This work was supported by the National Institute of Diabetes and Digestive and Kidney Diseases (NIDDK) grants F31-DK-113673 (C.L.F.), T32-GM-095421 (C.L.F.), DK-089056 (G.J.M.), an American Diabetes Association Innovative Basic Science Award (#1-19-IBS-192 to G.J.M.) and the NIDDK-funded Nutrition Obesity Research Center (DK-035816), Diabetes Research Center (DK-017047) and Diabetes, Obesity and Metabolism Training Grant T32 DK0007247 (T.H.M) at the University of Washington.

All procedures were approved in accordance with the National Institutes of Health, the Guide for the Care and Use of Animals and were approved by both the Institutional Animal Care and Use Committee (IACUC) and Environmental Health and Safety at the University of Washington.
1. Calculation of angled coordinates
<ol>
	<li>Using a coronal brain atlas, mark a right triangle so that the hypotenuse passes through the target region of interest. In the representative example (Figure 1), the hypothalamic ventromedial nucleus (VMN) is targeted at a 15&#176; angle from the coronal midline. 
	NOTE: The placement of the axis of rotation depicted in Figure 1 (and thus, the length of side C) is arbitrary and can be modified to target any brain region. Although this may seem counterintuitive, later steps in the protocol will adjust the position of the head in the z-axis such that this point aligns with the stereotactic center of rotation (see section 6). However, it is recommended not to exceed a coronal rotation angle of 15&#176; due to physical constraints of the head holder apparatus.</li>
	<li>Establish the desired angle (a) and estimated length of side B and use trigonometry to calculate the length of sides A and C. This step is important for properly positioning the head during rotation. 
	NOTE: In the example in Figure 1, atlas gridlines are used to approximate the length of side B, yielding a length of 7.576 mm. This information is used to calculate the length of side A: 
	<img alt="Equation 1" src="/files/ftp_upload/60965/60965equ01.jpg" /> 
	In this example, 2.03 mm indicates the R/L distance from the midline at which the fiberoptic cannula enters the brain when the head is rotated by 15&#176;.
	<ol>
		<li>Optionally, calculate the length of side C to approximate the D/V coordinate: 
		<img alt="Equation 2" src="/files/ftp_upload/60965/60965equ02.jpg" /> 
		NOTE: 1) The length of the hypotenuse (C) does not represent the depth of injection but will be helpful in determining the D/V coordinate, which may need to be adjusted to accommodate for the increased length vs. side B for straight-in injections. It is therefore recommended to perform test injections to optimize the D/V coordinate. 2) In this example targeting the VMN, two sets of coordinates are obtained: one for the microinjection that is non-angled (A/P = -1.4, R/L = 0.4 at 0&#176;, D/V = -5.7) and one for the angled fiberoptic implantation (A/P = -1.4, R/L = 0.0 at 15&#176;, D/V = -5.4).</li>
	</ol>
	</li>
</ol>
2. Preparation of the stereotax for angled procedure
<ol>
	<li>Confirm that the stereotactic frame and micromanipulator have been calibrated (see Kopf manual for full protocol).</li>
	<li>Place the center height gauge into the socket of the head holder base plate.</li>
	<li>Secure the centering scope in the tool holder, then sight down the scope. Adjust the position of the micromanipulator until the crosshairs are aligned and focused on the gauge crosshairs. 
	NOTE: During this step, the scope is being positioned into the focal plane of the head holder&#8217;s center of rotation. Once established, the micromanipulator should not be moved during the remaining steps.</li>
	<li>Place the ear bars into the holders and center them such that the indicator lines on both sides are at 0 (Figure 3A).</li>
	<li>Use the medial-lateral and anterior-posterior knobs on the head holder (Figure 2) to center-align the ear bars in the x- and y-planes above the crosshair of the center height gauge (Figure 3A).</li>
	<li>To align the ear bar position in the z-axis, remove the ear bars from the holder and remove the center height gauge. Replace the ear bars and center them again at 0.</li>
	<li>Sight down the scope. Use the vertical shift knob (Figure 3B) and coronal tilt knob, respectively, to lower and rotate the ear bars until the scope crosshairs remain centered between the ear bars throughout coronal rotation.</li>
	<li>The stereotax is now calibrated and ready. Do not make any further adjustments to the position of the head holder.</li>
</ol>
3. Preparation of materials for injection/implantation
<ol>
	<li>Ensure that all instruments, surgical tools, and materials are sterilized and placed in a sterile surgical field next to the stereotax.</li>
	<li>Handle and store viral constructs according to their biosafety level and relevant institutional biosafety regulatory requirements.</li>
	<li>Draw up the virus into the syringe, taking care to use proper handling practices and personal protective equipment.</li>
</ol>
4. Anesthesia
<ol>
	<li>Record the mouse&#8217;s body weight prior to surgery.</li>
	<li>Deeply anesthetize the mouse using isoflurane.</li>
	<li>Ensure that the mouse is deeply anesthetized by performing a toe pinch test until the flinching response is absent. If the animal continues to show strong reflexes, increase the concentration and/or duration of anesthesia.</li>
	<li>Apply eye ointment to each eye to keep them moist during surgery.</li>
	<li>Shave the scalp from just behind the ears to just behind the eyes with a hair clipper.</li>
	<li>Provide the mouse with IACUC-approved analgesic.</li>
	<li>Continuously monitor the animal throughout the surgical procedure and provide thermal support.</li>
</ol>
5. Surgical procedure
<ol>
	<li>Place the head into the head holder by placing the upper incisors into the gap in the bite bar, making sure that the tongue is below the bite bar.</li>
	<li>Secure the head in the ear bars by gently inserting the ear bars into the external auditory meatus, ensuring that the ear bars are symmetrically placed (typically between three and four for an adult mouse). This step is critical to ensure the head is stable and centered for rotation.</li>
	<li>Aseptically prepare&#160;the shaved incision area with three alternating scrubs of betadine and alcohol swabs, or with alternative institutionally approved surgical site preparation.</li>
	<li>Place a surgical drape over the animal to maintain a sterile surgical field and to reduce the risk of post-operative infection.</li>
	<li>Expose the skull by making an incision along the sagittal midline of the scalp. Gently scrape the surface of the skull to remove any fascia and expose the sutures. 
	NOTE: If suture lines are difficult to visualize, hydrogen peroxide can be applied to the skull using a sterile cotton-tipped applicator to improve suture visualization.</li>
	<li>Place the centering scope into the holder and center the crosshairs on bregma (Figure 4, left panel). Zero the micromanipulator.</li>
	<li>Move the crosshairs caudally to lambda, noting the bregma-lambda (B-L) distance. 
	NOTE: If the suture lines do not follow a straight line along the midline, it is recommended to establish the midline using the &#8220;line of best fit&#8221; through both bregma and lambda. However, if the above steps are followed, the initial placement of the scope reticle should be halfway between the ear bars and closely approximate the B-L midline suture.</li>
	<li>If the B-L distance is significantly less or greater than 4.21 mm, incrementally adjust the assigned bregma to obtain a B-L distance of 4.21 mm &#177; 0.2 mm.</li>
	<li>Replace the centering scope with the alignment indicator. Place the probes on lambda and bregma and adjust the dorsal tilt knob on the head holder to level in the sagittal plane (nose facing up or down), then use the centering scope to reassign bregma.</li>
	<li>Use the alignment indicator to level in the coronal plane using the coronal tilt knob. Measure at multiple points throughout the rostral/caudal axis to account for surface deformations in the skull.</li>
	<li>Note the position on the dial of the coronal tilt knob, as this is the 0&#176; rotation position.</li>
</ol>
6. Aligning the central axes of rotation for angled coordinates
<ol>
	<li>Secure the centering scope in the tool holder and position the micromanipulator to the calculated coordinate from section 1. Note that the R/L coordinate for the angled implantation corresponds to the length of side A.
	<ol>
		<li>In the example in Figure 1, the angled coordinates for fiberoptic placement targeting the VMN are (A/P = -1.4, R/L = [2.03] at 0&#176; coronal rotation, [0.00] at 15&#176; coronal rotation, D/V = -5.4).</li>
	</ol>
	</li>
	<li>Sighting down the scope and mark this coordinate (R/L 2.03 mm from the midline per the VMN example; Figure 4, middle panel). This mark represents the point at which the cannula will enter the brain once the head is rotated.</li>
	<li>Reposition the micromanipulator over the midline (R/L = 0.00). Use the coronal tilt knob to rotate the head to the angle calculated in section 1.
	<ol>
		<li>If the scope crosshairs already line up with the mark, proceed to section 7.</li>
		<li>If the scope crosshairs do not line up with the reference mark, adjust the head position in the z-axis using the vertical shift knob (Figure 2) until the crosshairs line up as close as possible to the mark.</li>
	</ol>
	</li>
	<li>Rotate the head back to the 0&#176; coronal position. If the vertical shift was adjusted in step 6.3, reassign bregma using the centering scope.</li>
	<li>Repeat steps 6.3 and 6.4 until the crosshairs consistently hit the reference mark when the head is rotated (Figure 4C).</li>
	<li>At this point, the arbitrary point of rotation established in section 1 should now be aligned with the stereotactic center of rotation.</li>
</ol>
7. Microinjection
<ol>
	<li>Place the stereotactic drill in the holder and maneuver the micromanipulator to the first injection coordinate.
	<ol>
		<li>Per the example for targeting the VMN, drill at A/P = -1.4 and R/L = 0.4 while the head is level.</li>
	</ol>
	</li>
	<li>Lower the drill until the bit is just above the skull. Turn on the drill, and gently lower until the bit has just drilled through the skull (not the dura).</li>
	<li>Repeat for the contralateral injection site.</li>
	<li>Use a sterile needle-driver to introduce a 90&#176; bend into a 27-30G needle (e.g., of a sterile 0.5 mL insulin syringe), and use the bent needle to gently poke through the dura mater.</li>
	<li>NOTE: If bleeding occurs, apply pressure with a sterile cotton-tipped applicator and clean with sterile water until the bleeding has stopped.</li>
	<li>When ready to inject, carefully place a filled Hamilton syringe into the stereotactic holder. 
	NOTE: The coordinates on the micromanipulator no longer apply after switching to a new tool. Use the center of the burr hole as the new target for injection.</li>
	<li>Carefully position the needle above the burr hole.</li>
	<li>Lower the needle until it slightly touches the dura within the center of the burr hole. CRITICAL: Zero the micromanipulator only in the z-axis, such that the coordinates on the micromanipulator for the stereotactic centering scope and drill are maintained.</li>
	<li>Slowly lower the needle into the brain, watching closely to ensure that the needle does not deflect on the edge of the burr hole. Continue to lower until 0.05 mm ventral to the D/V injection coordinate and wait 1 min. This extra step creates a small &#8220;pocket&#8221; to minimize viral backflow on needle removal.</li>
	<li>Slowly raise the needle to the D/V coordinate and start the injection. 
	NOTE: Flow rate and volume will vary depending on target region and experimental design. For optogenetic silencing of VMN neurons, sufficient coverage is desired, so 200 nL of virus is injected at a rate of 1 nL/s.</li>
	<li>Following microinjection, wait 10 min at the injection site to minimize efflux of virus during withdrawal.</li>
	<li>Slowly withdraw the micropipette from the brain at an approximate rate of 1 mm/min.</li>
	<li>Once the needle is clear of the skull, eject a small volume of virus to ensure the needle has not clogged with blood or tissue. Use a sterile cotton-tipped applicator to remove the virus before continuing.</li>
	<li>Repeat steps 7.6&#8211;7.12 for the contralateral side.</li>
	<li>Seal the microinjection burr holes with bone wax to improve healing (Figure 5B).</li>
</ol>
8. Fiberoptic implantation
NOTE: After viral injection, bilateral fiberoptic cannulas are implanted at the calculated angle per section 1. Note that these coordinates should already be marked on the skull from section 6.
<ol>
	<li>Repeat steps 7.1 &#8211;7.4 for the angled coordinates.</li>
	<li>Return the head to the level 0&#176; position.</li>
	<li>Next, use the hand drill to produce four additional holes for the bone screws: two should be placed anteriorly and two posteriorly (Figure 5A). These will serve as anchors to affix the fiberoptics to the skull (Figure 5D). 
	NOTE: Make sure to space the holes far enough away from the angled coordinate burr holes to accommodate the ferrule portion of the fiberoptic that sits above the skull.</li>
	<li>As gently as possible, use the small flathead screwdriver to set the bone screws such that they sit firmly in the skull but do not penetrate the brain.</li>
	<li>Clamp a fiberoptic cannula into the cannula holder and place it in the stereotactic holder.</li>
	<li>Rotate the head to the calculated angle, noting again that the coordinates on the micromanipulator do not apply to the new tool. Use the center of the angled burr holes as the implantation target.</li>
	<li>Lower the fiberoptic until it just touches the dura within the center of the burr hole (Figure 5C). Zero the micromanipulator in the z-axis, then slowly slower the fiberoptic to the D/V angled coordinate (-5.4 per the VMN example).</li>
	<li>Use cyanoacrylate gel to connect the fiberoptic ferrule to the ipsilateral anchor screws, then apply an accelerant with a micropipette tip (Figure 5D).</li>
	<li>Once the cyanoacrylate gel has completely hardened, gently loosen the cannula holder and raise until clear of the fiberoptic ferrule.</li>
	<li>Repeat steps 8.5&#8211;8.9 for the contralateral angled coordinate, then level the head. For extra security, make an additional connection between the two angled fiberoptic cannulas with the cyanoacrylate gel and accelerant (Figure 5D).</li>
	<li>Prepare a small, relatively thin amount of dental cement. Apply to the surface of the skull, making sure to thoroughly cover the anchor screws and base of the fiberoptic cannulas. Leave enough of the ferrule clean for subsequent mating with the fiberoptic patch cables.</li>
	<li>Once the cement is completed dry, remove the mouse from the stereotactic apparatus.</li>
	<li>Place mouse in a recovery cage with thermal support. Allow it to recover and transfer to the home cage once it appears alert, mobile, and is grooming.</li>
</ol>
9. Post-surgical care
<ol>
	<li>Monitor animals daily for 3 days post-operatively for behavior, posture, activity, and grooming, and keep records of food intake and body weight.</li>
	<li>If animals exhibit any general indicators of pain or poor health, consult with veterinarian services.</li>
	<li>Allow mice at least 2 weeks for recovery and for viral expression before starting behavioral studies.</li>
</ol>
10. Optogenetics
<ol>
	<li>For the performance of optogenetics studies, refer to Sidor et al.8.</li>
	<li>Validate viral expression and fiber placement at the completion of studies.</li>
</ol>

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	<li>King, B. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+rise,+fall,+and+resurrection+of+the+ventromedial+hypothalamus+in+the+regulation+of+feeding+behavior+and+body+weight.">The rise, fall, and resurrection of the ventromedial hypothalamus in the regulation of feeding behavior and body weight.</a> Physiology and Behavior. 87, 221-244 (2006).</li><li>Boyden, E. S., Zhang, F., Bamberg, E., Nagel, G., Deisseroth, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Millisecond-timescale,+genetically+targeted+optical+control+of+neural+activity.">Millisecond-timescale, genetically targeted optical control of neural activity.</a> Nature Neuroscience. 8, 1263-1268 (2005).</li><li>Roth, B. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=DREADDs+for+Neuroscientists.">DREADDs for Neuroscientists.</a> Neuron. 89, 683-694 (2016).</li><li>Richevaux, L., Schenberg, L., Beraneck, M., Fricker, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+Vivo+Intracerebral+Stereotaxic+Injections+for+Optogenetic+Stimulation+of+Long-Range+Inputs+in+Mouse+Brain+Slices.">In Vivo Intracerebral Stereotaxic Injections for Optogenetic Stimulation of Long-Range Inputs in Mouse Brain Slices.</a> Journal of Visualized Experiments. , e59534 (2019).</li><li>Fricano-Kugler, C. J., Williams, M. R., Luikart, B., Salinaro, J. R., Li, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Designing,+packaging,+and+delivery+of+high+titer+crispr+retro+and+lentiviruses+via+stereotaxic+injection.">Designing, packaging, and delivery of high titer crispr retro and lentiviruses via stereotaxic injection.</a> Journal of Visualized Experiments. , e53783 (2016).</li><li>McSweeney, C., Mao, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Applying+Stereotactic+Injection+Technique+to+Study+Genetic+Effects+on+Animal+Behaviors.">Applying Stereotactic Injection Technique to Study Genetic Effects on Animal Behaviors.</a> Journal of Visualized Experiments. (99), e52653 (2015).</li><li>Lowell, B. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+Neuroscience+of+Homeostasis+and+Drives+for+Food,+Water,+and+Salt.">New Neuroscience of Homeostasis and Drives for Food, Water, and Salt.</a> New England Journal of Medicine. 380, 459-471 (2019).</li><li>Sidor, M. M., 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=In+vivo+optogenetic+stimulation+of+the+rodent+central+nervous+system.">In vivo optogenetic stimulation of the rodent central nervous system.</a> Journal of Visualized Experiments. , e51483 (2015).</li><li>Faber, C. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+Neuronal+Projections+from+the+Hypothalamic+Ventromedial+Nucleus+Mediate+Glycemic+and+Behavioral+Effects.">Distinct Neuronal Projections from the Hypothalamic Ventromedial Nucleus Mediate Glycemic and Behavioral Effects.</a> Diabetes. 67, 2518-2529 (2018).</li><li>Berndt, A., 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=Structural+foundations+of+optogenetics:+Determinants+of+channelrhodopsin+ion+selectivity.">Structural foundations of optogenetics: Determinants of channelrhodopsin ion selectivity.</a> Proceedings of the National Academy of Scences. 113, 822-829 (2016).</li><li>Faber, C. L., Matsen, M. E., Meek, T. H., Krull, J. E., Morton, G. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+customizable+procedure+for+angled+stereotaxic+implantation+and+microinjection+in+the+rodent+brain.">A customizable procedure for angled stereotaxic implantation and microinjection in the rodent brain.</a> Kopf Carrier. 96, (2019).</li><li>Correia, P., Matias, S., Mainen, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stereotaxic+Adeno-associated+Virus+Injection+and+Cannula+Implantation+in+the+Dorsal+Raphe+Nucleus+of+Mice.">Stereotaxic Adeno-associated Virus Injection and Cannula Implantation in the Dorsal Raphe Nucleus of Mice.</a> Bio-Protocol. 7, 2549 (2017).</li><li>Cardozo Pinto, D. F., Lammel, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hot+topic+in+optogenetics:+new+implications+of+in+vivo+tissue+heating.">Hot topic in optogenetics: new implications of in vivo tissue heating.</a> Nature Neuroscience. 22, 1039-1041 (2019).</li></ol>

The authors have nothing to disclose.

Recent advances in neuroscience have supported advanced insight and understanding into the activity and function of brain neurocircuits. This includes the application of optogenetic and chemogenetic technologies to activate or silence discrete neuronal populations and their projection sites in vivo. More recently, this has included the development of genetically encoded calcium indicators (e.g., GCaMP, RCaMP) and other fluorometric biosensors (e.g., dopamine, norepinephrine) for in vivo recording of neuronal activity in a defined cell type in freely moving animals. However, effective employment of these technologies relies upon successful stereotactic surgery to target the region of interest. While there are several established protocols describing these methods, which are suitable for targeting many brain regions, targeting deep brain regions along the midline represents significant additional challenges. Demonstrated here is a detailed surgical technique for targeting discrete brain regions via an angled stereotactic approach. Importantly, this technique can be adapted and applied to a diverse range of neuroscience techniques (i.e., optogenetics, chemogenetics, and fiber photometry approaches).
Using this approach, it is shown that acute optogenetic silencing of VMN neurons expressing neuronal nitric oxide synthase (VMNNOS1 neurons) blunts glucagon responses to insulin-induced hypoglycemia in mice9. Using a slightly modified approach, it is further demonstrated that unilateral activation of VMNNOS1 neurons 1) elicits robust hyperglycemia that is driven by counterregulatory responses that are normally reserved for the response to hypoglycemia, and 2) elicits defensive immobility behavior. Furthermore, these behavioral and metabolic responses involve neuronal projections to distinct brain areas. Specifically, the activation of VMNNOS1 neurons projecting to the anterior bed nucleus of the stria terminalis are involved in glycemic responses, whereas VMNNOS1 neurons projecting to the periaqueductal gray are linked to fear-induced behavior responses9.
It should be noted that the protocol is highly specific to the Kopf Model 1900 stereotax and accompanying accessories. While this system enables precise, reproducible implantation as well as microinjection to discrete brain regions (with a common centerline position across multiple tools), the strategy and approach can be adapted to suit other stereotaxic frames. Specifically, instead of rotating the head to perform angled microinjections and implantations, an alternative approach is to utilize the same principles and rotate the dorsal-ventral manipulator instead (see Correia et al.12).
As with any new method, it is critical for individuals to optimize the technique to improve an experiment&#8217;s reliability, consistency, and accuracy. In addition, it is important to include the necessary appropriate controls for proper analysis and interpretation of data. These include the use of Cre-negative littermate controls, viral reporter controls (i.e., AAV-GFP), verification of light-dependent neuronal firing modulation using electrophysiology, and (upon study completion) the validation of viral targeting and fiberoptic placement in the region of interest. It is recommended to refer to the publication by Cardozo and Lammel13 for a detailed review of technical considerations and suggested controls.
In summary, the introduction of more advanced and precise neuroscience techniques has supported a significant advancement and understanding of the role of the brain in behavior, cognition, and physiology, and these advancements may lead to potential therapies for CNS-related disorders.

Neural control of behavior, feeding, and metabolism involves coordination of highly complex, integrative, and redundant neurocircuits. A driving goal of the neuroscience field is to dissect the relationship between neuronal circuit structure and function. Although classical neuroscience tools (i.e., lesioning, local pharmacological injections, and electrical stimulation) have uncovered vital knowledge regarding the role of specific brain regions that control behavior and metabolism, these tools are limited by their lack of specificity and reversibility1.
Recent advances in the neuroscience field have greatly improved the ability to interrogate and manipulate circuit function in a cell-type specific manner with high spatiotemporal resolution. Optogenetic2 and chemogenetic3 approaches, for instance, allow the rapid and reversible manipulation of activity in genetically defined cell types of freely moving animals. Optogenetics involves the use of light-sensitive ion channels, termed channelrhodopsins, to control neuronal activity. Key to this technique is the gene delivery of channelrhodopsin and a source of light to activate the opsin. A common strategy for gene delivery is through a combination of 1) genetically engineered mice expressing Cre-recombinase in discrete neurons, and 2) Cre-dependent viral vectors encoding channelrhodopsin.
While optogenetics provides an elegant, highly precise means to control neuronal activity, the method is contingent upon successful stereotactic microinjection of the viral vector and fiberoptic placement into a defined brain region. Although stereotactic procedures are commonplace within the modern neuroscience lab (and there are several excellent protocols describing this procedure)4,5,6, being able to consistently and reproducibly target discrete brain regions along the midline (i.e., the mediobasal hypothalamus, a brain area critical to the regulation of homeostatic functions7) presents additional challenges. These challenges include avoiding of the superior sagittal sinus, third ventricle, and adjacent hypothalamic nuclei. In addition, there are significant spatial limitations to the bilateral implantation of hardware that is required for inhibition studies. With these challenges in mind, this protocol herein presents a modifiable procedure for targeting discrete brain regions via an angled stereotactic approach.

<table>
	<tbody>
		<tr>
			<td>Fiberoptic Cannulae</td>
			<td>Doric Lenses</td>
			<td>MFC_200/230-0.57_###_MF1.25_FLT</td>
			<td>Customizable</td>
		</tr>
		<tr>
			<td>Kopf Model 1900 Stereotaxic Alignment System</td>
			<td>Kopf</td>
			<td>Model 1900</td>
			<td></td>
		</tr>
		<tr>
			<td>Kopf Model 1900-51 Center Height Gauge</td>
			<td>Kopf</td>
			<td>Model 1900-51</td>
			<td></td>
		</tr>
		<tr>
			<td>Kopf Model 1905 Alignment Indicator</td>
			<td>Kopf</td>
			<td>Model 1905</td>
			<td></td>
		</tr>
		<tr>
			<td>Kopf Model 1911 Stereotaxic Drill</td>
			<td>Kopf</td>
			<td>Model 1911</td>
			<td></td>
		</tr>
		<tr>
			<td>Kopf Model 1915 Centering Scope</td>
			<td>Kopf</td>
			<td>Model 1915</td>
			<td></td>
		</tr>
		<tr>
			<td>Kopf Model 1922 60-Degree Non-Rupture Ear Bars</td>
			<td>Kopf</td>
			<td>Model 1922</td>
			<td></td>
		</tr>
		<tr>
			<td>Kopf Model 1923-B Mouse Gas Anesthesia Head Holder</td>
			<td>Kopf</td>
			<td>Model 1923-B</td>
			<td></td>
		</tr>
		<tr>
			<td>Kopf Model 1940 Micro Manipulator</td>
			<td>Kopf</td>
			<td>Model 1940</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro4 Microinjection System</td>
			<td>World Precision Instruments</td>
			<td>--</td>
			<td></td>
		</tr>
		<tr>
			<td>Mouse bone screws</td>
			<td>Plastics One</td>
			<td>00-96 X 1/16</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotaxic Cannula Holder, 1.25mm ferrule</td>
			<td>Thor Labs</td>
			<td>XCL</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical Drill</td>
			<td>Cell Point Scientific</td>
			<td>Ideal Micro Drill</td>
			<td></td>
		</tr>
	</tbody>
</table>

adaptable angled stereotactic approach for versatile neuroscience techniques

Stereotactic surgery is an essential tool in the modern neuroscience lab. However, the ability to precisely and accurately target difficult-to-reach brain regions still presents a challenge, particularly when targeting brain structures along the midline. These challenges include avoiding of the superior sagittal sinus and third ventricle and the ability to consistently target selective and discrete brain nuclei. In addition, more advanced neuroscience techniques (e.g., optogenetics, fiber photometry, and two-photon imaging) rely on targeted implantation of significant hardware to the brain, and spatial limitations are a common hindrance. Presented here is a modifiable protocol for stereotactic targeting of rodent brain structures using an angled coronal approach. It can be adapted to 1) mouse or rat models, 2) various neuroscience techniques, and 3) multiple brain regions. As a representative example, it includes the calculation of stereotactic coordinates for targeting of the mouse hypothalamic ventromedial nucleus (VMN) for an optogenetic inhibition experiment. This procedure begins with the bilateral microinjection of an adeno-associated virus (AAV) encoding a light-sensitive chloride channel (SwiChR++) to a Cre-dependent mouse model, followed by the angled bilateral implantation of fiberoptic cannulae. Using this approach, findings show that activation of a subset of VMN neurons is required for intact glucose counterregulatory responses to insulin-induced hypoglycemia.

This protocol describes a surgical procedure for performing optogenetics studies to interrogate the role of hypothalamic VMN neurons in glycemic control9. First utilized was a standard (non-angled) stereotactic approach for the bilateral microinjection of an inhibitory channelrhodopsin virus to the VMN. While an angled approach would also be suitable, the standard (non-angled) approach was selected because it is sufficient to target the brain region of interest and is an easy, reliable and consistent approach. However, given the VMN&#8217;s proximity to the midline, space constraints did not permit the non-angled implantation of bilateral fiberoptics, necessitating the development of a surgical strategy for precisely implanting fiberoptics at an angle (Figure 6).
Using this surgical strategy, we microinjected a Cre-dependent AAV expressing a modified channelrhodopsin anion-conducting channel fused with the fluorescent reporter, referred to as a &#8221;SwiChR++&#8221; virus10, bilaterally to the VMN of Nos1-cre mice. This was followed by implantation of an optic fiber dorsolateral to each injection site at a 15&#176; angle from the midline. As expected, viral expression was restricted to the VMN and not detected in other brain areas.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/60965/60965fig01.jpg" /> 
Figure 1: Representative example of calculating angled coordinates targeting the hypothalamic ventromedial nucleus.&#160;Angles and line segments are not drawn to scale. (A) This length should be calculated using basic trigonometry. In this example, A = 2.03 mm. (B) Estimated length based upon assignment of arbitrary axis of rotation. In this example, B = 7.576 mm. (C) Calculated hypotenuse. It should be noted that the depth of fiberoptic/needle insertion depends upon the desired proximity to the target region, which requires optimization. This figure has been modified from Faber et al. 201911. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60965/60965fig1v2large.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/60965/60965fig02.jpg" /> 
Figure 2: Adjustment knobs for the stereotactic head holder apparatus.&#160;This figure has been modified from Faber et al. 201911. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60965/60965fig2v2large.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/60965/60965fig03.jpg" /> 
Figure 3: Aligning the head holder center of rotation.&#160;(A) Positioning the ear bars. (B) Sighting down the scope during 0&#176; level coronal rotation (left), during 15&#176; rotation before adjusting the vertical shift, and the center of rotation is misaligned (middle), and during 15&#176; rotation after adjusting the vertical shift, and the center of rotation is properly aligned (right). This figure has been modified from Faber et al. 201911. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60965/60965fig03large.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/60965/60965fig04.jpg" /> 
Figure 4: Assigning bregma and aligning the animal head with central axes of rotation.&#160;(A) Representative image indicating typical bregma placement. (B) Drawing a reference mark while head is level, before alignment. (C) Properly aligned axis of rotation, after adjusting the vertical shift and readjusting bregma. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60965/60965fig4v2large.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/60965/60965fig05.jpg" /> 
Figure 5: Fiberoptic implantation procedure.&#160;(A) Centering scope view of pilot holes for microinjection (m), fiberoptic (f), and anchor screws (*). (B) Centering scope view of implanted anchor screws, and bone wax covered microinjection drill holes. (C) Positioning the fiberoptic into place during angled implantation. (D) Representative bilateral angled fiberoptic placement. Dotted black arrows indicate areas in which super glue is used to anchor the fiberoptic to the anchor screws and ipsilateral fiberoptic. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60965/60965fig5v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/60965/60965fig06.jpg" /> 
Figure 6: Representative results for bilateral targeting of the ventromedial hypothalamus.&#160;(A) Schematic representing bilateral microinjection and angled fiberoptic strategy for targeting the VMN. (B) Representative image showing bilateral expression of SwiChR-GFP and tissue damage from angled fiberoptic tracts. 3V = third ventricle, ARC = arcuate nucleus, and VMN = ventromedial nucleus. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/60965/60965fig6v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Adaptable Angled Stereotactic Approach for Versatile Neuroscience Techniques at JoVE.com

Adaptable Angled Stereotactic Approach for Versatile Neuroscience Techniques

Described here is a stereotactic procedure that can target challenging and difficult-to-reach brain regions (due to spatial limitations) using an angled coronal approach. This protocol is adaptable to both mouse and rat models and can be applied to diverse neuroscientific applications, including cannula implantation and microinjections of viral constructs.

Stereotactic surgery is an essential tool in the modern neuroscience lab. However, the ability to precisely and accurately target difficult-to-reach brain ...

adaptable-angled-stereotactic-approach-for-versatile-neuroscience

Nanyang Technological University

Research

JoVE Journal

Neuroscience

4.9K Views.  University of Washington. Described here is a stereotactic procedure that can target challenging and difficult-to-reach brain regions (due to spatial limitations) using an angled coronal approach. This protocol is adaptable to both mouse and rat models and can be applied to diverse neuroscientific applications, including cannula implantation and microinjections of viral constructs.

Video: Adaptable Angled Stereotactic Approach for Versatile Neuroscience Techniques

A Novel Approach for Documenting Phosphenes Induced by Transcranial Magnetic Stimulation

Stimulation of the human visual cortex produces a transient perception of light, known as a phosphene. Phosphenes are induced by invasive electrical stimulation of the occipital cortex, but also by non-invasive Transcranial Magnetic Stimulation (TMS)1 of the same cortical regions. The intensity at which a phosphene is induced (phosphene threshold) is a well established measure of visual cortical excitability and is used to study cortico-cortical interactions, functional organization 2, susceptibility to pathology 3,4 and visual processing 5-7. Phosphenes are typically defined by three characteristics: they are observed in the visual hemifield contralateral to stimulation; they are induced when the subject s eyes are open or closed, and their spatial location changes with the direction of gaze 2. Various methods have been used to document phosphenes, but a standardized methodology is lacking. We demonstrate a reliable procedure to obtain phosphene threshold values and introduce a novel system for the documentation and analysis of phosphenes. We developed the Laser Tracking and Painting system (LTaP), a low cost, easily built and operated system that records the location and size of perceived phosphenes in real-time. The LTaP system provides a stable and customizable environment for quantification and analysis of phosphenes.

Phosphenes are transient percepts of light that can be induced by applying Transcranial Magnetic Stimulation (TMS) to visually sensitive regions of cortex. We demonstrate a standard protocol for determining the phosphene threshold value and introduce a novel method for quantifying and analyzing perceived phosphenes.

Stimulation of the human visual cortex produces a transient perception of light, known as a phosphene. Phosphenes are induced by invasive electrical ...

High Density Event-related Potential  Data Acquisition in Cognitive Neuroscience

Functional magnetic resonance imaging (fMRI) is currently the standard method of evaluating brain function in the field of Cognitive Neuroscience, in part because fMRI data acquisition and analysis techniques are readily available. Because fMRI has excellent spatial resolution but poor temporal resolution, this method can only be used to identify the spatial location of brain activity associated with a given cognitive process (and reveals virtually nothing about the time course of brain activity). By contrast, event-related potential (ERP) recording, a method that is used much less frequently than fMRI, has excellent temporal resolution and thus can track rapid temporal modulations in neural activity. Unfortunately, ERPs are under utilized in Cognitive Neuroscience because data acquisition techniques are not readily available and low density ERP recording has poor spatial resolution. In an effort to foster the increased use of ERPs in Cognitive Neuroscience, the present article details key techniques involved in high density ERP data acquisition. Critically, high density ERPs offer the promise of excellent temporal resolution and good spatial resolution (or excellent spatial resolution if coupled with fMRI), which is necessary to capture the spatial-temporal dynamics of human brain function.

Event-related potential (ERP) recording is under utilized in Cognitive Neuroscience because data acquisition techniques are not readily available and this method often has poor spatial resolution. To foster the increased use of ERPs in Cognitive Neuroscience, the present article details key techniques involved in high density ERP data acquisition.

Functional magnetic resonance imaging (fMRI) is currently the standard method of evaluating brain function in the field of Cognitive Neuroscience, in part ...

Rodent Stereotaxic Surgery and Animal Welfare Outcome Improvements for Behavioral Neuroscience

Stereotaxic surgery for the implantation of cannulae into specific brain regions has for many decades been a very successful experimental technique to investigate the effects of locally manipulated neurotransmitter and signaling pathways in awake, behaving animals. Moreover, the stereotaxic implantation of electrodes for electrophysiological stimulation and recording studies has been instrumental to our current understanding of neuroplasticity and brain networks in behaving animals. Ever-increasing knowledge about optimizing surgical techniques in rodents1-4, public awareness concerning animal welfare issues and stringent legislation (e.g., the 2010 European Union Directive on the use of laboratory animals5) prompted us to refine these surgical procedures, particularly with respect to implementing new procedures for oxygen supplementation and the continuous monitoring of blood oxygenation and heart rate levels during the surgery as well as introducing a standardized protocol for post-surgical care. Our observations indicate that these modifications resulted in an increased survival rate and an improvement in the general condition of the animals after surgery (e.g. less weight loss and a more active animal). This video presentation will show the general procedures involved in this type of stereotaxic surgery with special attention to our several modifications. We will illustrate these surgical procedures in rats, but it is also possible to perform this type of surgery in mice or other small laboratory animals by using special adaptors for the stereotaxic apparatus6.

Stereotaxic surgery on rodents allows for targeted administration of drugs or electrical stimulation and recordings in awake, behaving animals. In this video presentation we will demonstrate recent procedural refinements to this long-standing procedure that successfully improved survival rate and reduced post-surgical weight loss.

Stereotaxic surgery for the implantation of cannulae into specific brain regions has for many decades been a very successful experimental technique to ...

VisioTracker, an Innovative Automated Approach to Oculomotor Analysis

Investigations into the visual system development and function necessitate quantifiable behavioral models of visual performance that are easy to elicit, robust, and simple to manipulate. A suitable model has been found in the optokinetic response (OKR), a reflexive behavior present in all vertebrates due to its high selection value. The OKR involves slow stimulus-following movements of eyes alternated with rapid resetting saccades. The measurement of this behavior is easily carried out in zebrafish larvae, due to its early and stable onset (fully developed after 96 hours post fertilization (hpf)), and benefitting from the thorough knowledge about zebrafish genetics, for decades one of the favored model organisms in this field. Meanwhile the analysis of similar mechanisms in adult fish has gained importance, particularly for pharmacological and toxicological applications.
Here we describe VisioTracker, a fully automated, high-throughput system for quantitative analysis of visual performance. The system is based on research carried out in the group of Prof. Stephan Neuhauss and was re-designed by TSE Systems. It consists of an immobilizing device for small fish monitored by a high-quality video camera equipped with a high-resolution zoom lens. The fish container is surrounded by a drum screen, upon which computer-generated stimulus patterns can be projected. Eye movements are recorded and automatically analyzed by the VisioTracker software package in real time.
Data analysis enables immediate recognition of parameters such as slow and fast phase duration, movement cycle frequency, slow-phase gain, visual acuity, and contrast sensitivity.
Typical results allow for example the rapid identification of visual system mutants that show no apparent alteration in wild type morphology, or the determination of quantitative effects of pharmacological or toxic and mutagenic agents on visual system performance.

The VisioTracker is an automated system for the quantitative analysis of visual performance of larval and small adult fish based on the recording of eye movements. It features full control over visual stimulus properties and real-time analysis, enabling high-throughput research in fields such as visual system development and function, pharmacology, neural circuit studies and sensorimotor integration.

Investigations into the visual system development and function necessitate quantifiable behavioral models of visual performance that are easy to elicit, ...

A Fully Automated and Highly Versatile System for Testing Multi-cognitive Functions and Recording Neuronal Activities in Rodents

We have developed a fully automated system for operant behavior testing and neuronal activity recording by which multiple cognitive brain functions can be investigated in a single task sequence. The unique feature of this system is a custom-made, acoustically transparent chamber that eliminates many of the issues associated with auditory cue control in most commercially available chambers. The ease with which operant devices can be added or replaced makes this system quite versatile, allowing for the implementation of a variety of auditory, visual, and olfactory behavioral tasks. Automation of the system allows fine temporal (10 ms) control and precise time-stamping of each event in a predesigned behavioral sequence. When combined with a multi-channel electrophysiology recording system, multiple cognitive brain functions, such as motivation, attention, decision-making, patience, and rewards, can be examined sequentially or independently.

In this report, we present a fully automated and highly versatile system capable of simultaneously testing multi-cognitive behaviors and recording neuronal activities for rodents.

We have developed a fully automated system for operant behavior testing and neuronal activity recording by which multiple cognitive brain functions can be ...

A Simple Alternative to Stereotactic Injection for Brain Specific Knockdown of miRNA

MicroRNAs (miRNAs) are key regulators of gene expression. In the brain, vital processes like neurodevelopment and neuronal functions depend on the correct expression of microRNAs. Perturbation of microRNAs in the brain can be used to model neurodegenerative diseases by modulating neuronal cell death. Currently, stereotactic injection is used to deliver miRNA knockdown agents to specific location in the brain. Here, we discuss strategies to design antagomirs against miRNA with locked nucleotide modifications (LNA). Subsequently describe a method for brain specific delivery of antagomirs, uniformly across different regions of the brain. This method is simple and widely applicable since it overcomes the surgery, associated injury and limitation of local delivery in stereotactic injections. We prepared a complex of neurotropic, cell-penetrating peptide Rabies Virus Glycoprotein (RVG) with antagomir against miRNA-29 and injected through tail vein, to specifically deliver in the brain. The antagomir design incorporated features that allow specific targeting of the miRNA and formation of non-covalent complexes with the peptide. The knock-down of the miRNA in neuronal cells, resulted in apoptotic cell death and associated behavioural defects. Thus, the method can be used for acute models of neuro-degeneration through the perturbation of miRNAs.

MicroRNAs play crucial roles in the brain and are potential targets for modeling neuro-degeneration. However, perturbing miRNA levels is challenging due to the short length of miRNA and inaccessibility of the brain tissue. This video presents a method for antagomir design and brain specific delivery using a neuropeptide in mice.

MicroRNAs (miRNAs) are key regulators of gene expression.  In the brain, vital processes like neurodevelopment and neuronal functions depend on the ...

A New Approach that Eliminates Handling for Studying Aggression and the "Loser" Effect in Drosophila melanogaster

Aggressive behavior in Drosophila melanogaster is composed of the sequential expression of stereotypical behavioral patterns (for analysis see 1). This complex behavior is influenced by genetic, hormonal and environmental factors. As in many organisms, previous fighting experience influences the fighting strategy of flies and the outcome of later contests: losing a fight increases the probability of losing later contests, revealing &#34;loser&#34; effects that likely involve learning and memory 2-4. The learning and memory that accompanies expression of complex social behaviors like aggression, is sensitive to pre-test handling of animals 5,6. Many experimental procedures are used in different laboratories to study aggression 7-9, however, no routinely used protocol that excludes handling of flies is currently available. Here, we report a new behavioral apparatus that eliminates handling of flies, using instead their innate negative geotactic responses to move animals into or out of fighting chambers. In this protocol, small circular fight arenas containing a food cup are divided into two equal halves by a removable plastic slider prior to introduction of flies. Flies enter chambers from their home isolation vials via sliding chamber doors and geotaxis. Upon removal of plastic sliders, flies are free to interact. After specified time periods, flies are separated again by sliders for subsequent experimentation. All of this is done easily without handling of individual flies. This apparatus offers a novel approach to study aggression and the associated learning and memory, including the formation of &#34;loser&#34; effects in fly fights. In addition, this new general-purpose behavioral apparatus can be employed to study other social behaviors of flies and should, in general, be of interest for investigating experience-related changes in fundamental behavioral processes.

A New Approach that Eliminates Handling for Studying Aggression and the "Loser" Effect in Drosophila melanogaster

During fruit fly fights, the behavioral patterns observed, fight dynamics, and associated learning and memory are influenced by experimental conditions. The protocol presented here describes a novel procedure that entirely eliminates handling of flies during experiments. This improves fight dynamics and allows formation of strong &#34;loser&#34; effects.

Aggressive behavior in Drosophila melanogaster is composed of the sequential expression of stereotypical behavioral patterns (for analysis see 1). This ...

Endoscopic Endonasal Trans-sphenoidal Approach: Minimally Invasive Surgery for Pituitary Adenomas

Endoscopic endonasal trans-sphenoidal surgery has become the gold standard for the surgical treatment of pituitary adenomas and many other pituitary lesions. Refinements in surgical techniques, technological advancements, and incorporation of neuronavigation have rendered this surgery minimally invasive. The complication rates of this surgery are very low while excellent results are consistently obtained through this approach.
This paper focuses on the step-by-step surgical approach to pituitary adenomas, which is based on personal experience, and details the results obtained with this minimally invasive surgery.

The aim of this paper is to describe the different steps of the endoscopic endonasal approach to the sella turcica.

Endoscopic endonasal trans-sphenoidal surgery has become the gold standard for the surgical treatment of pituitary adenomas and many other pituitary ...

Stereotactic Atlas-Guided Laser Capture Microdissection of Brain Regions Affected by Traumatic Injury

The ability to isolate specific brain regions of interest can be impeded in tissue disassociation techniques that do not preserve their spatial distribution. Such techniques also potentially skew gene expression analysis because the process itself can alter expression patterns in individual cells. Here we describe a laser capture microdissection (LCM) method to selectively collect specific brain regions affected by traumatic brain injury (TBI) by using a modified Nissl (cresyl violet) staining protocol and the guidance of a rat brain atlas. LCM provides access to brain regions in their native positions and the ability to use anatomical landmarks for identification of each specific region. To this end, LCM has been used previously to examine brain region specific gene expression in TBI. This protocol allows examination of TBI-induced alterations in gene and microRNA expression in distinct brain areas within the same animal. The principles of this protocol can be amended and applied to a wide range of studies examining genomic expression in other disease and/or animal models.

We describe the use of laser capture microdissection to obtain samples of distinct cell populations from different brain regions for gene and microRNA analysis. This technique allows the study of differential effects of traumatic brain injury in specific regions of the rat brain.

The ability to isolate specific brain regions of interest can be impeded in tissue disassociation techniques that do not preserve their spatial ...

A Simple Approach to Induce Experimental Autoimmune Neuritis in C57BL/6 Mice for Functional and Neuropathological Assessments

Experimental autoimmune neuritis (EAN) is a well-appreciated experimental model of autoimmune peripheral demyelinating diseases. EAN disease is induced by immunizing mice with neurogenic peptides to direct an inflammatory attack toward components of the peripheral nervous system (PNS). Recent advances have enabled the induction of EAN in the relatively resistant C57BL/6 mouse line using myelin protein zero (P0)106-125 or P0180-199 peptides delivered in adjuvant combined with the injection of pertussis toxin. The ability to induce EAN in the C57BL/6 strain allows for the use of the numerous genetic tools that exist on this mouse background, and thus allows the sophisticated study of disease pathogenesis and interrogation of the mechanistic action of novel therapeutics in combination with transgenic approaches. In this study, we demonstrate a simple approach to successfully induce EAN using the P0180-199 peptide in C57BL/6 mice. We also outline a protocol for the assessment of functional deficits that occur in this model, accompanied by an array of neuropathological features. Thus, this model is a powerful experimental model to study the pathogenesis of human peripheral demyelinating neuropathies, and to determine the efficacy of potential therapies that aim to promote myelin repair and protect against nerve damage in autoimmune neuritis.

This report outlines a simple approach to successfully induce experimental autoimmune neuritis (EAN) using the myelin protein zero (P0)180-199 peptide in combination with Freund&#39;s complete adjuvant and pertussis toxin. We present a sophisticated paradigm capable of accurately assessing the extent of functional deficits and neuropathology that occur in this EAN.

Experimental autoimmune neuritis (EAN) is a well-appreciated experimental model of autoimmune peripheral demyelinating diseases. EAN disease is induced by ...