All animal experiments have been conducted in accordance with the National Institutes of Health Animal Use Guidelines. This work was supported in part by grants from National Institutes of Health (R01NS093073 and R21AA024873) to C. Gu.

All methods described below have been approved by the Institutional Animal Care and Use Committee (IACUC) of the Ohio State University.
1. Coverslip Preparation
<ol>
	<li>Place one or more boxes of 12 mm or 25 mm coverslips into a glass beaker containing 70% nitric acid and incubate the coverslips at room temperature overnight. 
	NOTE: Do not wash the 25 mm coverslips in the same beaker as the 12 mm coverslips. It is better to wash them separately.</li>
	<li>Transfer all coverslips into a 4 L beaker filled with 2.5 L of ddH2O and shake the beaker containing the coverslips for 1 h at 100 rpm. Use rubber bands to hold the beaker during shaking. Repeat the rinse with fresh 2.5 L ddH2O four times.</li>
	<li>Transfer cleaned coverslips to a dry flask with a metal cover. Bake the coverslips in an oven at 225 &#176;C for 6 h and allow them to cool to room temperature. 
	NOTE: The 25 mm coverslip is prone to breaking. Separate coverslips one by one and air dry, then bake them in an oven.</li>
	<li>Store the dried and cleaned coverslips at room temperature in a sterilized plastic container until usage.</li>
	<li>Coat the coverslips as follows:
	<ol>
		<li>Prepare fresh coating solution in a centrifuge tube.</li>
		<li>Place each coverslip into one well of a 24-well (or 6-well) plate. Place 30 (or 100) &#181;L of coverslip coating solution (Table 1) onto a 12 mm (or 25 mm) coverslip and swirl. Add 1 mL of sterile phosphate-buffered saline (PBS, tissue culture grade) to the well. These plates can either be used immediately or stored at 4 &#176;C in sterile PBS for up to a week.</li>
		<li>On the day of dissection, remove the PBS. Air-dry and place the plates under UV light (30W) for 2-4 h. 
		NOTE: The last step should be completed with the UV light on in a cell culture laminar flow hood. The door of the hood should be lifted slightly to allow air flow for drying the coverslips.</li>
	</ol>
	</li>
</ol>
2. Dissection, Dissociation, and Culture of Hippocampal Neurons from Pregnant Mouse/Rat
<ol>
	<li>Dissection and dissociation of hippocampal neurons for culture
	<ol>
		<li>Thaw protease enzyme solution (3 mg/mL protease 23 in SLDS, Table 1) and pre-warm plating media (Table 2) at 37 &#176;C.</li>
		<li>Dissect hippocampus and rinse with SLDS, according to the methods described in previous publication14. 
		NOTE: Four hippocampi provide enough cells for one 24-well or one 6-well plate.</li>
		<li>Remove SLDS, add 2 mL of protease enzyme solution, and incubate the sample at 37 &#176;C, 5% CO2 for 15 min.</li>
		<li>Wash the hippocampus explants with 5-10 mL of plating medium twice. Add 5 mL of plating medium. Dissociate hippocampal explants into single cells by generating a small vortex using a pipette with a 1-mL plastic tip. Pipette up and down about 40 times, avoiding the formation of oxygen bubbles, until the solution becomes cloudy and no large pieces of explant remain. 
		NOTE: Remember to leave some media during washes, the hippocampal explants should remain submerged in the medium during the whole process.</li>
		<li>Centrifuge the cells at 1,125 x g for 3 min. Remove the supernatant with the cells submerged in media. Resuspend the cells in 145 mL of plating media. Add 1 mL/well of the cell suspension to a 24-well plate (3 mL/well to a 6-well plate). 
		NOTE: 145 mL of plating media is used to produce a cell suspension that has a low cell density. This volume will allow the plating of cells into six 24-well plates, eight 6-well plates, or various combinations of 24- or 6-well plates depending on the experiments to be conducted. Each well of the 24-well plate will have about 3 x 104 cells9.</li>
		<li>Incubate the cells in the plating media at 37 &#176;C, 5% CO2 for 2-4 h, then remove all plating media and replace it with fresh maintenance media.</li>
		<li>After 2 days (2 days in vitro, 2DIV), replace half of the media with 2 &#181;M Ara-C (arabinosylcytosine) dissolved in maintenance media which inhibits growth of fibroblasts, endothelial, and glial cells. After exposing cells to Ara-C for two days, replace the media with fresh maintenance media.</li>
	</ol>
	</li>
	<li>Transfection for the visualization of cell morphology 
	NOTE: Transfect the cells with fluorescent constructs of green fluorescent protein (GFP), yellow fluorescent protein (YFP), mCherry, etc. to illuminate the morphology of individual neurons.
	<ol>
		<li>Dilute the constructs from stock (1 &#181;g/&#181;L) into Opti-MEM media (0.8 &#181;g of the construct in 50 &#181;L of Opti-MEM media) and vortex. Prepare 1 construct dilution for each well. 
		NOTE: A 24-well plate requires 0.8 &#181;g of construct in 50 &#181;L of Opti-MEM media. A 6-well plate requires 2.4 &#181;g of construct in 150 &#181;L Opti-MEM media.</li>
		<li>Dilute liposome-mediated transfection reagent into Opti-MEM media (1.5 &#181;L of the transfection reagent in 50 &#181;L of Opti-MEM media) and vortex. Prepare 1 transfection dilution for each well. 
		NOTE: A 24-well plate requires 1.5 &#181;L of transfection reagent in 50 &#181;L of Opti-MEM media. A 6-well plate requires 4.5 &#181;L of transfection reagent in 150 &#181;L of Opti-MEM media.</li>
		<li>Prepare a 1:1 mixture (100 &#181;L) of the construct in Opti-Mem media and transfection reagent in Opti-MEM media and vortex. Incubate the mixture at room temperature for 20 min. 
		NOTE: Per well, there should be about 100 &#181;L of solution containing construct, transfection reagent, and Opti-MEM media if using a 24-well plate (300 &#181;L for 6-well plate).</li>
		<li>Remove half of the media (500 &#181;L for 24-well plate, 1.5 mL for 6-well plate) from each well and transfer this media into a clean conical tube. Keep this tube in the incubator. Then add the 100 &#181;L mixture (step 2.2.3) to the remaining media in the well. Allow the cells to incubate at 37 &#176;C for 20-30 min.</li>
		<li>During the incubation (step 2.2.4), add one volume of fresh maintenance media to the media in the conical tube (step 2.2.4). Place the mixture in the same incubator (37 &#176;C and 5% CO2).</li>
		<li>Following the incubation, remove all media containing the transfection solution from the wells and replace it with the 1:1 mixture of old and fresh maintenance media in the conical tube (step 2.2.5). 
		NOTE: Depending on the size of the constructs, cells may begin expressing within 12 h; for larger constructs, 3-4 days may be needed before the peak expression of the constructs is reached.</li>
	</ol>
	</li>
</ol>
3. Setting Up the Puffing Pipette Apparatus
NOTE: There are five basic components to this set-up: the glass pipette, the tubing, the syringe, the micromanipulator, and the buffer (Hank&#39;s). Any buffer that has physiologically relevant salt concentrations could be used in this set-up.
<ol>
	<li>Pull a borosilicate pipette to have around 45-&#181;m-diameter opening tip using a micropipette puller and the specifications in Table 3. 
	NOTE: It was found that 45 &#181;m is a suitable size for the diameter of the pipette opening under experimental conditions. Slight variations in the opening size should not significantly affect results. It is important to note that larger departures from this number should still work for varicosity induction by adjusting the height of the syringe that is linked to the pipette via tubing, but this will require recalibration of the pressure value.</li>
	<li>Insert the un-pulled end of the pipette into thin rubber tubing. 
	NOTE: The tubing wall should be thin and rigid to minimize the fluid resistance, ensuring that the height of the syringe is directly proportional to the pressure of fluid flowing through the pipette with negligible resistance. The tubing should be no longer than about 0.6 m in length to further minimize the resistance.</li>
	<li>Connect the other end of the tubing to a 10-mL plastic syringe through a connector with a turn-able valve. 
	NOTE: The plastic syringe needs to be held at a constant, measurable height to ensure an accurate measure for the assumed pressure flowing from the pipette. A height of 190 mm was utilized, since this provides minimal pressure but reliably induces varicosities in axons. Other height values may also be used if the pressure cause the cells to detach from the coverslip surface. It is recommended to use the same setting throughout the experiment.</li>
	<li>Insert the pipette into the screw top of the micromanipulator to hold the pipette in place. Screw the metal, flat topped screw attached to the micromanipulator arm so that it holds the un-pulled end of the pipette just above where the rubber tubing is attached. Angle the pipette at a 45&#176; angle with the surface of the neuron-containing coverslips. 
	NOTE: The micromanipulator should be constructed so that the pipette can be held without constricting the rubber tubing. The micromanipulator must allow motion of the pipette in the x, y, and z directions to accurately position it within range of the cells. A graphic as well as a photograph of the apparatus is provided below (Figure 1).</li>
	<li>Turn on a fluorescence filter, open the aperture, and ensure a fluorescent spot can be seen coming through the objective. Using the micromanipulator, move the pipette into a position that lines up the tip with the fluorescent spot and lower the pipette into a position just above the height of the cell culture dish (35 mm x 10 mm). Once it is in position, turn on the transmitted light and turn off fluorescence.</li>
	<li>Using tweezers, add one coverslip (with the cells facing up toward the pipette) from the cell culture plate to the cell culture dish containing about 2 mL of Hank&#39;s buffer at room temperature.</li>
	<li>Place the culture dish containing the coverslip onto the scope.</li>
	<li>Using the fine focus knob and 20X objective, focus on a plane about four full turns above the cells (about 0.4 mm). Use the micromanipulator to position the pipette tip so that it is focused and in the center, left-hand side of the plane as seen through the eye-pieces. 
	NOTE: Dendrites are projections that should be in the same frame as the cell body they originate from. To induce axonal varicosities, one should follow longer projections from the cell body to medial and distal axons.</li>
</ol>
4. Calibrating the Pressure of the Pipette Utilizing a Stretchable-membrane System
<ol>
	<li>Setup the puffing pipette apparatus with the exact same parameters as described above over a system containing a microfabricated silicone membrane (500 &#181;m in diameter and 50 &#181;m thick) and focus the microscope onto the surface of the membrane. 
	NOTE: This system was designed to measure small pressure values and an explanation of the system has been published before10. This measurement only needs to be done once for puffing experiments if the exact same setting of puffing is used.</li>
	<li>Focus the microscope on the arrays of microdots (4 &#181;m in diameter and 10 &#181;m apart, as measured between centers). Turn the stop valve and allow fluid to flow out of the pipette. Examine the deflection of the membrane in response to the fluid pressure with a confocal microscope.</li>
	<li>Utilize a linear model to determine the corresponding pressure associated with deflection of the membrane. 
	NOTE: While a recent study found that a deflection of w= -3.143 &#177; 0.69 &#181;m was obtained for 190 mm syringe height, this produced a pressure of 0.25 &#177; 0.06 nN/&#181;m2 from the current pipette setup, which is within the physiological and pathological range, though the syringe height is not the sole determinant. Other factors influence the exact pressure value on the neurons as well, including pipette opening, positioning (distance and angle) of the pipette tip relative to the cells, and even the flexibility of tubing4,10.</li>
</ol>
5. Puffing to Induce Varicosities
<ol>
	<li>Once the pipette is in position, focus the microscope on a plane that contains a region of interest. Position the cells and processes in the left half of the imaging field (seen by live capture of the camera) within the puffing area, while the right half is outside of puffing area. 
	NOTE: Due to extensive outgrowth of axons and dendrites of neurons, it is unlikely that all the processes of a neuron are within the same puffing area. This is actually one advantage of this system, which allows the examination of the local effect of puffing. There are two types of puffing that can be utilized to induce varicosities: long duration puffing and pulse puffing.</li>
	<li>Puffing
	<ol>
		<li>Long duration puffing
		<ol>
			<li>Position the syringe at the height, 190 mm above the coverslip containing cultured neurons4. Begin time-lapse imaging of area of interest with an optimized exposure time revealing clear neuronal morphology. Capture at least 15 frames at a 2 s interval (30 s total) as the baseline. The interval can be adjusted based on the experiment. At frame 16, open the turn-able stop valve of the syringe and allow fluid to flow for 75 frames (150 s). At frame 75, turn the valve off so that fluid flow stops. Stop video capture. 
			Note: This should induce axonal varicosities in neurons 7-9 DIV within 5-10 s, whereas older neurons take longer to form varicosities.</li>
		</ol>
		</li>
		<li>Pulse (2 s duration) puffing
		<ol>
			<li>Position the syringe at the proper height (190 mm). Begin time-lapse imaging and capture at least 15 frames (30 s) at a 2 s interval. At frame 16, open the valve of the syringe and allow fluid to flow, then close the valve after 2 s. Wait 5-20 s (depending on the frequency to be tested) before opening the valve again. Repeat opening and closing of the valve to create fluid pulses, and continue for a total period of 150-300 s. Continue time-lapse imaging for 20 min to capture the recovery stage. 
			&#8203;NOTE: When the interval is longer than 10 s, the pulses induce drastically less varicosities. At a 20 s interval, no varicosities can be induced by the pulses4. To obtain reliable, consistent varicosity formation, it is advised the syringe height should be around 190 mm. Syringe height and fluid pressure should not be raised to a point where cells begin detaching from the coverslip surface.</li>
		</ol>
		</li>
		<li>Before a day of use, clean the set-up (pipette, tubing, and syringe) by flowing about 10 mL 70% Ethanol through the set-up. Following the ethanol wash, flow 10 mL of Hank&#39;s buffer (or desired buffer) through the set-up to prime the system for that day&#39;s use. After finishing experiments for the day, clean the set-up as previously described with ethanol to prevent growth of contaminants overnight.</li>
	</ol>
	</li>
</ol>
6. Complimenting Methods
NOTE: The puffing assay can be combined with a variety of different techniques that are discussed in the Introduction section. Here, the focus is on the core techniques that are most frequently used together with the puffing assay, including high-resolution fluorescence timelapse imaging, calcium imaging, and recovery assays. These are discussed below.
<ol>
	<li>Conduct high-resolution fluorescent time-lapse imaging by following the protocol below
	<ol>
		<li>Track the movement of fluorescently-tagged proteins within neurons with a 100X oil lens. Neurons are transfected with the proper cDNA construct. Capture for 150 frames (total 300 s) with a 2 s interval. 
		NOTE: Sometimes, multiple-color time-lapse imaging is performed to image the movement of more than one fluorescently-tagged protein simultaneously.</li>
		<li>Generate a kymograph either through the video capturing software or through ImageJ (NIH). 
		NOTE: The speed and directionality of travel can be determined using the kymograph. *Be sure to note the direction the soma lies from the region captured. Travel towards the soma is retrograde, while travel away from the soma to the axonal tip is anterograde.</li>
		<li>Repeat this process following puffing, or a kymograph can be created using the image capture during the puffing assay. 
		NOTE: The effects of puffing on axonal transport of mitochondria and synaptic proteins were shown in a recent paper4.</li>
	</ol>
	</li>
	<li>Calcium imaging
	<ol>
		<li>Add 1 &#181;L (for a 24-well plate) or 3 &#181;L (for a 6-well plate) of 5 mM Fluo-4 AM to the culture media in one well and incubate at 37 &#176;C for 30 min.</li>
		<li>Wash the cells twice with 1 mL Hank&#39;s buffer. 
		NOTE: Loaded neurons emit green fluorescence through a FITC filter set. Loci denote regions of appreciable calcium within the cell. When combined with puffing, an increase in intensity (increase in internal calcium level) can be seen.</li>
	</ol>
	</li>
	<li>Varicosity recovery 
	NOTE: Immediately following a puffing experiment, a 20 min recovery experiment can be carried out.
	<ol>
		<li>Capture images for 300 frames (1,200 s) with a 4 s interval. 
		NOTE: A cell transfected with soluble YFP/mCherry/GFP should show a moderate level (less than 50%) of recovery by the end of the imaging session. Axonal recovery depends on a number of factors, such as the developmental stage of cultured neurons4. This can also be combined with high-resolution fluorescent imaging and calcium imaging described above.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Niki&#263;, I., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+reversible+form+of+axon+damage+in+experimental+autoimmune+encephalomyelitis+and+multiple+sclerosis.">A reversible form of axon damage in experimental autoimmune encephalomyelitis and multiple sclerosis.</a> Nat. Med. 17, 495-499 (2011).</li><li>Yang, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulation+of+axon+degeneration+after+injury+and+in+development+by+the+endogenous+calpain+inhibitor+calpastatin.">Regulation of axon degeneration after injury and in development by the endogenous calpain inhibitor calpastatin.</a> Neuron. 80 (5), 1175-1189 (2013).</li><li>Debanne, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Information+processing+in+the+axon.">Information processing in the axon.</a> Nat. Rev. Neurosci. 5, 304-316 (2004).</li><li>Gu, Y., Jukkola, P., Wang, Q., Esparza, T., Zhao, Y., Brody, D., Gu, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Polarity+of+varicosity+initiation+in+central+neuron+mechanosensation.">Polarity of varicosity initiation in central neuron mechanosensation.</a> J Cell Biol. 216 (7), 2179-2199 (2017).</li><li>Chung, R. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mild+axonal+stretch+injury+in+vitro+induces+a+progressive+series+of+neurofilament+alterations+ultimately+leading+to+delayed+axotomy.">Mild axonal stretch injury in vitro induces a progressive series of neurofilament alterations ultimately leading to delayed axotomy.</a> J. Neurotrauma. 22 (10), 1081-1091 (2005).</li><li>Staal, J. A., Dickson, T. C., Chung, R. S., Vickers, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cyclosporin-A+treatment+attenuates+delayed+cytoskeletal+alterations+and+secondary+axotomy+following+mild+axonal+stretch+injury.">Cyclosporin-A treatment attenuates delayed cytoskeletal alterations and secondary axotomy following mild axonal stretch injury.</a> Dev. Neurobiol. 67 (14), 1831-1842 (2007).</li><li>Tang-Schomer, M. D., Johnson, V. E., Baas, P. W., Stewart, W., Smith, D. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Partial+interruption+of+axonal+transport+due+to+microtubule+breakage+accounts+for+the+formation+of+periodic+varicosities+after+traumatic+axonal+injury.">Partial interruption of axonal transport due to microtubule breakage accounts for the formation of periodic varicosities after traumatic axonal injury.</a> Exp. Neurol. 233 (1), 364-372 (2012).</li><li>Donkin, J. J., Vink, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanisms+of+cerebral+edema+in+traumatic+brain+injury:+therapeutic+developments.">Mechanisms of cerebral edema in traumatic brain injury: therapeutic developments.</a> Curr Opin Neurol. 23 (3), 293-299 (2010).</li><li>Gardner, A., Jukkola, P., Gu, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Myelination+of+rodent+hippocampal+neurons+in+culture.">Myelination of rodent hippocampal neurons in culture.</a> Nat Protoc. 7 (10), 1774-1782 (2012).</li><li>Wang, Q., Zhang, X., Zhao, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Micromechanical+stimulator+for+localized+cell+loading:+fabrication+and+strain+analysis.">Micromechanical stimulator for localized cell loading: fabrication and strain analysis.</a> J. Micromech. Microeng. 23, 015002 (2013).</li><li>Lu, Y. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Viscoelastic+properties+of+individual+glial+cells+and+neurons+in+the+CNS.">Viscoelastic properties of individual glial cells and neurons in the CNS.</a> Proc. Natl. Acad. Sci. USA. 103 (47), 17759-17764 (2006).</li><li>Elkin, B. S., Azeloglu, E. U., Costa, K. D., Morrison, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanical+heterogeneity+of+the+rat+hippocampus+measured+by+atomic+force+microscope+indentation.">Mechanical heterogeneity of the rat hippocampus measured by atomic force microscope indentation.</a> J. Neurotrauma. 24 (5), 812-822 (2007).</li><li>Franze, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neurite+branch+retraction+is+caused+by+a+threshold-dependent+mechanical+impact.">Neurite branch retraction is caused by a threshold-dependent mechanical impact.</a> Biophys. J. 97 (7), 1883-1890 (2009).</li><li>Camp&#224;s, O., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantifying+cell-generated+mechanical+forces+within+living+embryonic+tissues.">Quantifying cell-generated mechanical forces within living embryonic tissues.</a> Nat. Methods. 11 (2), 183-189 (2014).</li></ol>

The authors have nothing to disclose.

The procedure of this microbiomechanical assay is straight forward. It will produce reliable results, if all its steps are carefully carried out. There are several key steps that, if improperly performed, will hinder successful data collection. The critical steps begin upstream of the actual application of the puffing stimulus. Careful dissection, culturing, and care of the primary neuron culture are paramount. If the cultured neurons are not healthy, they will not react consistently, since they might have already been p...

Varicosity formation, or swelling/beading, along axons, is a prominent feature of neurodegeneration observed in many disorders or injuries of the central nervous system, including multiple sclerosis, Alzheimer&#39;s disease, Parkinson&#39;s disease, and traumatic brain injury1,2. Despite the significant physiological impacts of axonal varicosities on action potential propagation and synaptic transmission3, how the varicosities are generated remains unknown. Recently, using a newly-established microbiomechanical assay on cultured hippocampal neurons from rodents, we found that mechanical stimuli can induce varicosities in these neurons with highly intriguing features. First, varicosity induction is rapid (&#60;10 s) and this process is unexpectedly reversible. Second, varicosity initiation depends on the strength of puffing pressure: the higher the pressure, the faster the initiation. Third, varicosity initiation depends on neuronal age. The axons of younger neurons appear more responsive to mechanical stress, compared to those of older neurons. Fourth, the varicosities form along the axons of hippocampal neurons, while the dendrites and initial axon segments of these neurons display no change under the same puffing condition. Thus, our study revealed a novel feature of neuronal polarity. These findings with the in vitro system are physiologically relevant. Using an in vivo model for mild traumatic brain injury (mTBI), we showed that axonal varicosities developed in a multi-focal fashion in the somatosensory cortex of the mice immediately after close-skull impact, consistent with our in vitro results4. It is important to note that our staining and imaging of the mTBI mice only provide a snap shot of neuronal morphological changes, since performing in vivo time-lapse imaging of neuronal morphology during a mechanical impact is still not feasible.
This fluid-puffing system allowed us not only to capture unique features associated with mechanical stress-induced varicosity formation, but also to determine the underlying mechanism. By testing different extracellular solutions, the blockers and openers of different candidates of mechanosensitive ion channels, and cellular electrophysiology, we identified that the transient receptor potential cation channel subfamily V member 4 (TRPV4) channel that is permeable to Ca2+ and Na+ and activated by puffing is mainly responsible for detecting initial mechanical stress during axonal varicosity formation4. This was further confirmed with an siRNA knockout approach. Taken together, this new assay system we have developed with hippocampal neuron culture, is highly valuable for studying the micromechanical properties of central neurons, especially in combination with other techniques.
This micromechanical system we have established is unique and differs from the previously-existing systems in several major aspects. First, in this system, the neurons experience out-of-plane mechanical stress in the forms of compression and shearing. During the mechanical impact, neuronal processes remain attached to the coverslip surface and do not move. This differs from other experimental systems that mainly involved bending and in-plane stretching (or tension), for instance, the deflection of bundled axons like moving strings5,6 or stretching axons grown on micropatterned channels and stretchable membranes7,8. Moreover, although axonal varicosities can also be induced in these assays like in our fluid-puffing system, the process in these settings takes much more time (from 10 min to several hours6,7,8) and appears irreversible. Finally, our system using local fluid puffing allows the examination of the spatial features of varicosity formation (e.g., dendrites, dendritic spines, soma, axonal initial segments, axonal terminals), besides its temporal features. Using this system, we discovered several unexpected and unique features of axonal varicosity formation, especially rapid onset, slow reversibility, and axon-dendrite polarity.
The system that we discuss in this paper is compatible with many techniques of molecular and cell biology. For instance, to study the effects of mechanical stress on neuronal morphology and function, it can be used together with myelin coculture, time-lapse imaging of fluorescent resonance energy transfer (FRET) and total internal reflection fluorescence (TIRF), calcium imaging, and patch clamp recording. In this paper, we focus on the core components of the system. Hippocampal neuron culture, the fluid-puffing setup, high-resolution time-lapse imaging for axonal transport, and calcium imaging are illustrated step-by-step below.

<table border="0" cellpadding="0" cellspacing="0" style="width:765px;" width="765">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="40">
			<td height="40" style="height:40px;width:341px;">12 mm coverslips&#160;</td>
			<td style="width:139px;">Warner Instruments&#160;</td>
			<td style="width:119px;">64-0702</td>
			<td style="width:167px;">for 24-well plate&#160;</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">25 mm coverslips&#160;</td>
			<td>Fisher Scientific&#160;</td>
			<td>12-545-102&#160;</td>
			<td>for 6-well plate&#160;</td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Acetic acid</td>
			<td>Fisher Scientific&#160;</td>
			<td>A38-212</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Poly-D-lysine</td>
			<td>Sigma&#160;</td>
			<td>P6407</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Rat tail collagen&#160;</td>
			<td>Roche&#160;</td>
			<td>11 179 179 001&#160;</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">10X PBS&#160;</td>
			<td>National Diagnostics&#160;</td>
			<td>CL-253</td>
			<td></td>
		</tr>
		<tr height="40">
			<td align="left" height="40" style="height:40px;">Na2SO4</td>
			<td>Fisher Scientific&#160;</td>
			<td>S373-500</td>
			<td></td>
		</tr>
		<tr height="40">
			<td align="left" height="40" style="height:40px;">K2SO4</td>
			<td>Fisher Scientific&#160;</td>
			<td>P304-500</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">HEPES&#160;</td>
			<td>Fisher Scientific&#160;</td>
			<td>BP410-500</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">D-glucose&#160;</td>
			<td>Fisher Scientific&#160;</td>
			<td>D16-500</td>
			<td></td>
		</tr>
		<tr height="40">
			<td align="left" height="40" style="height:40px;">MgCl2</td>
			<td>Fisher Scientific&#160;</td>
			<td>BP214-500</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">NaOH</td>
			<td>Fisher Scientific&#160;</td>
			<td>SS255-1</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Protease enzyme&#160;</td>
			<td>Sigma&#160;</td>
			<td>P4032</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">FBS</td>
			<td>Gibco&#160;</td>
			<td>26140</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Sodium pyruvate&#160;</td>
			<td>Gibco&#160;</td>
			<td>11360-070</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">L-glutamine&#160;</td>
			<td>Gibco&#160;</td>
			<td>25030081</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Penicillin/Streptomycin 100x (P/S)</td>
			<td>Gibco&#160;</td>
			<td style="width:119px;">15140122</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">MEM Earle&#39;s Salts&#160;</td>
			<td>Gibco&#160;</td>
			<td>11090</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">B27 supplement&#160;</td>
			<td>Gibco&#160;</td>
			<td>17504-044</td>
			<td></td>
		</tr>
		<tr height="40">
			<td height="40" style="height:40px;">Neurobasal&#160;</td>
			<td>Gibco&#160;</td>
			<td>21103-049</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Arabinosylcytosine (Ara-C)</td>
			<td>Sigma&#160;</td>
			<td>147-94-4</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Opti-MEM media&#160;</td>
			<td>Gibco&#160;</td>
			<td>31985-070</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Lipofectamine 2000</td>
			<td>Invitrogen&#160;</td>
			<td>1854313</td>
			<td>transfection reagent&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Borosilicate rods&#160;</td>
			<td>World Precision Instruments Inc.&#160;</td>
			<td>PG52151-4</td>
			<td>for puffing pipette&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">rubber tubing&#160;</td>
			<td style="width:139px;">Fisher Scientific&#160;</td>
			<td style="width:119px;">14-169-1A</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">10cc plastic syringe and plunger</td>
			<td>Becton Dickinson&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">micromanipulator&#160;</td>
			<td>Sutter Instruments&#160;</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">NaCl&#160;</td>
			<td>Fisher Scientific&#160;</td>
			<td>S640-3</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">KCl&#160;</td>
			<td>Fisher Scientific&#160;</td>
			<td>BP366-500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;">CaCl2</td>
			<td>Fisher Scientific&#160;</td>
			<td>C70-500</td>
			<td></td>
		</tr>
		<tr height="19">
			<td height="19" style="height:19px;width:341px;">Cell culture dish (35 mm x 10 mm)</td>
			<td style="width:139px;">Corning&#160;</td>
			<td style="width:119px;">3294</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:341px;">Fluo-4 AM</td>
			<td style="width:139px;">Molecular Probes</td>
			<td style="width:119px;">F14201</td>
			<td style="width:167px;">for calcium imaging&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:341px;">Mito-YFP construct&#160;</td>
			<td style="width:139px;">Takara Bio Inc.&#160;</td>
			<td style="width:119px;"></td>
			<td style="width:167px;">for cell transfection&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:341px;">YFP-N1 construct&#160;</td>
			<td style="width:139px;">Takara Bio Inc.&#160;</td>
			<td style="width:119px;"></td>
			<td style="width:167px;">for cell transfection&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:341px;">Model P-1000 Flaming/Brown Micropipette puller&#160;</td>
			<td style="width:139px;">Sutter Instruments&#160;</td>
			<td style="width:119px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:341px;">Eclipse TE2000-U Mcroscope&#160;</td>
			<td style="width:139px;">Nikon</td>
			<td style="width:119px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:341px;">Plan Fluor ELWD 20x lens&#160;&#160;</td>
			<td style="width:139px;">Nikon</td>
			<td style="width:119px;">062933</td>
			<td style="width:167px;">objective&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:341px;">Apo TIRF 100x/1.49 oil lens&#160;</td>
			<td style="width:139px;">Nikon</td>
			<td style="width:119px;">MRD01991</td>
			<td style="width:167px;">objective&#160;</td>
		</tr>
	</tbody>
</table>

a microbiomechanical system for studying varicosity formation and recovery in central neuron axons

Axonal varicosities are enlarged structures along the shafts of axons with a high degree of heterogeneity. They are present not only in brains with neurodegenerative diseases or injuries, but also in the normal brain. Here, we describe a newly-established micromechanical system to rapidly, reliably, and reversibly induce axonal varicosities, allowing us to understand the mechanisms governing varicosity formation and heterogeneous protein composition. This system represents a novel means to evaluate the effects of compression and shear stress on different subcellular compartments of neurons, different from other in vitro systems that mainly focus on the effect of stretching. Importantly, owing to the unique features of our system, we recently made a novel discovery showing that the application of pressurized fluid can rapidly and reversibly induce axonal varicosities through the activation of a transient receptor potential channel. Our biomechanical system can be utilized conveniently in combination with drug perfusion, live cell imaging, calcium imaging, and patch clamp recording. Therefore, this method can be adopted for studying mechanosensitive ion channels, axonal transport regulation, axonal cytoskeleton dynamics, calcium signaling, and morphological changes related to traumatic brain injury.

Prior to puffing, axons normally show little varicosity formation. Following puffing with our standard pressure (190 mmH2O height), the axons start to develop many bead-like varicosities. The formation of varicosities is partially reversible, as shown by regions of the axon returning to their pre-puffed state following a 10 min recovery period (Figure 2A-B). After a longer period of recovery (&#62;20 min), some axons completely rec...

Watch this Scientific Journal Video about A Microbiomechanical System for Studying Varicosity Formation and Recovery in Central Neuron Axons at JoVE.com

A Microbiomechanical System for Studying Varicosity Formation and Recovery in Central Neuron Axons

This protocol describes a physiologically relevant, pressurized fluid approach for rapid and reversible induction of varicosities in neurons.

Axonal varicosities are enlarged structures along the shafts of axons with a high degree of heterogeneity. They are present not only in brains with ...

a-microbiomechanical-system-for-studying-varicosity-formation

Research

JoVE Journal

Neuroscience

6.2K Views.  The Ohio State University. This protocol describes a physiologically relevant, pressurized fluid approach for rapid and reversible induction of varicosities in neurons.

Video: A Microbiomechanical System for Studying Varicosity Formation and Recovery in Central Neuron Axons

Systemic and Local Drug Delivery for Treating Diseases of the Central Nervous System in Rodent Models

Thorough preclinical testing of central nervous system (CNS) therapeutics includes a consideration of routes of administration and agent biodistribution in assessing therapeutic efficacy. Between the two major classifications of administration, local vs. systemic, systemic delivery approaches are often preferred due to ease of administration. However, systemic delivery may result in suboptimal drug concentration being achieved in the CNS, and lead to erroneous conclusions regarding agent efficacy. Local drug delivery methods are more invasive, but may be necessary to achieve therapeutic CNS drug levels. Here, we demonstrate proper technique for three routes of systemic drug delivery: intravenous injection, intraperitoneal injection, and oral gavage. In addition, we show a method for local delivery to the brain: convection-enhanced delivery (CED). The use of fluorescently-labeled compounds is included for in vivo imaging and verification of proper drug administration. The methods are presented using murine models, but can easily be adapted for use in rats.

Thorough preclinical testing of drugs that act in the central nervous system often involves assessing and comparing drug biodistribution in association with specific routes of administration. Here, three commonly used methods of systemic delivery (intravenous, intraperitoneal, and oral) as well as a method for local delivery (convection-enhanced delivery) are demonstrated in mice.

Thorough preclinical testing of central nervous system (CNS) therapeutics includes a consideration of routes of administration and agent biodistribution ...

Optic Nerve Transection: A Model of Adult Neuron Apoptosis in the Central Nervous System

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. 
The optic nerve can be accessed within the orbit of the eye and completely transected (axotomized), cutting the axons of the entire RGC population. Optic nerve 
transection is a reproducible model of apoptotic neuronal cell death in the adult CNS 1-4. This model is particularly attractive because the vitreous
 chamber of the eye acts as a capsule for drug delivery to the retina, permitting experimental manipulations via intraocular injections. The diffusion of chemicals 
through the vitreous fluid ensures that they act upon the entire RGC population. Moreover, RGCs can be selectively transfected by applying short interfering RNAs 
(siRNAs), plasmids, or viral vectors to the cut end of the optic nerve 5-7 or injecting vectors into their target, the superior colliculus 8. 
This allows researchers to study apoptotic mechanisms in the desired neuronal population without confounding effects on other bystander neurons or surrounding glia. 
An additional benefit is the ease and accuracy with which cell survival can be quantified after injury. The retina is a flat, layered tissue and RGCs are localized in 
the innermost layer, the ganglion cell layer. The survival of RGCs can be tracked over time by applying a fluorescent tracer (3% Fluorogold) to the cut end of the 
optic nerve at the time of axotomy, or by injecting the tracer into the superior colliculus (RGC target) one week prior to axotomy. The tracer is retrogradely transported, labeling 
the entire RGC population. Because the ganglion cell layer is a monolayer (one cell thick), RGC densities can be quantified in flat-mounted tissue, without the need 
for stereology. Optic nerve transection leads to the apoptotic death of 90% of injured RGCs within 14 days postaxotomy 9-11. RGC apoptosis has a 
characteristic time-course whereby cell death is delayed 3-4 days postaxotomy, after which the cells rapidly degenerate. This provides a time window for 
experimental manipulations directed against pathways involved in apoptosis.

Optic Nerve transection is a widely used model of adult CNS injury. Ninety percent of retinal ganglion cells (RGCs) whose axons are completely transected (axotomy) die within 14 days after axotomy. This model is easily amenable to experimental manipulations and highly reproducible.

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. 
The optic nerve can be ...

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic development and axonal wiring. Recently the dendritic arborization (DA) sensory neurons of the Drosophila larval peripheral nervous system (PNS) have become powerful genetic models in which to elucidate both general and class-specific mechanisms of neuron differentiation.
There are four main DA neuron classes (I-IV)1. They are named in order of increasing dendrite arbor complexity, and have class-specific differences in the genetic control of their differentiation2-10. The DA sensory system is a practical model to investigate the molecular mechanisms behind the control of dendritic morphology11-13 because: 1) it can take advantage of the powerful genetic tools available in the fruit fly, 2) the DA neuron dendrite arbor spreads out in only 2 dimensions beneath an optically clear larval cuticle making it easy to visualize with high resolution in vivo, 3) the class-specific diversity in dendritic morphology facilitates a comparative analysis to find key elements controlling the formation of simple vs. highly branched dendritic trees, and 4) dendritic arbor stereotypical shapes of different DA neurons facilitate morphometric statistical analyses.
DA neuron activity modifies the output of a larval locomotion central pattern generator14-16. The different DA neuron classes have distinct sensory modalities, and their activation elicits different behavioral responses14,16-20. Furthermore different classes send axonal projections stereotypically into the Drosophila larval central nervous system in the ventral nerve cord (VNC)21. These projections terminate with topographic representations of both DA neuron sensory modality and the position in the body wall of the dendritic field7,22,23. Hence examination of DA axonal projections can be used to elucidate mechanisms underlying topographic mapping7,22,23, as well as the wiring of a simple circuit modulating larval locomotion14-17.
We present here a practical guide to generate and analyze genetic mosaics24 marking DA neurons via MARCM (Mosaic Analysis with a Repressible Cell Marker)1,10,25 and Flp-out22,26,27 techniques (summarized in Fig. 1).

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

The dendritic arborization sensory neurons of the Drosophila larval peripheral nervous system are useful models to elucidate both general and neuron class-specific mechanisms of neuron differentiation. We present a practical guide to generate and analyze dendritic arborization neuron genetic mosaics.

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic ...

In ovo Electroporation in Chick Midbrain for Studying Gene Function in Dopaminergic Neuron Development

Dopaminergic neurons located in the ventral midbrain control movement, emotional behavior, and reward mechanisms1-3. The dysfunction of ventral midbrain dopaminergic neurons is implicated in Parkinson's disease, Schizophrenia, depression, and dementia1-5. Thus, studying the regulation of midbrain dopaminergic neuron differentiation could not only provide important insight into mechanisms regulating midbrain development and neural progenitor fate specification, but also help develop new therapeutic strategies for treating a variety of human neurological disorders.
Dopaminergic neurons differentiate from neural progenitors lining the ventricular zone of embryonic ventral midbrain. The development of neural progenitors is controlled by gene expression programs6,7. Here we report techniques utilizing electroporation to express genes specifically in the midbrain of Hamburger Hamilton (HH) stage 11 (thirteen somites, 42 hours) chick embryos8,9. The external development of chick embryos allows for convenient experimental manipulations at specific embryonic stages, with the effects determined at later developmental time points10-13. Chick embryonic neural tubes earlier than HH stage 13 (nineteen somites, 48 hours) consist of multipotent neural progenitors that are capable of differentiating into distinct cell types of the nervous system. The pCAG vector, which contains both a CMV promoter and a chick &beta;-actin enhancer, allows for robust expression of Flag or other epitope-tagged constructs in embryonic chick neural tubes14. In this report, we emphasize special measures to achieve regionally restricted gene expression in embryonic midbrain dopaminergic neuron progenitors, including how to inject DNA constructs specifically into the embryonic midbrain region and how to pinpoint electroporation with small custom-made electrodes. Analyzing chick midbrain at later stages provides an excellent in vivo system for plasmid vector-mediated gain-of-function and loss-of-function studies of midbrain development. Modification of the experimental system may extend the assay to other parts of the nervous system for performing fate mapping analysis and for investigating the regulation of gene expression.

In ovo Electroporation in Chick Midbrain for Studying Gene Function in Dopaminergic Neuron Development

To assess the function and the regulation of genes during the development of midbrain dopaminergic neurons, we describe a method that involves in ovo electroporation of plasmid DNA constructs into embryonic chick ventral midbrain dopaminergic neuron progenitors. This technique can be used to achieve efficient expression of genes of interest to study different aspects of midbrain development and dopaminergic neuron differentiation.

Dopaminergic neurons located in the ventral midbrain control movement, emotional behavior, and reward mechanisms1-3. The dysfunction of ventral midbrain ...

Production of Lentiviral Vectors for Transducing Cells from the Central Nervous System

Efficient gene delivery in the central nervous system (CNS) is important in studying gene functions, modeling neurological diseases and developing therapeutic approaches. Lentiviral vectors are attractive tools in transduction of neurons and other cell types in CNS as they transduce both dividing and non-dividing cells, support sustained expression of transgenes, and have relatively large packaging capacity and low toxicity 1-3. Lentiviral vectors have been successfully used in transducing many neural cell types in vitro 4-6 and in animals 7-10.
Great efforts have been made to develop lentiviral vectors with improved biosafety and efficiency for gene delivery. The current third generation replication-defective and self-inactivating (SIN) lentiviral vectors are depicted in Figure 1. The required elements for vector packaging are split into four plasmids. In the lentiviral transfer plasmid, the U3 region in the 5' long terminal repeat (LTR) is replaced with a strong promoter from another virus. This modification allows the transcription of the vector sequence independent of HIV-1 Tat protein that is normally required for HIV gene expression 11. The packaging signal (&Psi;) is essential for encapsidation and the Rev-responsive element (RRE) is required for producing high titer vectors. The central polypurine tract (cPPT) is important for nuclear import of the vector DNA, a feature required for transducing non-dividing cells 12. In the 3' LTR, the cis-regulatory sequences are completely removed from the U3 region. This deletion is copied to 5' LTR after reverse transcription, resulting in transcriptional inactivation of both LTRs. Plasmid pMDLg/pRRE contains HIV-1 gag/pol genes, which provide structural proteins and reverse transcriptase. pRSV-Rev encodes Rev which binds to the RRE for efficient RNA export from the nucleus. pCMV-G encodes the vesicular stomatitis virus glycoprotein (VSV-G) that replaces HIV-1 Env. VSV-G expands the tropism of the vectors and allows concentration via ultracentrifugation 13. All the genes encoding the accessory proteins, including Vif, Vpr, Vpu, and Nef are excluded in the packaging system. The production and manipulation of lentiviral vectors should be carried out according to NIH guidelines for research involving recombinant DNA (<a href="http://oba.od.nih.gov/oba/rac/Guidelines/NIH_Guidelines.pdf" target="_blank">http://oba.od.nih.gov/oba/rac/Guidelines/NIH_Guidelines.pdf</a>). An approval from individual Institutional Biological and Chemical Safety Committee may be required before using lentiviral vectors. Lentiviral vectors are commonly produced by cotransfection of 293T cells with lentiviral transfer plasmid and the helper plasmids encoding the proteins required for vector packaging. Many lentiviral transfer plasmids and helper plasmids can be obtained from Addgene, a non-profit plasmid repository (<a href="http://www.addgene.org/" target="_blank">http://www.addgene.org/</a>). Some stable packaging cell lines have been developed, but these systems provide less flexibility and their packaging efficiency generally declines over time 14, 15. Commercially available transfection kits may support high efficiency of transfection 16, but they can be very expensive for large scale vector preparations. Calcium phosphate precipitation methods provide highly efficient transfection of 293T cells and thus provide a reliable and cost effective approach for lentiviral vector production.
In this protocol, we produce lentiviral vectors by cotransfection of 293T cells with four plasmids based on the calcium phosphate precipitation principle, followed by purification and concentration with ultracentrifugation through a 20% sucrose cushion. The vector titers are determined by fluorescence- activated cell sorting (FACS) analysis or by real time qPCR. The production and titration of lentiviral vectors in this protocol can be finished with 9 days. We provide an example of transducing these vectors into murine neocortical cultures containing both neurons and astrocytes. We demonstrate that lentiviral vectors support high efficiency of transduction and cell type-specific gene expression in primary cultured cells from CNS.

In this protocol we describe production, purification and titration of lentiviral vectors. We provide an example of lentiviral vector-mediated gene delivery in primary cultured neurons and astrocytes. Our methods may also apply to other cell types in vitro and in vivo.

Efficient gene delivery in the central nervous system (CNS) is important in studying gene functions, modeling neurological diseases and developing ...

A Lightweight, Headphones-based System for Manipulating Auditory Feedback in Songbirds

Experimental manipulations of sensory feedback during complex behavior have provided valuable insights into the computations underlying motor control and sensorimotor plasticity1. Consistent sensory perturbations result in compensatory changes in motor output, reflecting changes in feedforward motor control that reduce the experienced feedback error. By quantifying how different sensory feedback errors affect human behavior, prior studies have explored how visual signals are used to recalibrate arm movements2,3 and auditory feedback is used to modify speech production4-7. The strength of this approach rests on the ability to mimic naturalistic errors in behavior, allowing the experimenter to observe how experienced errors in production are used to recalibrate motor output.
Songbirds provide an excellent animal model for investigating the neural basis of sensorimotor control and plasticity8,9. The songbird brain provides a well-defined circuit in which the areas necessary for song learning are spatially separated from those required for song production, and neural recording and lesion studies have made significant advances in understanding how different brain areas contribute to vocal behavior9-12. However, the lack of a naturalistic error-correction paradigm - in which a known acoustic parameter is perturbed by the experimenter and then corrected by the songbird - has made it difficult to understand the computations underlying vocal learning or how different elements of the neural circuit contribute to the correction of vocal errors13.
The technique described here gives the experimenter precise control over auditory feedback errors in singing birds, allowing the introduction of arbitrary sensory errors that can be used to drive vocal learning. Online sound-processing equipment is used to introduce a known perturbation to the acoustics of song, and a miniaturized headphones apparatus is used to replace a songbird's natural auditory feedback with the perturbed signal in real time. We have used this paradigm to perturb the fundamental frequency (pitch) of auditory feedback in adult songbirds, providing the first demonstration that adult birds maintain vocal performance using error correction14. The present protocol can be used to implement a wide range of sensory feedback perturbations (including but not limited to pitch shifts) to investigate the computational and neurophysiological basis of vocal learning.

We describe the design and assembly of miniaturized headphones suitable for replacing a songbird&#x2019;s natural auditory feedback with a manipulated acoustic signal. Online sound processing hardware is used to manipulate song output, introduce real-time errors in auditory feedback via the headphones, and drive vocal motor learning.

Experimental manipulations of sensory feedback during complex behavior have provided valuable insights into the computations underlying motor control and ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

Immunohistochemical Analysis in the Rat Central Nervous System and Peripheral Lymph Node Tissue Sections

Immunohistochemistry (IHC) provides highly specific, reliable and attractive protein visualization. Correct performance and interpretation of an IHC-based multicolor labeling is challenging, especially when utilized for assessing interrelations between target proteins in the tissue with a high fat content such as the central nervous system (CNS).
Our protocol represents a refinement of the standard immunolabeling technique particularly adjusted for detection of both structural and soluble proteins in the rat CNS and peripheral lymph nodes (LN) affected by neuroinflammation. Nonetheless, with or without further modifications, our protocol could likely be used for detection of other related protein targets, even in other organs and species than here presented.

We here present an optimized, detailed protocol for double immunostaining in formalin-fixed, paraffin-embedded rat central nervous system (CNS) and peripheral lymph node (LN) tissue sections.

Immunohistochemistry (IHC) provides highly specific, reliable and attractive protein visualization. Correct performance and interpretation of an IHC-based ...

Laser Nanosurgery of Cerebellar Axons In Vivo

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

Laser Nanosurgery of Cerebellar Axons In Vivo

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

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

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...