We would like to thank Curt Mazer from Isis Pharmaceuticals for providing advice pertaining to the ICV bolus surgery, as well as Isis Pharmaceuticals as a whole for supplying our lab with ASOs. Further, we would like to thank Carey Shaner for reviewing this article. TMM and SLD are supported by NIH Grants P50AG005681, K08NS074194, and R01NS078398.

The protocol below has been approved by the Institutional Animal Care and Use Committees at Washington University in St Louis and is in compliance with the National Institutes of Health guidelines for the use of experimental animals. If this is your first time performing surgeries, we recommend looking at the JoVE article on rodent surgeries as an introduction before beginning 14.
ASOs can be delivered to the mouse CNS through both osmotic pump infusion and a single ICV bolus injection. Because of this, the procedure detailed below is broken up into two segments. Pros and cons for each method will be touched on in the Discussion section. The coordinates used in the protocols are for adult C57BL6 and B6C3 mice. Corresponding rat coordinates can be found in Table 1.
ALZET OSMOTIC PUMP
1. Preparation of Alzet Osmotic Pumps
<ol>
	<li>Lay down a sterile drape and carefully drop the following items onto it while taking care not to directly touch anything in order to maintain sterility: osmotic pump(s), flow modulator(s), 1 ml sterile syringe(s), scalpel blade, and sections of tubing (1 piece of tubing = 5 pumps). For mice, there are three options for pumps: 14 days, 28 days, and 42 days (Figure 2A).</li>
	<li>Uncap and fill sterile 50 ml conical tubes with 15-20 ml saline solution. Up to 6 assembled pumps can be added per conical tube.</li>
	<li>Fill a p60 Petri dish with 100% ethanol. Add appropriate number of catheters to the ethanol. For mice, catheter length is 2.5 mm.</li>
	<li>Filter each ASO into as many 1.5 ml Eppendorf tubes as necessary. Include an Eppendorf tube filled with sterile 0.9% saline to fill the tubing with.</li>
	<li>Once everything is prepared, put on sterile gloves. Make sure not to touch anything that is not sterile. Change gloves as necessary.</li>
	<li>For each ASO to be used, fill a 1 ml syringe and loadpumps by placing the needle into the top hole and slowly filling until excess leaks out. Continue until all of the pumps are filled.</li>
	<li>Uncap the flow-modulators and insert into the pumps. Push the flow-modulator almost all of the way in, leaving a 3-5 mm gap.</li>
	<li>Using the scalpel blade, cut each piece of tubing into 5 equal segments. Take one piece of tubing, fill with sterile saline using a 1 ml syringe, and insert the top of the flow modulator into one end of the tubing. Then take a cannula from the ethanol and connect it to the other end of the tubing. The pump is now fully assembled.</li>
	<li>Place the fully assembled pump into the sterile 50 ml conical tube with saline for priming. The priming allows for the pumps to soak up fluid to initiate the drug delivery as well as come to the proper temperature. Priming time: 14 day pumps: 4-6 hr, 28 day pumps: 40 hr, 42 day pumps: 60 hr. When finished assembling the pumps, cap the conical tubes and place in a clean 37 &deg;C water bath until surgery.</li>
</ol>
2. Pre-surgical Procedure
<ol>
	<li>Wipe everything down with 70% ethanol to sterilize the area and lay down sterile drapes on the table top and stereotax to create a sterile field. It is important to be mindful of the sterile field throughout the entire surgery. Turn on the following: Bead Sterilizer, Heating Pad, Light, Digital Readout, and Oxygen/Isoflurane System (Figure 1).</li>
	<li>The following items will be required if doing either a single surgery or a whole batch of surgeries as we routinely do: 1 pair of forceps, 1 curved hemostat, 1 pair of small scissors, 1 pair of small curved blunted scissors, 1 rat bone cutter, 1 straight hemostat, 5-0 nylon sutures, 1 conical tubes with 70% ethanol, 1 conical tube with 95% ethanol, 1 conical tube with iodine, 1 conical tube with hydrogen peroxide, package of sterile cotton swabs, 1 bottle of super glue, 1 tube antibiotic ointment, 1 tube of eye lubricant, and 1 electric shaver.</li>
	<li>Place the tips of both pairs of scissors, forceps, and curved hemostat into the bead sterilizer for 15 sec. Remove and place in 70% ethanol. When performing more than one surgery, re-sterilize the tips of the instruments in between animals.</li>
	<li>Turn the isoflurane to 4% and O2 to 0.4 L/min and direct gas flow to the chamber. We have our isoflurane/oxygen system calibrated at least one a year to ensure that we are delivering the proper amount of anesthesia. Place mouse in the chamber and wait until breathing is significantly slowed. Remove the mouse and perform a toe pinch to ensure the mouse is completely unconscious.</li>
	<li>Shave the hair from the top of the shoulder up to in between the eyes. If the mouse begins to wake up, place the mouse back into the chamber until the mouse is completely unconscious again, confirming with a toe pinch. Place the anesthetized mouse on the stereotax and push the nose cone over the nose. Be gentle as the teeth and bones are fragile.</li>
	<li>Direct the gas flow to the stereotax to keep the mouse anesthetized. It is important to occasionally check to make sure the mouse is completely unconscious with a toe pinch throughout the surgery. Secure the head with the ear bars. We prefer leveling ear bars, though to target the lateral ventricle, a perfectly level skull is not necessary due to the large size of the ventricle.</li>
	<li>Turn the isoflurane level down to 2.0% for maintenance. Dab eye ointment onto each eye. As seen in the video, lay a sterile drape over the mouse so that only the base of the neck and head are exposed. Then disinfect the surgical area by wiping the top of the head and neck in a circular motion starting at the center of the shaved area and moving outwards first with a cotton swab dipped in 95% ethanol, followed by a cotton swab dipped in iodine.</li>
	<li>We recommend using a rectal probe feedback temperature controlled heating system that ensures the temperature of the mouse remains at a stable 37 &deg;C. If the temperature of the mouse decreases during the surgery, the post-surgery recovery may take longer and the resulting hypothermia can lead to aberrant protein activity, including hyperphosphorylation of the microtubule associated protein Tau15. After setting up the heating system, you are now ready to begin the surgery.</li>
</ol>
3. Implantation of Alzet Osmotic Pump
<ol>
	<li>Before beginning the surgery, put on a new pair of sterile gloves. Then, using forceps and a small pair of scissors, make an incision in the skin from the base of the neck above the animal's skull to up in between the eyes</li>
	<li>Take the blunted scissors with the curve facing up and slide underneath the skin at the base of the neck back towards the left hindlimb. There should be no resistance. If there is, remove the scissors and try again. This forms the subcutaneous pocket for the osmotic pump.</li>
	<li>Wipe the skull clean with sterile cotton swabs, followed by a cotton swab dipped in hydrogen peroxide to enhance bregma.</li>
	<li>Wipe down the 50 ml conical tubes that contain pumps to maintain sterility of the pumps. Carefully take a pump from the conical tube using the curved hemostat, taking extra care that the pump does not touch the sides of the conical tube. Hold the base of the pump with forceps and push the flow modulator in the rest of the way with the curved hemostat.</li>
	<li>Hold the pump where the flow modulator and tubing meet with the curved hemostat. Insert the pump under the skin at the base of the neck and push it back toward the left hindlimb as far as it will go without resistance. Be careful to not let the catheter touch anything.</li>
	<li>With the curved hemostat, grab the cannula at the groove where the top meets the pedestal. Move the cannula driver into position and secure into place.</li>
	<li>Place a single drop of super glue on the base of the cannula. Push the top of the cannula into the driver and position so that the tubing is pointed straight back.</li>
	<li>Touch the catheter tip to bregma and zero the coordinates on the Digital display. Raise the catheter and move 1.1 mm laterally to the right and 0.5 mm posterior (Figure 2B). Hold the skin out of the way with the curved hemostat.</li>
	<li>Drive the thin metal catheter through the skull until the plastic cannula base is securely pressed against the top of the skull. The metal catheter can be driven directly through the skull in mice due to the relative thin skull. If using rats, drill a hole in the skull before lowering the catheter.</li>
	<li>Pull any skin that has glue on it away from the skull. With the cannula driver holding the cannula/catheter in place, wait 1-2 min for the glue to fully dry.</li>
	<li>When practicing, to assess catheter placement, inject 20-40 &mu;l of 2.5% FastGreen Dye through the tubing (Figure 2C). Wait 2-3 min, perfuse the mouse with 1X PBS 0.03% heparin, and remove the brain. Cut coronally at the catheter site. If the catheter was placed in the ventricle, the dye will be perfused throughout the mouse ventricular system (Figure 2D).</li>
	<li>Hold the catheter in place with the curved hemostat while raising the driver. Slowly release the hemostat to ensure that the cannula is properly secured to the skull.</li>
	<li>With a cotton swab, press down on the top of the cannula. Fit the rat bone clippers into the grove between the top and base of the cannula. Clip off the top of the cannula while still pressing down with the cotton swab. Try and keep the clippers level so as not to detach the cannula from the skull. If the cannula does come unglued, quickly re-glue and apply pressure with a cotton swab for an additional 2 min.</li>
	<li>Using the 5-0 suture thread, regular hemostat, and forceps, suture along the full opening, paying extra attention right over the cannula. We use a running horizontal mattress suture (see Figure 5) due to the larger incision over the skull.</li>
	<li>Apply a dab of antibiotic ointment over the head and neck. Unscrew the ear bars and loosen the nose cone. Remove the mouse from the stereotax and place on a warming pad for recovery. Monitor the mice closely for the duration of recovery, typically ranging from 10-30 minutes. Check every 2-3 minutes until the mouse begins walking around and grooming itself.</li>
</ol>
4. Post-operative Care
<ol>
	<li>Monitor the mice daily after the surgery for the first week.</li>
	<li>If a mouse appears to be in pain or distress, we recommend providing the mouse with a 5 mg/kg subcutaneous injection of Carprofen once every 24 hr for up to 5 days in order to relieve the pain.</li>
	<li>If there appears to be an infection, generously apply antibiotic ointment to the area daily and consult an Attending Veterinarian to ensure that the wound properly heals.</li>
	<li>Remove the 5-0 nylon suture 7-10 days following surgery to prevent irritation from the suture thread.</li>
</ol>
5. Changing or Removing Alzet Osmotic Pump
After the pump is done actively infusing ASO, it can be either changed or completely removed.
<ol>
	<li>Follow the same pre-surgery procedure as above with the following changes: 1) you will not need the curved scissors, curved hemostat, rat bone cutters, nor the Digital Display, and 2) instead of prepping the head of the mouse, shave and disinfect the back of the mouse at the junction of the pump and tubing.</li>
	<li>Make a 1.5 cm incision on the back of the mouse, perpendicular to the pump/tubing junction, using forceps and scissors. Carefully pull the pump out of the incision without significantly pulling/pushing on the tubing as it may jar the catheter.</li>
	<li>To remove the pump, clip the tubing about 0.5 cm above the flow modulator and touch the superglue to the tubing to seal it shut. Wait 1-2 min for the glue to dry. Place the tubing back into the incision and suture shut. Dab antibiotic ointment onto the sutured skin and place the mouse on a heated recovery pad.</li>
	<li>To change the pump, have freshly made pumps ready along with a 10 cm dish filled with 95% ethanol. Catheters are not needed with the new pumps.</li>
	<li>Grab a new pump with the forceps. Dip your fingers into the ethanol, take hold of the pump, and discard the flow-modulator. Then, carefully and slowly pull off the old pump from the flow modulator attached to the tubing and cannula. As soon as it is off, slowly slide the new pump on, ensuring that the flow modulator never touches the outside of the mouse.</li>
	<li>Reinsert the new pump through the incision back into the subcutaneous packet and suture the skin. Dab antibiotic ointment on the wound and place mouse on recovery pad.</li>
	<li>Just as with the pump implantation, monitor the mice daily after the surgery to check for pain/discomfort and infections. Treat with Carprofen and antibiotics as deemed necessary.</li>
</ol>
ICV BOLUS
6. Pre-surgical Procedure
<ol>
	<li>Follow the same pre-surgery procedure as in Section 2 above with the following changes: 1) you will not need curved scissors, curved hemostat, or rat bone cutters; 2) clean a 10 &mu;l Hamilton syringe and needle with sterilized water and load with ASO; and 3) attach the syringe and needle to the stereotaxic apparatus where the cannula driver was located.</li>
</ol>
7. Bolus Injection of ASO
<ol>
	<li>Using forceps and a small pair of scissors, make an incision from the base of the neck up to in between the eyes.</li>
	<li>Wipe the skull clean with sterile cotton swabs, followed by a cotton swab dipped in hydrogen peroxide to enhance bregma.</li>
	<li>Move the syringe/needle into place and secure. Make sure that the beveled portion of the needle faces posterior. Lower the needle until it touches bregma. Zero all of the coordinates. Move the needle laterally to the right 1.0 mm and anterior 0.3 mm (Figure 3A).</li>
	<li>Slowly drive the needle through the skull just until the needle hole is flush with the top of the skull. Again, this can be done due to the thinness of the mouse skull. Zero the z-coordinate and lower the needle to -3.0 mm at a rate of 1 mm/sec (Figure 3B). Wait 2-3 min for the brain to seal around the needle.</li>
	<li>Deliver the full 10 &mu;l of ASO at a rate of 1 &mu;l per sec. Wait 2-3 min. We use a 1 &mu;l infusion rate for ASOs into the ventricle, though this rate may differ between different drugs. When practicing, check for proper placement of needle by delivering 10-20 &mu;l of 2.5% FastGreen Dye. Perfuse the mouse with 1X PBS 0.03% heparin soon after dye infusion and make coronal sections of the brain to verify that the dye is in the ventricular system (Figure 3C).</li>
	<li>Hold a cotton swab against the skull at the base of the needle. Raise the needle at a rate of 1 mm per sec. As soon as the needle is out of the skull, roll the cotton swab over the needle hole and hold for 1 min to prevent any ASO from leaking out.</li>
	<li>Suture the skin closed, apply antibiotic ointment, and place mouse on heated recovery pad. Depending on ASO concentration, it may take 20 min to a couple hours to fully recover.</li>
	<li>Just as with the pump implantation, monitor the mice daily after the surgery to check for pain/discomfort and infections. Treat with Carprofen and antibiotics as deemed necessary.</li>
</ol>

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The authors receive their Antisense Oligonucleotides from Isis Pharmaceuticals. Open Access fees sponsored by, Alzet Osmotic Pumps.

The ability to deliver drugs globally in the CNS as shown in the video is an extremely powerful technique that is easy to both learn and use. With practice, a single pump implantation or an ICV bolus can be completed in 10 min, allowing for large cohorts of mice to be treated at the same time. This is especially useful for studies with a behavioral readout, as larger numbers for mouse behavior are critical to help see significant differences.
Based on our experiences delivering ASOs via pumps and bolus injections, we have observed some pros and cons to each method. It should be noted that these are the opinions in our lab and may not be true for all mouse and rat models.
We find an advantage of using the pumps is their ability to deliver a high amount of ASO since the ASO is distributed over a much longer period of time. This usually equates to longer sustained knockdown or splicing after active ASO infusion, though this may not always be the case. The pumps also allow for a precise time frame of ASO delivery (14 days, 28 days, or 42 days) and the pumps can be changed once to allow for even longer active ASO infusion. However, we have noticed that changing pumps more than once increases the variability due to the formation of fibrous pockets around the pump that prevent the pump from properly absorbing fluid. A disadvantage of the pumps is that some mice do not tolerate the pump as well as others. If the transgenic line you are working with is more fragile, a pump may be too cumbersome. The pumps also need to be removed after the final infusion, subjecting the mice to another surgery and added anesthesia. If doing behavior work, it is especially important to remove the pump and allow for at least 1-2 weeks of recovery since the presence of the pump will affect some behaviors in the mice.
With ICV bolus injections, one advantage is cost. There is an upfront investment to purchase the syringes and needles, but over time, bolus injections are more cost-effective since there are no pumps/tubing/catheters to purchase. Overall, there is less up-keep with the ICV bolus injections due to the lack of cannulas and pumps. We also find that ICV bolus can be used to deliver ASOs to younger and/or more fragile mice. A disadvantage of the ICV bolus is that it is a single injection. Not as much overall ASO can be delivered through this route, and if the duration of action of the ASO being used is short, the knockdown/splicing effects will also be short-lived.
Both the osmotic pumps as well as the ICV bolus have the capability to deliver ASOs that can knockdown proteins or alter splicing of genes in the entire rodent CNS, a technique that has wide applications in multiple neuroscience-related fields. We suggest piloting both methods of delivery if you are unsure which route of administration is best suited for your specific study.

Some drugs are unable to cross the blood brain barrier (BBB), requiring direct Central Nervous System (CNS) delivery. To circumvent the BBB, drugs can be delivered directly into the brain using the described methods. While our lab and the paper detailed herein focus on Antisense Oligonucleotides (ASOs), other drugs, such as small molecules, antibodies, gene therapy vectors, etc., can also be delivered through the exact same approach.
Certain proteins play an instrumental role in the pathogenesis of neurodegenerative diseases. Such proteins often form toxic species and accumulate into aggregates, leading to eventual neuronal death and subsequent neurological disease 3-4. In an effort to slow or even halt the progression of these diseases, one therapeutic option may be to directly target and decrease the causative protein. However, these proteins are often found ubiquitously through the CNS, making it difficult to effectively target them on a global scale.
In order to target genes throughout the entire CNS, we administer ASOs into mouse cerebrospinal fluid (CSF) via the lateral ventricle to bypass the BBB. This specific method takes advantage of the mouse ventricular system that bathes the entire brain and spinal cord, allowing widespread distribution of the ASOs. We use ASOs that are 18-20 mer RNA-like molecules that directly bind the target mRNA sequence and, depending on the ASO chemical modifications, either A) recruit RNase-H to degrade the mRNA leading to knockdown or B) shift alternative splicing 1,2,16,17. It should be noted that multiple molecules exist for knocking down a specific protein in vivo, including shRNA. Since these molecules are not the focus of this article, we direct the reader to review articles that better detail knockdown mechanisms of action and the advantages/disadvantages of each 5-6.
In prior work, we have used ASOs to target the protein superoxide dismutase 1 (SOD1) in a transgenic rat model of Amyotrophic Lateral Sclerosis (ALS) 7 (Figure 4). Mutations in SOD1 occur in approximately 2% of all ALS cases 8, though it has been recently hypothesized that SOD1 may play an important role in sporadic ALS as well 9-10. By decreasing total SOD1 levels in the ALS transgenic rat, survival after onset was significantly increased 7. These important data were the first to show that an ASO treatment in the CNS could have a profound positive impact on a neurological disease model. Since then, ASOs targeted to human SOD1 have entered and successfully completed a Phase I human clinical trial with minimal side effects (Clnicaltrails.gov NCT01041222), as presented at the 2012 64th Annual American Academy of Neurology. Plans to move the ASOs forward to Phase II trials are currently underway.
While targeting SOD1 was the first demonstration of using ASOs to treat a neurological disease, several other studies have since been performed looking at different diseases and their respective protein targets. In 2010 and 2011, ASOs that shift splicing of the protein survival of motor neuron 2 (SMN2)were used in transgenic mouse models of spinal muscle atrophy (SMA) and resulted in a significant improvement in the disease phenotypes 11,12. These splicing ASOs are now in Phase I clinical trials in children with SMA (Clinicaltrails.gov NCT01494701). Additionally, it was recently shown that transient administration of ASOs targeted against the huntingtin gene were able to dramatically rescue the Huntington's mouse model, even after the huntingtin protein levels returned to baseline 13.
In all of these studies, ASOs were delivered to the lateral ventricle to decrease total gene levels or alter gene splicing through the entire CNS. Both osmotic pumps and a single bolus injection can be used to deliver ASOs to the CSF. Pumps allow for a slow, continuous delivery, whereas the intracerebroventricular (ICV) bolus is a fast, one-time injection. We have used both of these methods with success, though we have not reported the direct comparison between pump and bolus in a single transgenic line.
Using ASOs in the CNS is a powerful way to decrease total protein levels and/or change splicing of several proteins. While we use ASOs exclusively as a treatment for neurological disorders, we recognize that other fields may also benefit from this technique. As long as the protein of interest is expressed in the CNS and the ultimate goal is to achieve CNS-wide changes in gene expression, using ASOs in the demonstrated techniques may be very useful.

<table border="1">
 <tbody>
 <tr>
 <td>Name of Reagent/Material</td>
 <td>Company</td>
 <td>Catalog Number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>PREPARING ALZET OSMOTIC PUMPS </td>
 </tr>
 <tr>
 <td>Alzet Osmotic Pump 14 days</td>
 <td>DURECT</td>
 <td>Model 1002</td>
 <td></td>
 </tr>
 <tr>
 <td>Alzet Osmotic Pump 28 days</td>
 <td>DURECT</td>
 <td>Model 2004</td>
 <td></td>
 </tr>
 <tr>
 <td>Alzet Osmotic Pump 42 days</td>
 <td>DURECT</td>
 <td>Model 2006</td>
 <td></td>
 </tr>
 <tr>
 <td>2.5 mm Catheters</td>
 <td>PlasticsOne</td>
 <td>3280PM/SPC</td>
 <td>Custom ordered to 2.5 mm Catheter Length</td>
 </tr>
 <tr>
 <td>Vinyl Catheter Tubing</td>
 <td>DURECT</td>
 <td>7760</td>
 <td>ID: 0.027", OD: 0.045"</td>
 </tr>
 <tr>
 <td>0.9% Sodium Chloride, Irrigation, USP</td>
 <td>Baxter</td>
 <td>2F7124</td>
 <td>NOT to be used in pumps or tubing</td>
 </tr>
 <tr>
 <td>0.9% Sodium Chloride, Injection, USP</td>
 <td>Hospira</td>
 <td>NDC 0409-4888-10</td>
 <td></td>
 </tr>
 <tr>
 <td>p60 Petri Dish (Sterilized)</td>
 <td>TRP</td>
 <td>93060</td>
 <td></td>
 </tr>
 <tr>
 <td>Surgical Blades (Sterile)</td>
 <td>Butler Schein</td>
 <td>#007319</td>
 <td></td>
 </tr>
 <tr>
 <td>Latex Surgical Gloves (Sterile)</td>
 <td>Micro-Touch</td>
 <td></td>
 <td>CatNo will depend on size of the gloves needed</td>
 </tr>
 <tr>
 <td>Sterile Towel Drape</td>
 <td>Dynarex</td>
 <td>4410</td>
 <td></td>
 </tr>
 <tr>
 <td>.2um Syringe Filters</td>
 <td>PALL</td>
 <td>4192</td>
 <td></td>
 </tr>
 <tr>
 <td>1 ml Syringe (Sterile)</td>
 <td>BD</td>
 <td>309625</td>
 <td></td>
 </tr>
 <tr>
 <td>50 ml Conical Tubes</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>100% Ethanol</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>PUMP &amp; BOLUS SURGERY PROTOCOLS </td>
 </tr>
 <tr>
 <td>Curved Forceps</td>
 <td>Fine Science Tools</td>
 <td>11001-12</td>
 <td></td>
 </tr>
 <tr>
 <td>Curved Hemostat</td>
 <td>Fine Science Tools</td>
 <td>13009-12</td>
 <td></td>
 </tr>
 <tr>
 <td>Fine Sharp Scissors</td>
 <td>Fine Science Tools</td>
 <td>14060-09</td>
 <td></td>
 </tr>
 <tr>
 <td>Curved Blunt Scissors</td>
 <td>Fine Science Tools</td>
 <td>14029-10</td>
 <td></td>
 </tr>
 <tr>
 <td>Bone Cutter</td>
 <td>Fine Science Tools</td>
 <td>16104-14</td>
 <td></td>
 </tr>
 <tr>
 <td>Straight Hemostat</td>
 <td>Fine Science Tools</td>
 <td>12002-12</td>
 <td></td>
 </tr>
 <tr>
 <td>Syringe</td>
 <td>Hamilton</td>
 <td>7653-01</td>
 <td>10 &mu;l gas-tight with removable needles</td>
 </tr>
 <tr>
 <td>Needles</td>
 <td>Hamilton</td>
 <td>7758-04</td>
 <td>26 gauge, Point Style: 2</td>
 </tr>
 <tr>
 <td>5-0 Nylon Suture Thread</td>
 <td>Covidient</td>
 <td>SN-871</td>
 <td></td>
 </tr>
 <tr>
 <td>Alcohol Pads</td>
 <td>Select</td>
 <td>#521</td>
 <td></td>
 </tr>
 <tr>
 <td>Cotton Swabs (sterile)</td>
 <td>Puritan</td>
 <td>REF 806-WC</td>
 <td></td>
 </tr>
 <tr>
 <td>Super Glue</td>
 <td>Loctite</td>
 <td></td>
 <td>Longneck Bottles</td>
 </tr>
 <tr>
 <td>CAUTION: FastGreen Dye</td>
 <td>Sigma</td>
 <td>F7252-5G</td>
 <td>Wear Eyeshields and Gloves when handling this product</td>
 </tr>
 <tr>
 <td>Antibiotic Cream</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Eye Ointment</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Electric Shaver</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>70% Ethanol</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>10% Provadone Iodine</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>3% Hydogen Peroxide</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Warming Pad</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Bead Sterilizer</td>
 <td>SouthPointe Surgical</td>
 <td>GRM5-1450</td>
 <td></td>
 </tr>
 <tr>
 <td>Small Animal Stereotaxic</td>
 <td>Kopf</td>
 <td>Model 940</td>
 <td></td>
 </tr>
 <tr>
 <td>Nose Cone</td>
 <td>Kopf</td>
 <td>Model 923-B</td>
 <td></td>
 </tr>
 <tr>
 <td>Ear Bars</td>
 <td>Kopf</td>
 <td>Model 921</td>
 <td>This model is optional</td>
 </tr>
 <tr>
 <td>Cannula Driver</td>
 <td>Kopf</td>
 <td>Model 1966</td>
 <td></td>
 </tr>
 <tr>
 <td>Syringe Holder</td>
 <td>Kopf</td>
 <td>Model 1972</td>
 <td></td>
 </tr>
 <tr>
 <td>Temperature Control System</td>
 <td>Kopf</td>
 <td>Model TCAT-2LV</td>
 <td>Optional</td>
 </tr>
 <tr>
 <td>Oxygen/Isoflurane System</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 </tbody>
</table>

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.

After pump infusion or a designated time after the ICV bolus injection (we routinely use four weeks), it is important to test the efficacy of the ASOs. We recommend taking various regions of brain and spinal cord to measure levels/isoforms of the target gene, both at the mRNA level using quantitative real-time PCR as well as the protein level using either Western Blot or ELISA (Figure 4 for a published example). This will help determine ASO efficacy throughout the different CNS regions. If the half-life of the targeted protein is long, we advise waiting longer after ASO infusion in order to assess maximum protein knockdown or splicing.
It is also important to test oligo distribution. Pump infusions result in distribution through both ipsilateral and contralateral hemispheres, though a higher ASO concentration is often seen in areas closer to the ventricular system 16. ICV bolus injections yield a more uniform distribution of oligo through the CNS 16, though the effects may not last as long. We direct the reader to Southwell et al. to see a direct comparison of pump versus bolus oligo distribution 16.
We also recommend testing multiple doses as well as the duration of action of the ASOs since each ASO will behave slightly differently in vivo. ASOs typically have a relatively long duration of action, with target knockdown or splicing lasting several months post ASO delivery. In addition, some ASOs will work better than others and some gene targets will be easier to knockdown than others. Because of this, when screening ASOs to determine the lead candidate, we recommend testing at least 3-5 ASOs in vivo with an n = 4-6 adult mice per ASO as a first pass.
If using a drug other than ASOs, it will also be important to pilot the ideal drug concentration, drug duration of action, and CNS drug distribution for that specific compound.
<img alt="Table 1" fo:content-width="5in" fo:src="/files/ftp_upload/50326/50326table1highres.jpg" src="/files/ftp_upload/50326/50326table1.jpg" /> 
Table 1. Surgery Coordinates. The coordinates used to hit the right lateral ventricle in both mice and rats when implanting osmotic pumps as well as performing an intracerebroventricular (ICV) bolus. Both coordinates, for pumps and ICV bolus, target the right lateral ventricle. We use two different sets because these are what we have the most experience with and know provide excellent distribution of ASO.
<img alt="Figure 1" src="/files/ftp_upload/50326/50326fig1.jpg" /> 
 Figure 1. Stereotaxic Setup. (A) The necessary stereotaxic equipment for Alzet pump and Bolus injection surgeries. i) Glass Bead Sterilizer. ii) Digital Readout. iii) Stereotaxic base with moving arms. iv)Temperature Controlled Heating Pad. v)Isoflurane/Oxygen System. vi)Isoflurane Chamber. (B) Close up of the nose cone and ear bars. (C) Two attachments required. Left: Syringe/needle holder for ICV bolus injections. Right: Cannula Driver for Osmotic pump implantation.
<img alt="Figure 2" src="/files/ftp_upload/50326/50326fig2.jpg" /> 
 Figure 2. Alzet Osmotic Pump Implantation. (A) Alzet osmotic pump options for mice. 14 day pumps (0.25 &mu;l/hr), 28 day pumps (0.25 &mu;l/hr), 42 day pumps (0.15 &mu;l/hr). (B) Coordinates for pump implantation from bregma: -0.5 mm Posterior, -1.1 mm Lateral (Right), and -2.5 mm Ventral (length of catheter). (C) After driving the catheter through the skull and gluing the cannula, dye is flushed through the catheter to assess proper placement of the catheter in the lateral ventricle. (D) Perfused brain immediately after dye administered as in (C). If catheter is successfully placed in ventricle, the dye will be distributed throughout the mouse ventricular system.
<img alt="Figure 3" src="/files/ftp_upload/50326/50326fig3.jpg" /> 
 Figure 3. Intracerebroventricular Bolus Injection. (A) Coordinates for bolus injection from bregma: +0.3 mm Anterior, -1.0 mm Lateral (Right), and -3.0 mm Ventral. (B) After driving the needle through the skull -3.0 mm ventral, dye is pushed through the syringe to assess proper placement of the needle in the later ventricle. (C) Perfused brain shortly after dye administered as in (B). If needle is successfully placed in ventricle, the dye will be distributed throughout the mouse ventricular system.
<img alt="Figure 4" fo:content-width="5.5in" fo:src="/files/ftp_upload/50326/50326fig4highres.jpg" src="/files/ftp_upload/50326/50326fig4.jpg" /> 
 Figure 4. Antisense Oligonucleotides reduce rat SOD1 in vivo (figure taken directly from Smith et al. 7). (A-D) Antisense SOD1 oligonucleotides SODr146192 or SODscrambled were infused for 28 days into the right lateral ventricle of normal rats at 100 &mu;g/day. (A) Endogenous SOD1 mRNA levels from brain and spinal cord as measured by qRT-PCR. (B) SOD1 and a-tubulin protein levels following ASO infusion. (C and D) Protein levels for tubulin and SOD1 quantified for different regions of the brain following infusion. (E) ASOs against presenilin 1 or GSK-3&beta; were infused for 2 weeks into the right lateral ventricle of nontransgenic mice and mRNA levels assessed in the right frontal/temporal cortex. Reproduced with permission from the American Society for Clinical Investigation. <a href="https://www.jove.com/files/ftp_upload/50326/50326fig4large.jpg" target="_blank"> Click here to view larger figure</a>.
<img alt="Figure 5" src="/files/ftp_upload/50326/50326fig5.jpg" fo:content-width="3in" fo:src="/files/ftp_upload/50326/50326fig5highres.jpg"/> 
 Figure 5. Running Horizontal Mattress Suture. 5-0 nylon suture thread is weaved in and out of the skin, starting anterior and working posterior, concentrating more over the cannula. Once having reached the most posterior point, the suture thread is then brought back up and threaded over the cannula once more. This ensures proper wound closure and prevents the cannula/tubing from working through the skin.

Watch this Scientific Journal Video about Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System at JoVE.com

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

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 ...

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59.4K ビュー.  Washington University in St. Louis School of Medicine. 我々は、カテーテルを注入又はマウスにおける右側脳室へのボーラス注入を実行することにより、中枢神経系に薬物を標的化する方法について述べる。私たちは、アンチセンスオリゴヌクレオチドの配信に特に焦点を当てています。この技術は、他の薬剤に、ラットに容易に適用可能である。

ビデオ: Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

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 ...

Multimodal Imaging of Stem Cell Implantation in the Central Nervous System of Mice

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, both in preclinical and clinical studies. However, to date results regarding functional outcome and/or tissue regeneration following stem cell transplantation are quite diverse. Generally, a clinical benefit is observed without profound understanding of the underlying mechanism(s)1. Therefore, multiple efforts have led to the development of different molecular imaging modalities to monitor stem cell grafting with the ultimate aim to accurately evaluate survival, fate and physiology of grafted stem cells and/or their micro-environment. Changes observed in one or more parameters determined by molecular imaging might be related to the observed clinical effect. In this context, our studies focus on the combined use of bioluminescence imaging (BLI), magnetic resonance imaging (MRI) and histological analysis to evaluate stem cell grafting.
BLI is commonly used to non-invasively perform cell tracking and monitor cell survival in time following transplantation2-7, based on a biochemical reaction where cells expressing the Luciferase-reporter gene are able to emit light following interaction with its substrate (e.g. D-luciferin)8, 9. MRI on the other hand is a non-invasive technique which is clinically applicable10 and can be used to precisely locate cellular grafts with very high resolution11-15, although its sensitivity highly depends on the contrast generated after cell labeling with an MRI contrast agent. Finally, post-mortem histological analysis is the method of choice to validate research results obtained with non-invasive techniques with highest resolution and sensitivity. Moreover end-point histological analysis allows us to perform detailed phenotypic analysis of grafted cells and/or the surrounding tissue, based on the use of fluorescent reporter proteins and/or direct cell labeling with specific antibodies.
In summary, we here visually demonstrate the complementarities of BLI, MRI and histology to unravel different stem cell- and/or environment-associated characteristics following stem cell grafting in the CNS of mice. As an example, bone marrow-derived stromal cells, genetically engineered to express the enhanced Green Fluorescent Protein (eGFP) and firefly Luciferase (fLuc), and labeled with blue fluorescent micron-sized iron oxide particles (MPIOs), will be grafted in the CNS of immune-competent mice and outcome will be monitored by BLI, MRI and histology (Figure 1).

This article describes an optimized sequence of events for multimodal imaging of cellular grafts in rodent brain using: (i) in vivo bioluminescence and magnetic resonance imaging, and (ii) post mortem histological analysis. Combining these imaging modalities on a single animal allows cellular graft evaluation with high resolution, sensitivity and specificity.

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, ...

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 ...

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

In this article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the lipophilic fluorescent membrane marker DiI. This method allows the visualization of neuronal cell morphology in great detail. It is possible to label any cell in the CNS: cell bodies of target neurons are visualized under DIC optics or by expression of a fluorescent genetic marker such as GFP. After labeling, the DiI can be transformed into a permanent brown stain by photoconversion to allow visualization of cell morphology with transmitted light and DIC optics. Alternatively, the DiI-labeled cells can be observed directly with confocal microscopy, enabling genetically introduced fluorescent reporter proteins to be colocalised. The technique can be used in any animal, irrespective of genotype, making it possible to analyze mutant phenotypes at single cell resolution.

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

We present a technique for labeling single neurons in the central nervous system (CNS) of Drosophila embryos, which allows the analysis of neuronal morphology by either transmitted light or confocal microscopy.

In this  article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the ...

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. Understanding repair and regeneration in animals is a key question in biology. The damaged human CNS does not regenerate, and understanding how to promote the regeneration is one of main goals of medical neuroscience. The powerful genetic toolkit of Drosophila can be used to tackle the problem of CNS regeneration.
A lesion to the CNS ventral nerve cord (VNC, equivalent to the vertebrate spinal cord) is applied manually with a tungsten needle. The VNC can subsequently be filmed in time-lapse using laser scanning confocal microscopy for up to 24 hr to follow the development of the lesion over time. Alternatively, it can be cultured, then fixed and stained using immunofluorescence to visualize neuron and glial cells with confocal microscopy. Using appropriate markers, changes in cell morphology and cell state as a result of injury can be visualized. With ImageJ and purposely developed plug-ins, quantitative and statistical analyses can be carried out to measure changes in wound size over time and the effects of injury in cell proliferation and cell death. These methods allow the analysis of large sample sizes. They can be combined with the powerful genetics of Drosophila to investigate the molecular mechanisms underlying CNS regeneration and repair.

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An injury paradigm using the Drosophila larval ventral nerve cord to investigate central nervous system regeneration and repair is described. Stabbing followed by laser scanning confocal microscopy in time-lapse and fixed specimens, combined with quantitative analysis with purposefully developed software and genetics, are used to investigate the molecular mechanisms of CNS regeneration and repair.

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. ...

In vivo Interrogation of Central Nervous System Translatome by Polyribosome Fractionation

Multiple processes are involved in gene expression including transcription, translation and stability of mRNAs and proteins. Each of these steps are tightly regulated, affecting the final dynamics of protein abundance. Various regulatory mechanisms exist at the translation step, rendering mRNA levels alone an unreliable indicator of gene expression. In addition, local regulation of mRNA translation has been particularly implicated in neuronal functions, shifting &#39;translatomics&#39; to the focus of attention in neurobiology. The presented method can be used to bridge transcriptomics and proteomics.
Here we describe essential modifications to the technique of polyribosome fractionation, which interrogates the translatome based on the association of actively translated mRNAs to multiple ribosomes and their differential sedimentation in sucrose gradients. Traditionally, working with in vivo samples, particularly of the central nervous system (CNS), has proven challenging due to the restricted amounts of material and the presence of fatty tissue components. In order to address this, the described protocol is specifically optimized for use with minimal amount of CNS material, as demonstrated by the use of single mouse spinal cord and brain. Briefly, CNS tissues are extracted and translating ribosomes are immobilized on mRNAs with cycloheximide. Myelin flotation is then performed to remove lipid rich components. Fractionation is performed on a sucrose gradient where mRNAs are separated according to their ribosomal loading. Isolated fractions are suitable for a range of downstream assays, including new genome wide assay technologies.

In vivo Interrogation of Central Nervous System Translatome by Polyribosome Fractionation

This protocol illustrates essential modifications of polyribosome fractionation in order to study the translatome of in vivo CNS samples. It allows global assessment of translation and transcription regulation through the isolation and comparison of total RNA to ribosome bound RNA fractions.

Multiple processes are involved in gene expression including transcription, translation and stability of mRNAs and proteins. Each of these steps are ...

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 ...

Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique non-linear deformations, dramatically hampering one's ability to evaluate cellular morphology, distribution and connectivity in the central nervous system (CNS). However, optical clearing techniques are changing the way tissues are visualized. These approaches permit one to probe deeply into intact organ preparations, providing tremendous insight into the structural organization of tissues in health and disease. Techniques such as Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) achieve this goal by providing a matrix that binds important biomolecules while permitting light-scattering lipids to freely diffuse out. Lipid removal, followed by refractive index matching, renders the tissue transparent and readily imaged in 3 dimensions (3D). Nevertheless, the electrophoretic tissue clearing (ETC) used in the original CLARITY protocol can be challenging to implement successfully and the use of a proprietary refraction index matching solution makes it expensive to use the technique routinely. This report demonstrates the implementation of a simple and inexpensive optical clearing protocol that combines passive CLARITY for improved tissue integrity and 2,2&#8242;-thiodiethanol (TDE), a previously described refractive index matching solution.

Optical clearing techniques are revolutionizing the way tissues are visualized. In this report we describe modifications of the original Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) protocol that yields more consistent and less expensive results.

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique ...

A Computerized Test Battery to Study Pharmacodynamic Effects on the Central Nervous System of Cholinergic Drugs in Early Phase Drug Development

Investigating potential pharmacodynamic effects in an early phase of central nervous system (CNS) drug research can provide valuable information for further development of new compounds. A computerized and thoroughly validated battery of neuropsychological and neurophysiological tests has been shown to be sensitive to detect drug-induced effects of multiple new and existing compounds. The test battery covers the main CNS domains, which have been shown to respond to drug effects and can be repeatedly administered following drug administration to characterize the concentration-effect profile of a drug.
The standard tests in the battery are saccadic eye movement, smooth pursuit eye movement, the Bowdle visual analog scale (VAS), the Bond and Lader VAS, body sway, adaptive tracking, visual verbal learning, and quantitative electroencephalography (qEEG). However, the test battery is adaptive in nature, meaning that it can be composed and adjusted with tests fit to investigate specific drug classes, or even specific receptors.
Showing effects of new cholinergic drugs designed to have a pro-cognitive outcome has been difficult. The pharmacological challenge model is a tool for early proof-of-pharmacology. Here, a marketed drug is used to induce temporary and reversible disease-like symptoms in healthy subjects, via a pharmacological mechanism related to the disease that is targeted as indication for the new compound. The test battery was implemented to investigate the potential of the nicotinic receptor antagonist mecamylamine to be used as a challenge model for cholinergic dysfunction, as seen in neurodegenerative disorders.
A worsening of scores in a dose dependent manner on the visual verbal learning test (VVLT; a test for learning and memory abilities) and the adaptive tracking test (a measure of visuomotor control and arousal), in particular, showed that the test battery is sensitive to showing acute pharmacodynamic effect after administration of anti-cholinergic drugs.

A validated computerized battery of neuropsychological and neurophysiological tests is used to study pharmacodynamic effects on the central nervous system of newly developed drugs in early phase development. To demonstrate the test battery, the acute effects of mecamylamine and the reversal of these effects by two agonist drugs are described.

Investigating potential pharmacodynamic effects in an early phase of central nervous system (CNS) drug research can provide valuable information for ...

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar Drosophila Melanogaster

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield deleterious consequences, yet the current methods for recording nervous system activity of an individual animal is costly and laborious. This suction electrode preparation of the third-instar larval central nervous system of Drosophila melanogaster, is a tractable system for testing the physiological effects of neuroactive agents, determining the physiological role of various neural pathways to CNS activity, as well as the influence of genetic mutations to neural function. This ex vivo preparation requires only moderate dissecting skill and electrophysiological expertise to generate reproducible recordings of insect neuronal activity. A wide variety of chemical modulators, including peptides, can then be applied directly to the nervous system in solution with the physiological saline to measure the influence on the CNS activity. Further, genetic technologies, such as the GAL4/UAS system, can be applied independently or in tandem with pharmacological agents to determine the role of specific ion channels, transporters, or receptors to arthropod CNS function. In this context, the assays described herein are of significant interest to insecticide toxicologists, insect physiologists, and developmental biologists for which D. melanogaster is an established model organism. The goal of this protocol is to describe an electrophysiological method to enable the measurement of electrogenesis of the central nervous system in the model insect, Drosophila melanogaster, which is useful for testing a diversity of scientific hypotheses.

Electrophysiological Recording of The Central Nervous System Activity of Third-Instar  Drosophila Melanogaster

This protocol describes a method to record the descending electrical activity of the Drosophila melanogaster central nervous system to enable the cost-efficient and convenient testing of pharmacological agents, genetic mutations of neural proteins, and/or the role of unexplored physiological pathways.

The majority of the currently available insecticides target the nervous system and genetic mutations of invertebrate neural proteins oftentimes yield ...