranial optical imaging allows wide-field imaging of cerebrospinal fluid transport in the cortex of live mice through an intact skull.

Cerebrospinal fluid (CSF) flow in rodents has largely been studied using ex vivo quantification of tracers. Techniques such as two-photon microscopy and ...

3D Kinematic Analysis for the Functional Evaluation in the Rat Model of Sciatic Nerve Crush Injury

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of sciatic nerve injury rodent models. In this protocol, we describe a novel kinematic analysis method that uses a three-dimensional (3D) motion capture apparatus for functional evaluations using a rat sciatic nerve crush injury model. First, the rat is familiarized with treadmill walking. Markers are then attached to the designated bone landmarks and the rat is made to walk on the treadmill at the desired speed. Meanwhile, the posterior limb movements of the rat are recorded using four cameras. Depending on the software used, marker tracings are created using both automatic and manual modes and the desired data are produced after subtle adjustments. This method of kinematic analysis, which uses a 3D motion capture apparatus, offers numerous advantages, including superior precision and accuracy. Many more parameters can be investigated during the comprehensive functional evaluations. This method has several shortcomings that require consideration: The system is expensive, can be complicated to operate, and may produce data deviations due to skin shifting. Nevertheless, kinematic analysis using a 3D motion capture apparatus is useful for performing functional anterior and posterior limb evaluations. In the future, this method may become increasingly useful for generating accurate assessments of various traumas and diseases.

We introduce a kinematic analysis method that uses a three-dimensional motion capture apparatus containing four cameras and data processing software for performing functional evaluations during fundamental research involving rodent models.

Compared to the Sciatic Functional Index (SFI), kinematic analysis is a more reliable and sensitive method for performing functional evaluations of ...

Imaging and Analysis of Neurofilament Transport in Excised Mouse Tibial Nerve

Summary

Explore More Videos

Chapters in this video

Organotypic Slice Culture of GFP-expressing Mouse Embryos for Real-time Imaging of Peripheral Nerve Outgrowth

Multiple-mouse Neuroanatomical Magnetic Resonance Imaging

Neuromodulation and Mitochondrial Transport: Live Imaging in Hippocampal Neurons over Long Durations

Reproducible Mouse Sciatic Nerve Crush and Subsequent Assessment of Regeneration by Whole Mount Muscle Analysis

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

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

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

In Vivo Multimodal Imaging and Analysis of Mouse Laser-Induced Choroidal Neovascularization Model

In Vivo Imaging of Cerebrospinal Fluid Transport through the Intact Mouse Skull using Fluorescence Macroscopy

3D Kinematic Analysis for the Functional Evaluation in the Rat Model of Sciatic Nerve Crush Injury