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 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 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.
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 illustrates these two approaches.
This protocol demonstrates 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.
All methods described here have been approved by the Institutional Animal Care and Use Committee (IACUC) of The Ohio State University.
1. Preparation of nerve saline solution
2. Initial assembly of nerve perfusion chamber
3. Dissection and preparation of mouse tibial nerve
4. Final nerve perfusion chamber assembly
5. Fluorescence activation and image acquisition
6. Flatfield and darkfield image acquisition
7. Imaging glycolytically inhibited nerves for bleach correction
8. Image processing and analysis using ImageJ
9. Photobleach correction
Figure 3 shows representative images from pulse-escape and pulse-spread experiments. We have published several studies that describe data obtained using the pulse-escape method and our methods for the analysis of those data5,6,7,8,17. Below, we show how the pulse-spread data can yield information on the directionality and velocity of neurofilament transport, which we have not reported previously.
Neurofilament transport in the axon is intermittent and bidirectional. This transport can be described by the fraction of filaments moving at any given time in the anterograde and retrograde directions, and respectively, and their velocities in the anterograde and retrograde directions, denoted by and . If we take the total quantity of neurofilament polymer per unit length of axon to be , then the fluxes in the anterograde and retrograde directions through a given region are given by
and
,
respectively, and the total flux is given by
,
where has units µm-1, has units and has units . Since the flux at a given location along the axon is the quantity of neurofilament polymer that moves past that location in a unit of time, it is related to the average velocity through . Thus, we can write
.
In a pulse-spread experiment the flux can be determined from the rate of departure of the fluorescent neurofilaments from the activated region, which we refer to as the central window. The total loss of fluorescent neurofilament polymer from this central window per second is the sum of the losses due to fluorescent neurofilament polymers leaving in anterograde and retrograde directions, i.e.
Normalized to the initial content of fluorescent neurofilament polymer in the central window, i.e. , where is the length of the central window, this rate of loss then becomes
where is the slope of the decrease in fluorescence in the central window, which is initially linear for a time period given by 8. For a 40 µm central window, this equates to just tens of seconds. However, for windows this large, the transition to the exponential decay phase is gradual and the slope is effectively linear for several minutes or more8.
At early times, the flanking windows capture all the neurofilaments that exit the central window because these filaments do not have sufficient time to pass through the flanking windows and exit them on the other side. In this case, the quantities of fluorescent neurofilament polymer that leave the central window anterogradely and retrogradely per second, i.e. the fluxes and , are given by the increase in the neurofilament content and in the flanking windows. Normalized to the initial content of fluorescent neurofilament polymer in the central window, the rates of increase in the flanking windows become
where and are the slopes of the increase in fluorescence in the proximal and distal flanking windows, respectively.
Thus, we can express the average velocity in terms of the slopes in the flanking windows, i.e.
(Eq. 1)
and the ratio of the number of anterograde and retrograde moving neurofilaments in terms of the ratio of these slopes, i.e.
(Eq. 2)
where and denote the rates at which neurofilaments reverse from anterogradely to retrogradely moving and vice versa8. The values of and can be determined by measuring the movement of individual neurofilaments in cultured neurons, as reported previously17. Importantly, the expression for the velocity in Eq. 1 only applies at short times after activation during which fluorescent neurofilaments enter the flanking window but do not leave. The duration of this short time window will depend on the length of the flanking windows and the kinetics of the neurofilament movement. The longer the flanking windows, in principle, the longer the time window. Theoretically, one can test that this criterion is met by confirming that
(Eq. 3)
In contrast to the expression for the velocity in Eq. 1, which is given by the difference between the slopes in the flanking windows, the expression for the directionality in Eq. 2 is robust to flanking window size because it is given by the ratio of the slopes in the flanking windows.
Figure 3 shows representative results from pulse-escape and pulse-spread experiments on myelinated axons in the tibial nerve of an 8-week old male mouse from our hThy1 paGFP-NFM line on a C57Bl/6J background, using window sizes of 5 and 40 µm, respectively. Mice of at least 2 weeks of age, both male and female, have been used for these experimental paradigms. The appropriate age and sex should be determined by the researcher depending on what is being tested in the study. Over time, it can be seen that the edges of the activated regions blur due to the departure of neurofilaments in both anterograde and retrograde directions, resulting in loss of fluorescence from the central window.
For the pulse-escape method (Figure 3A, C, E), the fluorescence decay kinetics in the activated region can yield information on the long-term and short-term pausing behavior8,18. For these experiments, the activated region can be short (we typically use 5 µm; see yellow box in Figure 3A). For the pulse-spread method (Figure 3B, D, F), we use a longer activated region (40 µm here; see yellow box in Figure 3B) in order to provide a larger pool of fluorescent neurofilaments. This lengthens the time over which the fluorescence increase in the flanking regions remains linear (see above). The linear domain of this fluorescence increase in the flanking windows can also be increased by increasing the length of these windows, however this is limited by the size of the field of view. We used 15 μm flanking windows, shown in red and green, for the data shown in Figure 3D.
Figure 3E,3F shows the quantification of the total fluorescence intensities (i.e., the sum of the pixel intensities) for the measurement windows shown in Figure 3C,3D respectively. For the pulse-escape method the decay is biphasic, with an initial exponential decay which represents the departure of on-track neurofilaments. This transitions at around 10-20 minutes to a second slower exponential decay that represents the mobilization and departure of off-track filaments8. For comparison with the pulse-spread method, we show only a 12 minute time course in Figure 3E, but usually the time course of the analysis needs to be longer (typically 30-120 minutes) to capture the long-term pausing kinetics5,6,18. For the pulse-spread strategy, the calculations of the velocity and directionality of the movement are dependent only on the slopes in the flanking windows. For the window lengths used here, the linear phase extends for about 5 minutes. The duration of this time window of linearity should be assessed by incrementally shortening the time used to measure the slope, and determining the point at which the slope no longer increases. This duration can be increased by increasing the window lengths if the camera field of view permits, though the window lengths should be held constant for all axons in a given experiment. As an example, we have used the first 5 minutes of data to measure the slopes for the pulse-spread data in Figure 3F. This results in slopes of 72 A.U./min, -594 A.U./min, and 111 A.U./min for the proximal, central and distal windows, respectively (A.U. = arbitrary units). In order to normalize these values, they are divided by the initial fluorescence of the central window, resulting in rates of 0.108 %F0/min, -0.962 %F0/min and 0.173 %F0/min. Applying Eq. 3 we find that the conservation criterion is not met, indicating that we are not capturing all of the fluorescence in the flanking windows. This is a technical limitation due to the field of view of our EMCCD camera (82 µm x 82 µm). Cameras with sCMOS chips, which can have much larger fields of view, could permit the use of larger flanking window sizes. However, we cannot exclude the possibility that this discrepancy between the central and flanking slopes could also be due, at least in part, to underestimation of the extent of photobleaching (see Discussion), which would have the effect of underestimating the positive slopes in the flanking windows and overestimating the negative slope in the central window.
From the rates in the flanking windows (%F0/min) and Eq. 2 above, and using values of and 17, we calculate the ratio = 2.12. This indicates that 68% of the filaments were moving anterograde and 32% retrograde. Using the same rates and Eq. 1 above, we calculate the average net population velocity = 40*(0.00173-0.00108) = 0.026 μm/min, or 0.037 mm/day. Radioisotopic pulse-labeling studies in mice of similar age have reported a neurofilament population velocity of approximately 0.6 mm/day in the most proximal portions of the sciatic nerve, slowing to 0.12 mm/day over a distance of two centimeters19,20. Our pulse-spread data was gathered in the tibial nerve, roughly another 2-3 centimeters distal to these measures. For the reasons mentioned above, we believe that the estimate of 0.037 mm/day is an underestimate of the true velocity. However, extrapolating the spatial slowing observed by radioisotopic pulse-labeling into the tibial nerve this estimate not unreasonable, especially when we consider that it was determined using a very different methodology on a very different spatial and temporal scale.
To demonstrate the capability of the pulse-spread method to detect significant differences between populations, we compared the proximal and distal flanking window slopes measured from nerves perfused with both normal and inhibitor salines. Inhibition of glycolysis blocks movement of neurofilaments, as we have reported previously5,6,7. We will discuss later the selection of sample sizes for experiments, however here we used one nerve in normal saline and two in inhibitory saline, due to the limitation of one acquisition field per nerve during inhibition. Figure 4A shows an example timelapse from a glycolytically inhibited nerve, demonstrating an apparent reduction in transport out of the activated region. Indeed, we find significantly lower distal and proximal slopes (Figure 4B, p = 0.00000639 and 0.0121, respectively, Tukey’s pairwise comparison following ANOVA) in nerves treated with inhibitory saline versus those perfused with normal saline. We also find a significant decrease in the population velocity between these two conditions (Figure 4C, p = 0.0232, t-test with unequal variances, following F test for equal variance with p = 0.0190).
Figure 1: The perfusion chamber. (A) Diagram showing the assembly of the perfusion chamber housing and the outer gasket, with tubing connected to the saline syringe and waste flask. (B) Diagram showing the assembly of the chamber itself. The inner gasket is laid flat on top of the #1.5 coverslip. The nerve is placed on the coverslip in the well created by the rectangular opening of the inner gasket. The microaqueduct slide is placed over the nerve to sandwich the nerve between the #1.5 coverslip and the microaqueduct slide. Finally, the sandwich is flipped before mounting on top of the outer gasket of the assembly shown in (A) and secured with a locking ring (not shown). Please click here to view a larger version of this figure.
Figure 2: The tibial nerve preparation. (A) Image showing the location of the tibial nerve within the leg of a mouse, in which the gluteus superficialis, biceps femoris, semitendinosus, popliteus, tibialis caudalis and flexor digitorum longus muscles have been removed. The tibial nerve can be seen as one of three major branches of the sciatic nerve arising at the knee. (B) Schematic of the nerve in cross-section in the assembled chamber, showing the oil-immersion objective beneath the coverslip. The best optical quality is obtained by activating and imaging the layer of axons apposed to the surface of the coverslip; image quality declines deeper into the nerve due to light scattering. (C) Examples of a healthy axon (top), and of an unhealthy axon (bottom) immediately after activation. The dashed yellow line marks the activated region. The activated fluorescence has sharp boundaries in the healthy axon whereas in the unhealthy axon the activated fluorescence diffuses rapidly out of the activated region, filling the axon within seconds. Scale bar = 10 μm. (D) Examples of the mitochondrial appearance in a healthy axon (top) and an unhealthy axon (bottom). The mitochondria are visible due to flavin autofluorescence16. They appear linear (solid arrowheads) in healthy axons. In unhealthy axons, the mitochondria first become punctate (open arrowheads) and then subsequently fainter over time, providing an indicator of axon health. The dashed open arrowhead in the healthy axon points to a mitochondrion transitioning from linear to punctate. Scale bar = 10 μm. Please click here to view a larger version of this figure.
Figure 3: Pulse-escape and pulse-spread experiments. Example of pre-activation, post-activation and timelapse images of myelinated axons in the tibial nerve of an 8 week old mouse in (A) a pulse-escape experiment and (B) a pulse-spread experiment. The pre-activation image is the top panel, with the post-activation images below. The time stamps show the time elapsed after activation. The yellow boxes represent the region activated. These activations were performed using 5 scans with a 40 μs pixel dwell time and 5% laser power. Scale bars = 10 μm. (C) The measurement ROIs (yellow) for three axons in a pulse-escape experiment. (D) The proximal (red), central (yellow) and distal (green) measurement ROIs for three axons in a pulse-spread experiment. The measurement ROIs for each axon are shown in solid red (proximal), solid yellow (central window) and solid green (distal window). (E) Plot of the average fluorescence versus time, average of the 3 axons in C. Dashed line is an exponential function fit to the data, of the form Ft = Ae-τt, as described previously7. F) Plots of the fluorescence in the central, distal and proximal ROIs versus time, average of 14 axons including the three axons in D. The slope is calculated from trendlines fit to the first 5 minutes of data. Please click here to view a larger version of this figure.
Figure 4: The effect of metabolic inhibitors on neurofilament transport. (A) Example of timelapse images of a nerve pretreated with 5.6% (w/v) 2-deoxy-glucose and 0.5 mM sodium iodoacetate to inhibit glycolysis and deplete cellular ATP. Note that the distal and proximal boundaries of the activated regions remain sharp, indicative of an inhibition of neurofilament transport. Scale bar = 10 µm. (B) Quantification of the slopes in the distal (D) and proximal (P) flanking windows (anterograde and retrograde, respectively), expressed as percent of initial fluorescence in the central window (%F0/min), from one nerve (14 axons) using standard saline and from two nerves (8 axons) pretreated with saline containing the glycolytic inhibitors. (C) Neurofilament population velocities calculated from slopes in the flanking windows, showing a significant decrease in velocity following inhibitor treatment. *** – p < 0.005; ** – p < 0.01; * – p < 0.05. The data show significant impairment of neurofilament transport in the presence of the inhibitors. Please click here to view a larger version of this figure.
Supplemental Video. Please click here to download this video.
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 “dark state” 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 “post-activation” 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 “settle in” 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 “intrinsic bleaching correction”, 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’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 (>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 authors have nothing to disclose.
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’s Postdoctoral Scholars Program.
14 x 22 Rectangle Gasket 0.1mm | Bioptechs | 1907-1422-100 | inner gasket |
2-deoxy-D-glucose | Sigma | D6134 | |
30mm Round Gasket w/ Holes | Bioptechs | 1907-08-750 | outer gasket |
35 x 10mm dish | Thermo Fisher | 153066 | dissection dishes |
40mm round coverslips | Bioptechs | 40-1313-0319 | |
60mL syringe – Luer-lock tip | BD | 309653 | |
Andor Revolution WD spinning-disk confocal system | Andor | outfitted with Perfect Focus and FRAPPA systems | |
Calcium chloride | Fisher | C79 | |
Coverslips | Fisher | 12-541-B | for fluorescein slide |
D-(+)-glucose solution | Sigma | G8769 | |
Dissecting pins | Fine Science Tools | 26001-70 | |
Dissection forceps | Fine Science Tools | 11251-30 | fine tipped forceps |
Dissection microscope | Zeiss | 47 50 03 | |
Dissection pan with wax | Ginsberg Scientific | 568859 | |
Dissection scissors | Fine Science Tools | 14061-09 | initial dissection scissors |
FCS2 perfusion chamber | Bioptechs | 060319-2-03 | |
Fluorescein sodium | Fluka | 46960 | |
Inline solution heater | Warner Instruments | SH27-B | |
Laminectomy forceps | Fine Science Tools | 11223-20 | initial dissection forceps |
Magnesium sulfate | Sigma-Aldrich | M7506 | |
Microaqueduct slide | Bioptechs | 130119-5 | |
Microscope slides | Fisher | 12-544-3 | for fluorescein slide |
Microscope stage insert | Applied Scientific Instrumentation | I-3017 | |
Objective heater system | Okolab | Oko Touch with objective collar | |
Objective oil – type A | Nikon | discontinued | |
Plan Apo VC 100x 1.40 NA objective | Nikon | MRD01901 | |
Potassium chloride | Fisher | P217 | |
Potassium phosphate | Sigma-Aldrich | P0662 | |
Sodium bicarbonate | Sigma-Aldrich | S6297 | |
Sodium chloride | Sigma-Aldrich | S7653 | |
Sodium iodoacetate | Sigma-Aldrich | I2512 | |
Syringe pump | Sage Instruments | Model 355 | |
Tubing adapter – female | Small Parts Inc. | 1005109 | |
Tubing adapter – male | Small Parts Inc. | 1005012 | |
Tygon tubing | Bioptechs | 1/16" ID, 1/32" wall thickness | |
Vannas spring scissors | Fine Science Tools | 15018-10 | fine scissors |