Name: imaging and analysis of neurofilament transport in excised mouse tibial nerve
Uploaded: 2020-08-31T13:00:00
Description: Watch this Scientific Journal Video about Imaging and Analysis of Neurofilament Transport in Excised Mouse Tibial Nerve at JoVE.com

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.

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
<ol>
	<li>Make 100 mL of Breuer&#8217;s saline10: 98 mM NaCl, 1 mM KCl, 2 mM KH2PO4, 1 mM MgSO4, 1.5 mM CaCl2, 5.6% D-glucose, 23.8 mM NaHCO3 in double-distilled water.</li>
	<li>Bubble 95% oxygen/5% carbon dioxide (carbogen) through the sali...

<ol>
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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 perfor...

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>
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			<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>
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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.

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...

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 ...

This protocol allows direct imaging of the neurofilament population in myelinated axons from juvenile or adult mice using fluorescence microscopy, making it possible to measure the directionality and velocity of neurofilament transport in isolated nerve segments. This method depends on the availability of mice expressing a photoactivatable fluorescent fusion of the protein of interest, but can be adapted for use in any readily dissected nerve. To begin, pour oxygenated saline into a 60 milliliter syringe and ensure that there is minimal air remaining in the syringe.Connect the syringe to the tubing, then place the outflow tube into a waste flask. Place the outer gasket into the perfusion chamber housing, ensuring that the flow inlet and outlet posts are aligned with the holes in the gasket, then lay the inner gasket on a number 1.5 circular coverslip and carefully smooth out any wrinkles in the gasket to ensure a tight seal. Place the coverslip and gasket on a paper towel or a task wipe with the gasket facing up.Use a pair of large dissection scissors to make a dorsal incision in the skin near the middle of the spine and continue to cut around the ventral aspect of the animal. Slowly pull the skin away from the muscle and cut the fascia. Next, use microdissection scissors to make an incision in the thigh muscles midway between the tail and knee to expose the sciatic nerve.Make sure that the nerve, which is visible through the muscle, is not cut. Extend the incision dorsally and ventrally to remove the muscle, then remove the muscles of the calf, keeping the cuts shallow and short to avoid damaging nerves. Continue removing the muscles until the tibial nerve is fully exposed from the point where it branches from the sciatic nerve to the heel.Grasp the tibial nerve at the spine proximal end with a pair of forceps and cut it with a pair of microdissection scissors, making sure to not apply too much tension to the nerve. Carefully lift it away from the muscle and cut any attachments, taking care not to put tension on the nerve. Cut the spine distal end of the tibial nerve and transfer it to a small Petri dish of room temperature oxygenated saline.Starting from the proximal end of the nerve, gently grasp the exposed axon ends with a pair of very fine tip forceps. With a second pair of forceps, grasp the nerve sheath proximally and slowly pull towards the distal end of the nerve making sure that no undue tension is applied during this process. Grasp the proximal end of the nerve and slowly lay it down onto a coverslip, then straighten it under gentle tension.Place the microaqueduct slide over the nerve with the grooved slide down and position the direction of flow parallel to the nerve. Flip the coverslip and microaqueduct assembly over and place it in the perfusion chamber housing with the slide opposed to the outer gasket. To secure the profusion chamber, place it in the metal housing and rotate the locking ring.Slowly depress the saline syringe plunger and fill the perfusion chamber. Keep the inlet and outlet tubing, outlet flask, and syringe elevated above the chamber itself at all times during the setup and imaging. Transfer the profusion assembly to an inverted microscope stage and mount the saline syringe into the syringe pump, then start the motor and adjust the speed for a flow rate of 0.25 milliliters per minute.Connect the inline solution heater and set it to 37 degrees Celsius. Connect the objective heater and set it to 37 degrees Celsius. Apply oil to the objective and insert the profusion chamber into the stage mount.Apply oil to the chamber heater pad and attach it to the perfusion chamber, then connect it and set it to 37 degrees Celsius. Lock the profusion chamber into the stage adapter and bring the objective oil into contact with the coverslip on the underside of the chamber. Use Brightfield illumination to focus on the layer of axons on the bottom surface of the nerve closest to the coverslip surface.Myelinated axons can be identified by the presence of a myelin sheath which is visible under Brightfield transmitted light illumination without contrast enhancement. Acquire a Brightfield reference image, recording the directionality of the nerve, then acquire a confocal image using a 488 nanometer laser and an emission filter appropriate for photoactivatable GFP. Set the laser power to approximately five times normal imaging power and acquire an image with an exposure time of three to four minutes, then acquire another confocal image to record the pre-activation autofluorescence after the bleaching step.Draw a line parallel to the axons with a length equal to the desired activation window size of the Brightfield image. Using this line as a guide, draw a rectangular region of interest across the field of view perpendicular to the axons. Activate the GFP fluorescence in this region by pattern excitation with the 405 nanometer light, making sure that an image is acquired just prior to and immediately following activation.As the activation finishes, start a one-minute timer. When it goes off, start acquisition of a timelapse series. Add 250 milligrams of fluorescein powder to 0.5 milliliters of double distilled water and mix it until there are no visible particles, then spin the solution for 30 seconds in a tabletop centrifuge to sediment any undissolved material.Add eight microliters of fluorescein solution to a slide and apply a number 1.5 coverslip. Blot the excess liquid, seal the coverslip with nail polish, and allow the slide to dry. Place the slide coverslip side down on the inverted microscope stage and adjust the focus on the thin plane of fluorescence at the surface of the coverslip.Move the slide around to find a field of view that does not contain air bubbles or large fluorescein particles and acquire a Z-stack spanning six micrometers at 0.2 micrometer intervals such that the middle image is the original focus plane. Close all light path shutters including the camera shutter and set the laser power and the exposure time to zero seconds. Acquire a stack of 100 images with these settings, which will be averaged to generate the dark field image.Representative images from pulse-escape and pulse-spread experiments are shown here. For the pulse-escape method, the fluorescence decay kinetics in the activated region can yield information on long-term and short-term pausing behavior. The activated region can be short for these experiments.A longer activated region is used for the pulse-spread method 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. 15 micrometer flanking windows are used here.The total fluorescence from the measuring window was quantified. The decay is biphasic for the pulse-escape method with an initial exponential decay representing the departure of on-track neurofilaments and a second slower exponential decay at 10 to 20 minutes that represents the departure of off-track filaments. 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 five minutes. A timelapse from a glycolytically inhibited nerve is shown here, demonstrating an apparent reduction in transport out of the activated region. Indeed, the distal and proximal slopes are significantly lower in nerves treated with inhibitor.There is also a significant decrease in the population velocity between these two conditions. When attempting this protocol, remember that it is crucial to be hasty in conducting the dissection and chamber assembly portions as nerve health declines following sacrifice of the animal. Neuronal cell culture can allow measurement and analysis of the movement of individual neurofilaments, which would supplement the population scale data gathered using this protocol.

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Research

JoVE Journal

Neuroscience

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 ...

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 ...

In vivo Calcium Imaging in Mouse Inferior Olive

Inferior olive (IO), a nucleus in the ventral medulla, is the only source of climbing fibers that form one of the two input pathways entering the cerebellum. IO has long been proposed to be crucial for motor control and its activity is currently considered to be at the center of many hypotheses of both motor and cognitive functions of the cerebellum. While its physiology and function have been relatively well studied on single-cell level in vitro, presently there are no reports on the organization of the IO network activity in living animals. This is largely due to the extremely challenging anatomical location of the IO, making it difficult to subject to conventional fluorescent imaging methods, where an optic path must be created through the entire brain located dorsally to the region of interest.
Here we describe an alternative method for obtaining state-of-the-art -level calcium imaging data from the IO network. The method takes advantage of the extreme ventral location of the IO and involves a surgical procedure for inserting a gradient-refractive index (GRIN) lens through the neck viscera to come into contact with the ventral surface of the calcium sensor GCaMP6s-expressing IO in anesthetized mice. A representative calcium imaging recording is shown to demonstrate the feasibility to record IO neuron activity after the surgery. While this is a non-survival surgery and the recordings must be conducted under anesthesia, it avoids damage to life-critical brainstem nuclei and allows conducting large variety of experiments investigating spatiotemporal activity patterns and input integration in the IO. This procedure with modifications could be used for recordings in other, adjacent regions of the ventral brainstem.

In vivo Calcium Imaging in Mouse Inferior Olive

We present a protocol to expose the brainstem of adult mouse from the ventral side. By using a gradient-refractive index lens with a miniature microscope, calcium imaging can be used to examine the activity of inferior olive neural somata in vivo.