We would like to thank Robert M. Brownstone (Queen Square Institute of Neurology, University College London) for sharing the ChAT-eGFP, ChAT.Cre and Rosa26.tdTomato mice, and Pietro Fratta (Queen Square Institute of Neurology, University College London) for sharing the HB9.GFP mouse. We would like to thank Elena R.&#160;Rhymes, Charlotte J.P. Kremers, and Qiuhan Lang (Queen Square Institute of Neurology, University College London) for critical reading of the manuscript. This work was supported by a Junior Non-Clinical Fellowship from the Motor Neuron Disease Association (UK) (Tosolini/Oct20/973-799) (APT), the Wellcome Trust Senior Investigator Awards (107116/Z/15/Z and 223022/Z/21/Z) (GS), a UK Dementia Research Institute Foundation award (GS); and a Medical Research Council Career Development Award (MR/S006990/1) (JNS).

All mouse handling and experiments were performed in accordance with the Animals (Scientific Procedures) Act (1986) and were approved by the University College London - Queen Square Institute of Neurology Ethics Committee.
1. Animals
<ol>
	<li>House all animals in individually ventilated cages in a temperature- and humidity-controlled environment and maintain them on a 12 h light/dark cycle with ad libitum access to food and water.</li>
	<li>Use both male and female mice of the following transgenic strains: 1) heterozygous Tg(Chat-EGFP) GH293Gsat/Mmucd mice, referred to as ChAT.eGFP mice; 2) heterozygous B6.Cg-Tg(Hlxb9-GFP)1Tmj/J, referred to as HB9.GFP mice; and 3) heterozygous B6.Cg-Tg(Thy1-CFP/COX8A)S2Lich/J, referred to as Mito.CFP mice.</li>
	<li>Generate ChAT.tdTomato mice by crossing homozygous B6;129S6-Chattm2(cre)Lowl/J, referred to as ChAT.Cre mice, with homozygous B6.Cg-Gt(ROSA)26Sortm9(CAG-tdTomato)Hze/J, referred to as Rosa26.tdTomato mice.</li>
	<li>Generate ChAT.tdTomato::Mito.CFP mice by crossing heterozygous ChAT.tdTomato mice with heterozygous Mito.CFP mice.</li>
</ol>
2. Intramuscular injections of fluorescent HCT
<ol>
	<li>Pre-surgery preparation
	<ol>
		<li>Express HCT (HCT441, residues 875-1315) fused to an improved cysteine-rich tag in bacteria as a glutathione-S-transferase fusion protein as per 45. Label HCT with AlexaFlour555 C2 maleimide31, dialyze it in ice-cold dialysis buffer (10 mM HEPES-NaOH, 100 mM NaCl, pH 7.4), freeze it in liquid nitrogen, and store it at -80 &#176;C. Before performing in vivo experiments, first test HCT in vitro for successful uptake and transport in primary neurons.</li>
		<li>Dilute fluorescent HCT (e.g., HCT-555) to a final and experimentally consistent concentration ranging from 2.5 to 10 &#181;g/&#181;L in sterile phosphate-buffered saline (PBS) in a 0.2 mL tube. At this step, add more compounds/factors to the HCT solution if required (e.g., brain-derived neurotrophic factor). 
		NOTE: The final volume must be appropriate for the size of the muscle(s) of interest. For example, prepare an injection volume of 3-4 &#181;L for the tibialis anterior (TA) muscle and ~1 &#181;L for the smaller soleus muscle. Keep the working concentration of HCT between 2.5 and 10 &#181;g/&#181;L regardless of the final volume.</li>
		<li>Mix the HCT solution using a pipette or vortex, and briefly spin down at low speed using a desktop centrifuge to collect the liquid and remove large bubbles. Protect the HCT from light and transport on ice.</li>
		<li>Use a pulled glass micropipette for optimal intramuscular injections into smaller muscles (e.g., soleus) or for intrasciatic nerve injections. Pull graded glass micropipettes (as per 46) before surgery. 
		&#8203;NOTE: To enable pipetting and restrict flow up and out the back of the micropipette, carefully break off a small piece from the sharp tip using fine forceps under a dissecting microscope. Take care to dispose of the broken end in the appropriate bin.</li>
		<li>Sterilize and clean all surgical tools prior to use.</li>
	</ol>
	</li>
	<li>Surgery-intramuscular injections
	<ol>
		<li>Prepare for surgery by securing a sterile surgical drape on a heat mat set to 37&#176;C. Position and focus the operating microscope. For surgery to commence, unpack onto the surgical drape the presterilized surgical tools, surgical tape, sterile cotton swabs, 70% (v/v) ethanol in water, sterile saline, sutures, and a Hamilton needle or pulled glass micropipettes.</li>
		<li>Ensure the anesthetic machine has sufficient oxygen and isoflurane for the duration of the surgical procedure. Direct the flow of anesthesia to the induction chamber and switch on the anesthetic machine.
		<ol>
			<li>To begin, use an oxygen flow rate of 1-2 L/min and 5% isoflurane. Place the mouse in the induction chamber to initiate anesthesia. When the righting reflex is absent, reduce anesthesia to 2-3% isoflurane, direct the flow of anesthesia to the mouthpiece, and transfer the mouse to the mouthpiece located in a separate area of the surgical space.</li>
		</ol>
		</li>
		<li>Ensure both the corneal and pedal withdrawal reflexes are absent before shaving the area of fur covering the muscle(s) to be injected. When complete, remove as much shaved fur from the mouse as possible using the sticky side of surgical tape, and place the mouse on weighing scales to record its pre-surgical weight.</li>
		<li>Carefully apply eye lubricant using a cotton swab and transfer the mouse and mouthpiece to the surgical area. 
		NOTE: Try to limit the amount of shaved fur that is also transferred to the surgical area. Use surgical tape to secure the head to the mouthpiece to prevent the mouse from slipping out. Using a separate cotton swab, apply ethanol to the shaved region to sterilize and reduce fur contamination.</li>
		<li>Position the body according to the muscle to be injected. For example, for the TA, place the mouse on its back and stretch out the hindlimb at ~10&#176; from the midline. Alternatively, for soleus injections, place the animal on its side and extend the hindlimb at ~45&#176; from the midline. When the hindlimb is in the correct position, use surgical tape across the foot to prevent unwanted movement during surgery. 
		&#8203;NOTE: The injection procedures for TA, gastrocnemius, and soleus muscles have been previously detailed32.</li>
		<li>Before making an incision, confirm that the anesthesia is sufficient by testing the pedal withdrawal reflex. Monitor the anesthesia continually and maintain it throughout the surgical procedure with regular assessment of breathing and the withdrawal reflex.</li>
		<li>At this point, draw the working HCT solution into the Hamilton syringe or pulled glass micropipette.</li>
		<li>Make a small incision over the muscle(s) of interest in the area(s) that correspond(s) with the motor end plate regions46,47,48. Pierce the external fascia on the muscle and slowly inject the HCT as per 32. Leave the syringe/micropipette in position for 5-10 s before slowly withdrawing.</li>
		<li>Close the incisions with 1-2 sutures and transfer the mouse to an isolated recovery cage. Monitor the mouse post surgery for a minimum of 30 min, before returning it to the home cage. When the mouse has successfully recovered and post surgical monitoring is complete, return the cage to normal housing conditions.</li>
	</ol>
	</li>
</ol>
3. In vivo axonal transport
<ol>
	<li>Exposing the sciatic nerve
	<ol>
		<li>Set the microscope environmental chamber to 37 &#176;C at least 1 h prior to imaging.</li>
		<li>Prepare to expose the sciatic nerve by arranging the surgical drape, tools, tape, sterile cotton swabs, 70% ethanol, and sterile saline around the surgical area. Ensure the anesthetic machine has sufficient stores of oxygen and isoflurane for up to 2 h per mouse. Create a wedge out of parafilm or invisible tape by cutting it into a narrow rectangle (e.g., ~1 cm width for larger mice) with an angled tip and place it underneath the exposed sciatic nerve to aid the imaging process. Place the induction chamber on top of a heat mat and set it to body temperature. 
		NOTE: Four hours is ample time for HCT to have been taken up and retrogradely transported from the site of injection to the sciatic nerve; hence, a single mouse can be readied for re-anesthesia after this time.</li>
		<li>Direct the flow of anesthesia into the induction chamber, switch on the anesthetic machine with an oxygen flow rate of 1-2 L/min and 3-4% isoflurane, and place the mouse in the induction chamber to initiate anesthesia. 
		NOTE: As the in vivo axonal transport experiment is a terminal procedure, there is no need to lubricate the eyes.</li>
		<li>When the righting reflex is absent, reduce anesthesia to 2-3% isoflurane, direct the flow of anesthesia to the mouthpiece, and transfer the mouse to the mouthpiece. Use surgical tape to secure the head to the mouthpiece, extend the targeted hindlimb at ~45&#176; from the midline, and use surgical tape over the foot to maintain this position. 
		NOTE: Reduced anesthesia is advantageous at this point as it can limit the impact of breathing artifacts during the imaging process.</li>
		<li>Ensure corneal and pedal withdrawal reflexes are absent, and then use scissors to cut away the skin overlying the sciatic nerve32 (i.e., a large area extending from the central spinal cord to mid-lower hindlimb). Remove the overlying biceps femoris muscle, as well as any other musculature and connective tissue that is near the sciatic nerve. Avoid damaging the sciatic nerve and surrounding blood vessels, especially those located near the lateral aspect of the patella/proximal aspect of the lateral gastrocnemius head.</li>
		<li>When the intact sciatic nerve is sufficiently exposed, apply prewarmed sterile saline to the area around the sciatic nerve to prevent desiccation. Use curved forceps to disrupt the deep-lying connective tissue and place the pre-prepared parafilm &#39;wedge&#39; underneath the nerve. When complete, place saline-soaked cotton wool on the exposed area and move the mouse into the induction chamber positioned on top of the heat mat (set to 37 &#176;C), which should still be filled with isoflurane in O2.</li>
	</ol>
	</li>
	<li>In vivo axonal imaging
	<ol>
		<li>Place a 22 x 64 mm coverglass on the customized microscope stage and secure its position with tape. Select and apply immersion oil to the objective, and then connect the microscope stage to the inverted microscope. Slowly raise the oil-immersed objective until contact is made between the oil and coverglass. 
		NOTE: Either the 40x, 1.3 numerical aperture (NA) DIC Plan-Apochromat or 63x, 1.4 NA DIC Plan-Apochromat oil-immersion objectives can be used to image in vivo transport in the sciatic nerve.</li>
		<li>Move the anesthetic mouthpiece onto the microscope stage and secure the anesthesia hoses with tape to prevent disturbance to the anesthesia. Remove the cotton wool from the sciatic nerve and transfer the mouse from the induction chamber to the mouthpiece, with the exposed nerve facing the coverglass. Use surgical tape to ensure the mouse&#39;s head is fixed to the mouthpiece and maintain the lowest, effective level of anesthesia. Gently lift the mouse by its tail and add sterile saline to the coverslip near the exposed sciatic nerve to restrict desiccation and aid imaging. 
		NOTE: Close all doors of the environmental chamber to ensure the area remains at body temperature.</li>
		<li>Using the oculars, locate the sciatic nerve, determine the optimal focal point, and select an area of interest containing motile axonal organelles. 
		NOTE: A detailed explanation of this process has been previously described32.</li>
		<li>Switch to the computer software by clicking the Acquisition button (or equivalent), and select an area of interest. Use a digital zoom to obtain a total of &#62;80x magnification and rotate the selected area to horizontally visualize the axons (e.g., right-to-left moving retrograde cargo and left-to-right moving anterograde cargo). 
		NOTE: The directionality parameters are user-dependent, but must remain consistent throughout experiments.</li>
		<li>Optimize the signal intensity by adjusting parameters such as laser intensity (0.2 - 1%), pinhole aperture (1 AU - max), gain (Master) (700 - 1000), digital offset (-50 - 0), and digital gain (1.0 - 4.0). To reduce the potential influence of phototoxicity, maintain laser intensity at &#8804; 1% where possible, with a maximum laser intensity of 2%. Change all other parameters before adjusting the laser intensity for optimal signal detection.</li>
		<li>Click the Regions box (or equivalent), select a rectangular region of interest, then in the Acquisition Mode (or equivalent), set the frame size to a minimum 1024 x 1024 pixels, and commence time-lapse acquisition of 100-1,000 frames. 
		NOTE: The desired frame acquisition rate is user-dependent (e.g., transport can be assessed with frame rates between 0.1 and 6 s) and can be adjusted with software parameters, such as region of interest, scan speed time, acquisition averaging, and laser directionality. For example, to obtain a slower frame rate, increase the height/width of the region of interest, acquire slower scan speeds, increase the acquisition averaging, and use single laser directionality, and vice versa for a faster frame rate. The frame acquisition rate must remain consistent across comparable datasets because imaging at different frequencies may cause inconsistencies.&#160;Fast cargoes such as signaling endosomes require a faster frame rate (e.g. 0.1-3 s) compared with slower organelles like mitochondria which can be analysed using a slower rate (e.g. 2.5-6 s).</li>
		<li>Aim to capture a minimum of 10 motile cargoes from a minimum of three axons per mouse. 
		NOTE: Based on two-sample, two-sided power calculations (with standard power of 0.8 (1&#8722;&#946;) and type I error rate of 5% (&#945;)), sample sizes of 6-8 are sufficient to identify axonal transport differences between wild-type and disease models35,43.</li>
		<li>Once imaging is complete, euthanize&#160;the mouse immediately while under anesthesia (e.g., cervical dislocation). Post mortem tissue, such as muscles and sciatic nerves, can also be harvested for further analysis.</li>
	</ol>
	</li>
</ol>

The authors have no conflicts of interest.

This protocol details steps to assess in vivo axonal transport of signaling endosomes and mitochondria in intact axons of the mouse sciatic nerve. Indeed, an experimental setup is provided that enables users to 1) distinguish motor from sensory neurons in vivo, in situ, and ex vivo by using mice expressing fluorescent reporter proteins selectively expressed in motor neurons; 2) assess in vivo axonal transport of signaling endosomes specifically in motor neuron axons using three different transgenic mice; 3) investigate in vivo axonal transport of mitochondria specifically in motor neuron axons; and 4) concurrently assess in vivo transport dynamics of signaling endosomes and mitochondria within the same axon. This approach has vast potential for investigating axonal transport in basal conditions and can be used to assess pathological perturbations in different diseases affecting peripheral motor and sensory nerves.
Using previous experimental paradigms as a foundation31,32, here we have detailed novel, robust ways to differentiate axonal transport occurring in motor versus sensory neurons using transgenic reporter mice. Using the Mito.CFP mouse, this approach has been further developed to assess in vivo mitochondrial transport&#160;by avoiding intrasciatic nerve injections of TMRE36. This circumvents possible neural damage and perturbations in axonal transport caused by the intranerve injection of the probe. Furthermore, this protocol allows the visualization of axonal transport of multiple organelles in motor axons innervating muscles with distinct physiological properties (e.g., fast-twitch fatigable muscles vs. slow-twitch fatigue-resistant muscles). As such, signaling endosome and/or mitochondrial axonal transport dynamics can be assessed in different subsets of &#945;-motor neurons44. Moreover, the axonal transport of those organelles in pathological settings can also be assessed through crossbreeding with mouse models of different neurodegenerative diseases1,2,3.
The axonal transport toolkit is continuously expanding28,29, and ex vivo protocols have been developed to assess transport dynamics using cultured mouse ventral horn explants49 or excised mouse nerve-muscle preparations50. Furthermore, the development of protocols to assess axonal transport in induced human pluripotent stem cell (hiPSC)-derived cortical51 neurons or hiPSC-derived spinal motor neurons52 has enabled investigation of human neurons with disease-causing mutations. Such cutting-edge protocols in mouse tissue and human cells can provide critical insights into neuronal function, facilitate novel pathomechanistic discovery in neurodegenerative disease models, and be used to test therapeutic molecules and strategies.
Several critical steps need to be followed for the successful implementation of these techniques, and some important notes have been provided in the protocol section. The major requirements for intravital imaging are the inverted confocal microscope with customized stage insert and the equipment to maintain anesthesia and optimum temperature. Indeed, a specialized mobile anesthetic system is needed for 1) induction of anesthesia, 2) dissection/tissue processing (i.e., exposing the sciatic nerve), and 3) maintaining anesthesia during intravital imaging (as previously detailed in 31,32). Especially when using higher magnification objectives (e.g., 40x or 63x), the depth of anesthesia can impact the image quality, as deeper anesthesia induces large &#39;gaspy&#39; breaths that result in frequent shifts in focus. Such large movements will undoubtedly impact post imaging transport analyses (e.g., tracking cargoes using the Fiji plugins TrackMate53 or KymoAnalyzer54) as the breathing movements produce artifacts in time-lapse videos that can render them unsuitable&#160;for automated tracking or require more time-consuming assessment. Moreover, we have also observed imaging artifacts caused by pulsating arteries within the sciatic nerve, which can only be resolved by choosing a different imaging region. The microscope must be equipped with an environmental chamber capable of maintaining constant body temperature, as temperature and pH influence axonal transport55. Furthermore, the application of analgesics post surgery should be avoided, as they can alter transport dynamics56. If the experimental design is longitudinal and requires repeated imaging (e.g.,57), the dissection protocols need to be appropriately adjusted to be minimally invasive and may require additional ethical/licence approval.
Certain experimental considerations need to be kept in mind. First, most of the protocols detailed herein involve the use of transgenic mice that possess fluorescent reporter proteins in mitochondria or motor neuron axons. Each of these mouse lines should be bred and imaged as hemi-/heterozygote. The exceptions, however, are the ChAT.Cre and Rosa26.tdTomato mouse lines that can be separately maintained as homozygotes, with the resulting hemizygote offspring enabling tdTomato expression in cholinergic neurons after Cre-loxP recombination. When cross-breeding transgenic hemi-/heterozygote mice (e.g., Mito.CFP) with other transgenic hemi-/heterozygote mice (e.g., ChAT.eGFP), one needs to carefully consider the breeding strategy, as obtaining the desired numbers of double-mutant progeny can be time-consuming. Moreover, when breeding the F1 generation of ChAT.Cre and Rosa26.tdTomato mice (i.e., ChAT.tdTomato) with additional transgenic strains (e.g., Mito.CFP), one should expect even fewer mice carrying the desired triple transgenes. In addition, one must also consider the potential fluorophore overlap when breeding two-reporter mice with nearby wavelength properties (e.g., Mito-CFP-excitation: 435 nm, emission: 485 nm, bred with ChAT.eGFP-excitation: 488 nm, emission: 510 nm), although it may be possible to overcome this problem with spectral unmixing58.
This technique has some limitations to be considered. In this work and our previous protocols31,32, we have shown how several genetically encoded markers and different staining methods can be used to label and track distinct organelles in vivo. However, not all probes are suitable for this experimental approach. We assessed injections into either TA or soleus muscle of cholera toxin beta subunit (CTB)-488 (0.5-1.5 &#181;g/&#181;L ~4 h before imaging), a probe routinely used to label motor neuron cell bodies in in vivo retrograde tracer experiments59,60. However, when injected alone or co-injected with HCT-555, the CTB-488 labeling was poor despite using concentrations similar to those used for successful retrograde motor neuron tracing. Thus, we conclude that, despite CTB being an excellent in vitro marker of signaling endosomes in neuronal cultures61, HCT remains the gold-standard probe to identify signaling endosomes in vivo in sciatic nerve axons.
Using different routes, we also tested probes routinely used for labeling lysosomes, such as LysoTracker green DND-26, and markers of active lysosomal hydrolases, such as BODIPY-FL-pepstatin A for Cathepsin D62 and Magic Red for Cathepsin B, but with no success. We tried intramuscular delivery of BODIPY-FL-pepstatin A (2.5 &#181;g into the TA ~4 h before imaging), as well as intrasciatic&#160;nerve injection of 2 &#181;L of LysoTracker (10 &#181;M), BODIPY-FL-pepstatin A (10 &#181;M) or Magic Red (1/10) 30-60 min before imaging. Despite these probes highlighting the nerve, we were unable to find clearly labeled organelles. The probes accumulated around axons, likely being retained by Schwann cells. Hence, the unsuccessful labeling of lysosomes may be due to deficient probe delivery into neurons, although the existence of more suitable concentrations cannot be ruled out. Given that TMRE labeling works under similar conditions (i.e., intrasciatic&#160;nerve injections), the labeling intensity may be dye-dependent and must be tested for each marker independently. However, we conclude that targeting lysosomes in vivo with these probes is not feasible at the concentrations stated above.
Methods of anesthesia can alter distinct physiological readouts (e.g., cochlea function63 and cortical electrophysiology64); however, whether anesthesia influences in vivo axonal transport in the sciatic nerve is currently unknown. Given the reduced neuromuscular activity under isoflurane-induced anesthesia, it is possible that transport kinetics differ compared to the wakeful state. However, the only in vivo study that has directly investigated this revealed that transport of dense core vesicles in thalamocortical projections does not differ between anesthetized and awake mice65. Furthermore, because distinctions in transport between wild-type and disease model mice are detectable under anesthesia35,43, it is clear that isoflurane exposure does not prevent the identification of perturbances in signaling endosome or mitochondrial trafficking.
This protocol has other potential applications, which have been described below. Breeding of the transgenic mice outlined in this protocol (e.g., Mito.CFP, ChAT.eGFP) with neurodegenerative disease mouse models1,2,3 will enable neuron subtype- and/or cargo-specific investigations. Moreover, recently developed mouse Cre lines66 would also permit the visualization of fluorescent reporter proteins in distinct sensory axon populations. For example, Rosa26.tdTomato mice can be crossed with a neuropeptide Y receptor-2-expressing (Npy2r).Cre mouse to enable tdTomato fluorescence in myelinated A-fiber nociceptors67. Furthermore, temporal control can also be achieved by using inducible Cre systems (e.g., tamoxifen)68. Another potential application relies on the availability of transgenic mice expressing fluorescent reporter proteins in Schwann cells. Indeed, S100-GFP69 and PLP-GFP70 mice enable in vivo and/or in situ imaging of Schwann cells and have been at the forefront of research involved in Schwann cell migration during peripheral nerve regeneration.
In addition to these applications and complementing the Mito.CFP mouse is the availability of several transgenic mouse lines that express fluorescent proteins in distinct organelles, such as mitochondria and autophagosomes. For example, investigating in vivo mitochondrial transport might be possible with the mito::mKate2 mouse71 or the photoconvertible mitoDendra mouse57. Moreover, in vivo mitophagosome transport may be possible using the pH-sensitive mito-Keima mouse72 and the mito-QC mouse73 for mitophagy analyses. Furthermore, the lysosomal labeling difficulties we encountered may be overcome by using mice expressing LAMP1-GFP, with the caveat that LAMP1 is also present in endocytic organelles distinct from lysosomes74.
In summary, we have provided novel ways to assess in vivo axonal transport of several organelles in specific peripheral nerve axons from diverse transgenic mice. The concurrent imaging of different organelles will be particularly important, given recent findings of axonal interactions and co-trafficking of organelles such as mitochondria and endosomes75,76. We believe that the presented methods will be useful for improving understanding of the basal physiology of axons in vivo and untangling important pathomechanisms driving neurodegeneration of peripheral nerves.

The peripheral nervous system (PNS) connects the central nervous system (CNS) to its distal targets, permitting the relay of efferent signals to exert motor control and afferent signals to provide sensory feedback. Using the multitude of advances in mouse genetics, scientists have developed different mouse models to investigate many diseases/syndromes afflicting the PNS1,2,3. As most neurodegenerative pathologies are multifactorial with cell autonomous and non-autonomous contributions4,5, untangling cell-/neuron-specific pathologies can provide crucial, novel insights into disease mechanisms.
To this end, the development of bacterial artificial chromosome (BAC)-transgenic mice6 has enabled selective endogenous expression of fluorescent proteins in targeted subsets of neurons. For example, BAC-transgenic mice are available, which express green fluorescent protein (GFP) in cholinergic7 or glycinergic neurons8, or a variant red fluorescent protein (tdTomato) in parvalbumin-positive neurons9. Alternatively, selective neuronal expression of fluorescent proteins can be achieved via Cre-loxP technology10. For instance, mouse strains expressing Cre-recombinase in subsets of neurons (e.g., choline acetyltransferase (ChAT)-Cre) can be bred with mice that express a fluorescent protein (e.g., tdTomato or GFP) from a constitutive locus (e.g., Gt(ROSA)26Sor) under the control of a transcriptional repressor flanked by loxP sites11 (e.g., generating mice that express tdTomato only in cholinergic neurons). Indeed, using Cre-loxP recombination, transgenic mice have been generated that express yellow fluorescent protein in axons of the descending corticospinal tract12.
In addition, recent advances in CRISPR/Cas9 gene editing, such as ORANGE, enable the fluorescent tagging of multiple endogenous neuronal proteins, with expression achievable at nanoscale resolution13. Moreover, in combination with Cre-expressing mouse strains, ORANGE-CAKE can be used to tag multiple endogenous proteins in individual neurons13. Alternatively, viral-mediated neuronal tracing also allows the labeling of neuronal subsets and can be achieved with targeted combinations of viral serotypes and/or cell-specific promoters14,15,16,17.
Further to the neuronal labeling methods, mouse lines have also been engineered to express reporter proteins targeting specific organelles, such as mitochondria expressing cyan fluorescent protein (Mito.CFP)18 or autophagosomes expressing GFP (LC3.GFP)19. Moreover, mouse lines have been engineered to assess calcium dynamics specifically in neurons (e.g., Thy1.GCaMP)20,21. Altogether, with the advancement of such models, novel experimental applications enable scientists to ask more precise biological and pathological questions about the CNS and PNS.
The main role of peripheral motor nerves is to transmit electrical signals to skeletal muscle to elicit movement. In addition, and occurring over longer time-scales, neurochemical and physiological messages in the form of diverse organelles (e.g., mitochondria, endolysosomes, signaling endosomes) traverse the cytoskeletal network in a uni- or bi-directional manner to help maintain neuronal homeostasis22,23,24. Impairments in axonal transport have disastrous consequences for neuronal health and are linked to many neurodevelopmental and neurodegenerative diseases25. At the molecular level, impairments in axonal transport can disrupt physiological events regulating synaptic signaling and plasticity, gene transcription, and local translation throughout the axon26,27. While there is a multitude of tools to study these events in cultured cells/neurons28,29, assessing&#160;axonal transport dynamics and axonal-linked biological events in vivo are required to confirm key insights into physiological and pathological processes30.
Over the years, the Schiavo Laboratory has optimized protocols to ask diverse questions about axonal transport31,32,33,34,35,36. These experiments have expanded from the discovery that a fluorescently labeled atoxic fragment of tetanus neurotoxin (HCT) is internalized into axon terminals in skeletal muscle through interactions with nidogens and polysialogangliosides37. Once internalized, HCT is retrogradely transported in Rab7-positive, neurotrophin-containing signaling endosomes that are destined for the cell bodies of motor and sensory neurons38,39,40,41. In parallel, advances in imaging technology have enabled the real-time analysis of peripheral nerve bundles and individual axons in live, anesthetized mice30. The first foray into assessing in vivo axonal transport dynamics in pathology revealed presymptomatic impairments in the transport of signaling endosomes and mitochondria in the SOD1G93A mouse model of amyotrophic lateral sclerosis (ALS)35. Importantly, these defects are unlikely to simply represent secondary consequences of neurodegeneration, given the finding that motor neuron loss can occur in the absence of axonal transport perturbations in a mouse model of Kennedy&#39;s disease42 and a heterozygous mutant FUS model of ALS43. Such axonal transport deficits can be remedied in ALS mice using inhibitors of specific kinases33 or growth factor receptors34. Moreover, treating neurons with a specific histone deacetylase blocker alters mitochondrial transport in vivo36. Most recently, we report that the BDNF-dependent modulation of axonal transport is dysregulated in distinct motor neuron subtypes in ALS mice44.
By using an ever-expanding toolkit for assessing axonal transport dynamics28,29, this video protocol outlines several applications that will permit further insights into different biological and pathological scenarios. First, transgenic mice that selectively express fluorescent proteins in cholinergic neurons (i.e., motor neurons) are used to discriminate between motor and sensory axons both in vivo and ex vivo. Fluorescently labeled HCT is then loaded into signaling endosomes in three transgenic lines to differentiate axonal transport dynamics in distinct peripheral neurons. The next experimental protocol details a multiplex fluorescence approach to assess mitochondrial transport specifically in motor neurons by breeding ChAT.tdTomato mice with Mito-CFP mice. Finally, instructions are provided on how to concurrently image mitochondria and signaling endosomes within the same axon in vivo.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.2 mL PCR tube</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>70% (v/v) ethanol in distilled water</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>AlexaFlour555 C2 maleimide</td>
			<td>ThermoFisher Scientific</td>
			<td>A-20346</td>
			<td>Can also use AlexaFlour-488 or -647 Maleimide</td>
		</tr>
		<tr>
			<td>B6.Cg-Gt(ROSA)26Sortm9(CAG-tdTomato)Hze/J</td>
			<td>Jackson Laboratory</td>
			<td>7909</td>
			<td>Rosa26.tdTomato mice</td>
		</tr>
		<tr>
			<td>B6.Cg-Tg(Hlxb9-GFP)1Tmj/J mice</td>
			<td>Jackson Laboratory</td>
			<td>5029</td>
			<td>HB9.GFP mice</td>
		</tr>
		<tr>
			<td>B6.Cg-Tg(Thy1-CFP/COX8A)S2Lich/J mice</td>
			<td>Jackson Laboratory</td>
			<td>7967</td>
			<td>Mito.CFP mice</td>
		</tr>
		<tr>
			<td>B6;129S6-Chattm2(cre)Lowl/J mice</td>
			<td>Jackson Laboratory</td>
			<td>6410</td>
			<td>ChAT.Cre mice</td>
		</tr>
		<tr>
			<td>Computer with microscope control and image acquisition software</td>
			<td>Zeiss</td>
			<td>Zen</td>
			<td></td>
		</tr>
		<tr>
			<td>Cotton swab</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Desktop centrifuge</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Dissecting microscope</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Eye lubricant</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fine curved forceps</td>
			<td>Dumont</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Fine straight forceps</td>
			<td>Dumont</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glass coverslip (22 x 64 mm, thickness no. 1)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Graduated, glass micropipette with microliter markings and plunger</td>
			<td>Drummond Scientific</td>
			<td>5-000-1001-X10</td>
			<td></td>
		</tr>
		<tr>
			<td>Hair clippers</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hamilton microliter syringe (701 N, volume 10 &#956;L, needle size 26 s G, bevel tip, needle L 51 mm)</td>
			<td>Merck</td>
			<td>20779</td>
			<td></td>
		</tr>
		<tr>
			<td>HcT-441</td>
			<td>N/A</td>
			<td>N/A</td>
			<td>See Restani et al., 2012 for more details</td>
		</tr>
		<tr>
			<td>Heating pad</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Immersion oil for fluorescent imaging at 37 &#176;C</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Inverted confocal microscope with environmental chamber</td>
			<td>Zeiss</td>
			<td>LSM 780</td>
			<td>Most inverted confocals should be adaptable</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane vaporizer/anesthesia machine with induction cham-ber and mask stabilizer</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Magic tape</td>
			<td></td>
			<td></td>
			<td>invisible tape</td>
		</tr>
		<tr>
			<td>Micropipette puller</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>Parafilm</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate-buffered saline (PBS): 137 mM NaCl, 10 mM Na2HPO4, 2.7 mM KCl, 1.8 mM KH2PO4&#8211;HCl, pH 7.4</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Recombinant human brain-derived neurotrophic factor (BDNF)</td>
			<td>Peprotech</td>
			<td>450-02</td>
			<td>BDNF that can be used to co-inject with HcT-555</td>
		</tr>
		<tr>
			<td>Saline</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Scalpel blade</td>
			<td>Dumont</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Small spring scissors</td>
			<td>Dumont</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Surgery/operating microscope</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical drape</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical suture</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical tape</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tg(Chat-EGFP) GH293Gsat/Mmucd mice</td>
			<td>MMRRC</td>
			<td>000296-UCD</td>
			<td>ChAT.eGFP</td>
		</tr>
		<tr>
			<td>Vortex mixer</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

expanding the toolkit for in vivo imaging of axonal transport

Axonal transport maintains neuronal homeostasis by enabling the bidirectional trafficking of diverse organelles and cargoes. Disruptions in axonal transport have devastating consequences for individual neurons and their networks, and contribute to a plethora of neurological disorders. As many of these conditions involve both cell autonomous and non-autonomous mechanisms, and often display a spectrum of pathology across neuronal subtypes, methods to accurately identify and analyze neuronal subsets are imperative.
This paper details protocols to assess in vivo axonal transport of signaling endosomes and mitochondria in sciatic nerves of anesthetized mice. Stepwise instructions are provided to 1) distinguish motor from sensory neurons in vivo, in situ, and ex vivo by using mice that selectively express fluorescent proteins within cholinergic motor neurons; and 2) separately or concurrently assess in vivo axonal transport of signaling endosomes and mitochondria. These complementary intravital approaches facilitate the simultaneous imaging of different cargoes in distinct peripheral nerve axons to quantitatively monitor axonal transport in health and disease.

This paper details a versatile protocol that expands the in vivo axonal transport toolkit in rodent models. Figure 1 demonstrates that motor neuron axons can be differentiated from both sensory neuron axons and Schwann cells by using transgenic mice. Figure 1A depicts eGFP expression in cholinergic motor axons from a live, anesthetized ChAT.eGFP mouse. Figure 1B uses an alternative method to attain tdTomato expression in a freshly excised nerve (i.e., no additional tissue processing) from a ChAT.tdTomato mouse. Hence, using transgenic strains such as ChAT.eGFP, ChAT.tdTomato or Hb9.GFP enables motor axon-specific labeling in vivo.
Alternatively, axons can also be identified by injecting tracers/markers (e.g., HCT31,32 or viruses encoding eGFP15) into skeletal muscles. Figure 2 highlights such an application, depicting eight robustly expressing ChAT.eGFP-positive axons that contain HCT-555-positive signaling endosomes (white arrows), ~4 h post probe injection into the TA muscle. Using this experimental design, we could identify TA-innervating &#945;-motor neurons, which are predominantly fast-fatigable44. A further five ChAT.eGFP axons with less robust eGFP expression (Figure 2A,&#160;orange asterisks) were partially out of focus and likely to be located slightly deeper within the sciatic nerve.
Moreover, we identified HCT-555-positive signaling endosomes in eGFP-negative sensory axons (yellow arrows). As such, using this experimental paradigm, one can specifically assess and compare the axonal transport of signaling endosomes in motor versus sensory neurons in vivo. Indeed, using this transgenic reporter strain,&#160;we discovered that transport of signaling endosomes in ChAT.eGFP-positive motor axons is faster than in ChAT.eGFP-negative sensory axons, which can be reliably differentiated using axon widths43.
We have previously identified motor neuron axons using the ChAT.eGFP mouse in vivo43. We now report that HB9.GFP mice can also be used to achieve motor neuron axon identification in vivo. Indeed, Figure 3 presents a series of time-lapse images of HB9.GFP axons containing retrogradely moving HCT-555-positive signaling endosomes. Note that, unlike ChAT-driven expression, GFP has a more punctate/granular pattern in HB9.GFP axons; the reason for this is unclear.
We have previously described how to monitor in vivo mitochondrial dynamics in sciatic nerves via intrasciatic nerve injections of the mitochondrial-targeting dye, tetramethylrhodamine, ethyl ester, perchlorate (TMRE)32,36. To reliably differentiate motor versus sensory mitochondria, the Mito.CFP mouse, which expresses CFP under the Thy1 promoter18, can be crossed with transgenic mice that express a fluorescent reporter gene in specific neuronal types. Indeed, by breeding Mito.CFP mice with ChAT.tdTomato mice (referred to as ChAT.tdTomato::Mito.CFP), we could visualize mitochondria specifically in motor axons, as shown in Figure 4. In this live multiplex example, five ChAT.tdTomato axons could be visualized, four of which contain CFP-positive mitochondria. Moreover, the node of Ranvier (white arrow in panel iii) could also be identified. Furthermore, the nodes of Ranvier are clearly detectable in ChAT.eGFP, HB9.GFP and Mito.CFP mice (not shown). These double-transgenic strains enable time-lapse intravital imaging of live, anesthetized mice to monitor motor neuron-specific mitochondrial content and axonal transport dynamics.
Finally, signaling endosomes and mitochondria can be concurrently visualized within the same axons in vivo by injecting HCT into the muscles of Mito.CFP mice (Figure 5). Intramuscular injections of HCT-555 were performed in TA muscle in a Mito.CFP mouse ~4 h prior to imaging. Both mitochondria (i panels) and signaling endosomes (ii panels) were simultaneously visualized in muscle-specific axons (i.e., axons innervating the TA). Indeed, anterogradely (yellow triangles) and retrogradely (green triangles and circles) moving organelles as well as stalled organelles (orange triangles and circles) can be observed. Using this experimental paradigm, one can assess the complex functional interactions between axonal mitochondria and signaling endosomes in vivo. Overall, we demonstrate several different experimental approaches to assess axonal transport of signaling endosomes and/or mitochondria, specifically in cholinergic motor neurons in vivo.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63471/63471fig1v3.jpg" /> 
Figure 1: Sciatic nerve motor axons. (A) Representative single-plane image of eGFP-positive motor axons obtained in vivo from a ChAT.eGFP mouse. (B) Representative single-plane image of tdTomato-positive motor axons in an excised sciatic nerve from a ChAT.tdTomato mouse. Distinctions in axon caliber result from differences in mouse age and size. Scale bars = 50 &#181;m. Abbreviations: eGFP = enhanced green fluorescent protein; ChAT = choline acetyltransferase. <a href="https://www.jove.com/files/ftp_upload/63471/63471fig1v3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63471/63471fig2v2.jpg" /> 
Figure 2: In vivo axonal transport of signaling endosomes in live sciatic nerve motor and sensory neurons of a ChAT.eGFP mouse. (A-C) Representative images of cholinergic axons expressing eGFP (A) and containing HCT-555-positive signaling endosomes (B), and the merge (C). The white arrows highlight an eGFP-positive motor axon&#160;containing HCT-555-positive signaling endosomes, the cyan arrows identify a motor axon&#160;lacking HCT-555-positive signaling endosomes, and the yellow arrows highlight an eGFP-negative sensory axon&#160;transporting HCT-555-positive signaling endosomes. Orange asterisks identify motor axons with weaker eGFP expression. Scale bar = 25 &#181;m. Abbreviations: eGFP = enhanced green fluorescent protein; ChAT = choline acetyltransferase; HCT-555 = tetanus toxin-binding domain. <a href="https://www.jove.com/files/ftp_upload/63471/63471fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63471/63471fig3v2.jpg" /> 
Figure 3: Time-lapse image series representing in vivo axonal transport of signaling endosomes in live motor neurons of an HB9.GFP mouse. (A-D) Time-lapse images taken every 3 s depicting motor neuron axons expressing green fluorescent protein (i) and containing HCT-555-positive signaling endosomes (ii), and the merge (iii). Each circle of the same color identifies the same moving endosome across different frames. Retrograde movement is from right to left. Scale bar = 10 &#181;m. Abbreviations: GFP = green fluorescent protein; HCT-555 = tetanus toxin-binding domain. <a href="https://www.jove.com/files/ftp_upload/63471/63471fig3v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/63471/63471fig4v2.jpg" /> 
Figure 4: In vivo axonal transport of mitochondria in live sciatic nerve motor neurons of a ChAT.tdTomato:: Mito.CFP: mouse. (A-C) Representative images of tdTomato-positive motor axons (A), containing CFP-positive mitochondria (B), and the merge (C). The inset images i-iii contain a higher magnification from each panel. The white arrow represents a suspected node of Ranvier. Scale bars = 25 (A-C) and 10 &#181;m (i-iii). Abbreviations: ChAT = choline acetyltransferase; CFP = cyan fluorescent protein. <a href="https://www.jove.com/files/ftp_upload/63471/63471fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/63471/63471fig5v2.jpg" /> 
Figure 5: Time-lapse image series representing concurrent in vivo axonal transport of mitochondria and signaling endosomes in live sciatic nerve motor neurons of a Mito.CFP mouse. (A-C) Time-lapse images taken every 3 s depicting axonal transport of both mitochondria (i) and signaling endosomes (ii) within the same sciatic nerve axon (iii). The yellow triangles identify anterogradely moving cargoes, the green circles/triangles identify retrogradely moving cargoes, and the orange circles/triangles identify stationary cargoes. Anterograde movement is from left to right, whereas retrograde movement is in the opposite direction. Scale bar = 10 &#181;m. Abbreviations: HCT-555 = tetanus toxin-binding domain; CFP = cyan fluorescent protein. <a href="https://www.jove.com/files/ftp_upload/63471/63471fig5v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Expanding the Toolkit for In Vivo Imaging of Axonal Transport at JoVE.com

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Using transgenic fluorescent mice, detailed protocols are described to assess in vivo axonal transport of signaling endosomes and mitochondria within motor and sensory axons of the intact sciatic nerve in live animals.

Expanding the Toolkit for In Vivo Imaging of Axonal Transport

Axonal transport maintains neuronal homeostasis by enabling the bidirectional trafficking of diverse organelles and cargoes. Disruptions in axonal ...

expanding-the-toolkit-for-in-vivo-imaging-of-axonal-transport

Research

JoVE Journal

Neuroscience

3.2K Views.  University College London. Using transgenic fluorescent mice, detailed protocols are described to assess in vivo axonal transport of signaling endosomes and mitochondria within motor and sensory axons of the intact sciatic nerve in live animals. 

Video: Expanding the Toolkit for In Vivo Imaging of Axonal Transport

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 ...

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of the olfactory epithelium continuously give rise to new sensory neurons that extend their axons into the olfactory bulb, where they face the challenge to integrate into existing circuitry. Because of this particular feature, the olfactory system represents a unique opportunity to monitor axonal wiring and guidance, and to investigate synapse formation. Here we describe a procedure for in vivo labeling of sensory neurons and subsequent visualization of axons in the olfactory system of larvae of the amphibian Xenopus laevis. To stain sensory neurons in the olfactory organ we adopt the electroporation technique. In vivo electroporation is an established technique for delivering fluorophore-coupled dextrans or other macromolecules into living cells. Stained sensory neurons and their axonal processes can then be monitored in the living animal either using confocal laser-scanning or multiphoton microscopy. By reducing the number of labeled cells to few or single cells per animal, single axons can be tracked into the olfactory bulb and their morphological changes can be monitored over weeks by conducting series of in vivo time lapse imaging experiments. While the described protocol exemplifies the labeling and monitoring of olfactory sensory neurons, it can also be adopted to other cell types within the olfactory and other systems.

The Olfactory System as a Model to Study Axonal Growth Patterns and Morphology In Vivo

We describe a protocol for in vivo labeling of olfactory sensory neurons by electroporation and subsequent confocal laser-scanning or multiphoton microscopy to visualize neuronal morphology and its development over time.

The olfactory system has the unusual capacity to generate new neurons throughout the lifetime of an organism. Olfactory stem cells in the basal portion of ...

In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing

Electrophysiology enables the objective assessment of peripheral nerve function in vivo. Traditional nerve conduction measures such as amplitude and latency detect chronic axon loss and demyelination, respectively. Axonal excitability techniques &#34;by threshold tracking&#34; expand upon these measures by providing information regarding the activity of ion channels, pumps and exchangers that relate to acute function and may precede degenerative events. As such, the use of axonal excitability in animal models of neurological disorders may provide a useful in vivo measure to assess novel therapeutic interventions. Here we describe an experimental setup for multiple measures of motor axonal excitability techniques in the rat ulnar nerve.
The animals are anesthetized with isoflurane and carefully monitored to ensure constant and adequate depth of anesthesia. Body temperature, respiration rate, heart rate and saturation of oxygen in the blood are continuously monitored. Axonal excitability studies are performed using percutaneous stimulation of the ulnar nerve and recording from the hypothenar muscles of the forelimb paw. With correct electrode placement, a clear compound muscle action potential that increases in amplitude with increasing stimulus intensity is recorded. An automated program is then utilized to deliver a series of electrical pulses which generate 5 specific excitability measures in the following sequence: stimulus response behavior, strength duration time constant, threshold electrotonus, current-threshold relationship and the recovery cycle.
Data presented here indicate that these measures are repeatable and show similarity between left and right ulnar nerves when assessed on the same day. A limitation of these techniques in this setting is the effect of dose and time under anesthesia. Careful monitoring and recording of these variables should be undertaken for consideration at the time of analysis.

In Vivo Electrophysiological Measurement of the Rat Ulnar Nerve with Axonal Excitability Testing

Axonal excitability techniques provide a powerful tool to examine pathophysiology and biophysical changes that precede irreversible degenerative events. This manuscript demonstrates the use of these techniques on the ulnar nerve of anesthetized rats.

Electrophysiology enables the objective assessment of peripheral nerve function in vivo. Traditional nerve conduction measures such as amplitude and ...

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging methods exist for examining the brain tissue of live newborn mammals. Herein we summarize a protocol for imaging individual cortical neurons in living neonatal mice. This protocol includes the following two methodologies: (1) the Supernova system for sparse and bright labeling of cortical neurons in the developing brain, and (2) a surgical procedure for the fragile neonatal skull. This protocol allows the observation of temporal changes of individual cortical neurites during neonatal stages with a high signal-to-noise ratio. Labeled cell-specific gene silencing and knockout can also be achieved by combining the Supernova with RNA interference and CRISPR/Cas9 gene editing systems. This protocol can, thus, be used for analyzing the developmental dynamics of cortical neurons, molecular mechanisms that control the neuronal dynamics, and changes in neuronal dynamics in disease models.

In Vivo Two-photon Imaging of Cortical Neurons in Neonatal Mice

We present an in vivo two-photon imaging protocol for imaging the cerebral cortex of neonatal mice. This method is suitable for analyzing the developmental dynamics of cortical neurons, the molecular mechanisms that control the neuronal dynamics, and the changes in neuronal dynamics in disease models.

Two-photon imaging is a powerful tool for the in vivo analysis of neuronal circuits in the mammalian brain. However, a limited number of in vivo imaging ...

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 ...

Axonal Transport of Organelles in Motor Neuron Cultures using Microfluidic Chambers System

Motor neurons (MNs) are highly polarized cells with very long axons. Axonal transport is a crucial mechanism for MN health, contributing to neuronal growth, development, and survival. We describe a detailed method for the use of microfluidic chambers (MFCs) for tracking axonal transport of fluorescently labeled organelles in MN axons. This method is rapid, relatively inexpensive, and allows for the monitoring of intracellular cues in space and time. We describe a step by step protocol for: 1) Fabrication of polydimethylsiloxane (PDMS) MFCs; 2) Plating of ventral spinal cord explants and MN dissociated culture in MFCs; 3) Labeling of mitochondria and acidic compartments followed by live confocal imagining; 4) Manual and semiautomated axonal transport analysis. Lastly, we demonstrate a difference in the transport of mitochondria and acidic compartments of HB9::GFP ventral spinal cord explant axons as a proof of the system validity. Altogether, this protocol provides an efficient tool for studying the axonal transport of various axonal components, as well as a simplified manual for MFC usage to help discover spatial experimental possibilities.

Axonal transport is a crucial mechanism for motor neuron health. In this protocol we provide a detailed method for tracking the axonal transport of acidic compartments and mitochondria in motor neuron axons using microfluidic chambers.

Motor neurons (MNs) are highly polarized cells with very long axons. Axonal transport is a crucial mechanism for MN health, contributing to neuronal ...

Imaging and Analysis of Neurofilament Transport in Excised Mouse Tibial Nerve

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

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

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

Transpupillary Two-Photon In Vivo Imaging of the Mouse Retina

The retina transforms light signals from the environment into electrical signals that are propagated to the brain. Diseases of the retina are prevalent and cause visual impairment and blindness. Understanding how such diseases progress is critical to formulating new treatments. In vivo&#160;microscopy in animal models of disease is a powerful tool for understanding neurodegeneration and has led to important progress towards treatments of conditions ranging from Alzheimer&#39;s disease to stroke. Given that the retina is the only central nervous system structure inherently accessible by optical approaches, it naturally lends itself towards in vivo imaging. However, the native optics of the lens and cornea present some challenges for effective imaging access.
This protocol outlines methods for in vivo two-photon imaging of cellular cohorts and structures in the mouse retina at cellular resolution, applicable for both acute- and chronic-duration imaging experiments. It presents examples of retinal ganglion cell (RGC), amacrine cell, microglial, and vascular imaging using a suite of labeling techniques including adeno-associated virus (AAV) vectors, transgenic mice, and inorganic dyes. Importantly, these techniques extend to all cell types of the retina, and suggested methods for accessing other cellular populations of interest are described. Also detailed are example strategies for manual image postprocessing for display and quantification. These techniques are directly applicable to studies of retinal function in health and disease.

In vivo imaging is a powerful tool for the study of biology in health and disease. This protocol describes transpupillary imaging of the mouse retina with a standard two-photon microscope. It also demonstrates different in vivoimaging methods to fluorescently label multiple cellular cohorts of the retina.

The retina transforms light signals from the environment into electrical signals that are propagated to the brain. Diseases of the retina are prevalent ...

&#181;Tongue: A Microfluidics-Based Functional Imaging Platform for the Tongue In Vivo

Intravital fluorescence microscopy is a tool used widely to study multicellular dynamics in a live animal. However, it has not been successfully used in the taste sensory organ. By integrating microfluidics into the intravital tongue imaging window, the &#181;Tongue provides reliable functional images of taste cells in vivo under controlled exposure to multiple tastants. In this paper, a detailed step-by-step procedure to utilize the &#181;Tongue system is presented. There are five subsections: preparing of tastant solutions, setting up of a microfluidic module, sample mounting, acquiring functional image data, and data analysis. Some tips and techniques to solve the practical issues that may arise when using the &#181;Tongue are also presented.

 &#181;Tongue: A Microfluidics-Based Functional Imaging Platform for the Tongue In Vivo

The article introduces the &#181;Tongue (microfluidics-on-a-tongue) device for functional taste cell imaging in vivo by integrating microfluidics into an intravital imaging window on the tongue.

Intravital fluorescence microscopy is a tool used widely to study multicellular dynamics in a live animal. However, it has not been successfully used in ...

Monitoring Epileptiform Events in Drosophila via Calcium Imaging

Ex Vivo Calcium Imaging for Drosophila Model of Epilepsy

Epilepsy is a neurological disorder characterized by recurrent seizures, partially correlated with genetic origin, affecting over 70 million individuals worldwide. Despite the clinical importance of epilepsy, the functional analysis of neural activity in the central nervous system is still to be developed. Recent advancements in imaging technology, in combination with stable expression of genetically encoded calcium indicators, such as GCaMP6, have revolutionized the study of epilepsy at both brain-wide and single-cell resolution levels. Drosophila melanogaster has emerged as a tool for investigating the molecular and cellular mechanisms underlying epilepsy due to its sophisticated molecular genetics and behavioral assays. In this study, we present a novel and efficient protocol for ex vivo calcium imaging in GCaMP6-expressing adult Drosophila to monitor epileptiform activities. The whole brain is prepared from cac, a well-known epilepsy gene, knockdown flies for calcium imaging with a confocal microscope to identify the neural activity as a follow-up to the bang-sensitive seizure-like behavior assay. The cac knockdown flies showed a higher rate of seizure-like behavior and abnormal calcium activities, including more large spikes and fewer small spikes than wild-type flies. The calcium activities were correlated to seizure-like behavior. This methodology serves as an efficient methodology in screening the pathogenic genes for epilepsy and exploring the potential mechanism of epilepsy at the cellular level.

Author Spotlight: Insights into the Techniques and Findings of Recent Advancements in Epilepsy Research 

Here, we present a protocol for ex vivo calcium imaging in GCaMP6-expressing adult Drosophila to monitor epileptiform activities. The protocol provides a valuable tool for investigating ictal events in adult Drosophila through ex vivo calcium imaging, allowing for exploration of the potential mechanisms of epilepsy at the cellular levels.

Epilepsy is a neurological disorder characterized by recurrent seizures, partially correlated with genetic origin, affecting over 70 million individuals ...