Name: systemic and local drug delivery for treating diseases of the central nervous system in rodent models
Uploaded: 2010-08-16T17:00:00
Duration: 11 min 51 s
Description: Watch this Scientific Journal Video about Systemic and Local Drug Delivery for Treating Diseases of the Central Nervous System in Rodent Models at JoVE.com

NS65819 (CDJ, TO), NS049720 (CDJ), CA097257 (CDJ, TO), CIRM DR1-01426 We would like to thank Raquel Santos for technical assistance.

1. Systemic Delivery Methods
Although systemic delivery of drugs is not the most efficient approach for achieving high drug concentrations in the CNS, systemic deliveries are convenient and well accepted by patients. Here, we will demonstrate proper procedures for three routinely used systemic administration approaches: intravenous injection, intraperitoneal injection, and oral gavage.
A. Intravenous Injection (Tail Vein Injection)
<ol>
	<li>Before beginning the procedure, the body weight of each mouse should be recorded. As with all preclinical studies, body weight should be monitored regularly (twice a week or greater) to assess potential drug toxicity. No more than 1% of the mouse's body weight in volume should be injected at a single time. For example, no more than 0.2mL of fluid should be injected in a 20g mouse. All fluids injected intravenously should be sterilized by an appropriate method.</li>
	<li>The mouse should be warmed for 5-10 minutes using either a heating pad or a heat lamp to dilate the tail vein. If using a heat lamp, monitor the mouse at all times to avoid hyperthermia.</li>
	<li>The mouse is transferred to a holding device, which restrains the mouse while allowing access to the tail vein (see video). In the mouse tail there are four visible vessels: the vessels on the dorsal and lateral sides are veins, and the vessel on the ventral side is an artery. To access the tail vein, the tail of the mouse is held at the most distal point, and rotated 90 degrees so that the vein is facing upwards. The injection site is cleaned with an alcohol swab and a 28g insulin syringe is inserted, beveled side up, into the vein (see video). If the needle is properly placed in the vein, it should move freely and without pressure. Slowly inject the drug with even pressure over 5-10 seconds. If a blister appears on the tail, stop injecting, as this indicates that the needle is no longer in the vein.</li>
	<li>After injection, apply gentle pressure to the injection site until the bleeding stops. This normally takes 30-60 seconds. Monitor the mouse for 5-10 minutes after the injection to insure that there is no further bleeding.</li>
</ol>
B. Intraperitoneal Injection
<ol>
	<li>Before beginning the injection, the drug should be loaded into a syringe attached to a 28g needle. Make sure that there is space in the syringe for drawing back the plunger before the injection (e.g., if injecting 200&mu;L, make sure that the syringe capacity is 300&mu;L or greater).</li>
	<li>Remove the mouse from the cage by its tail and place it onto a textured surface, so that the mouse has something to grip. A cage lid is usually sufficient. Allow the mouse to stretch its body and then using your non-dominant hand, grasp the skin on the back of the mouse, taking care to lightly pinch as much skin as possible between your thumb and index and middle fingers (see video). Turn the mouse over so that the abdomen of the mouse is facing upwards. If the mouse can freely move its head, release the grip and try again, so as to avoid being bitten during the injection.</li>
	<li>With your dominant hand, pick up the syringe and insert the needle at a 30 degree angle into the lower left quadrant of the mouse (see video). Holding the mouse slightly inverted will help move the organs away from the injection site. To ensure that the needle is in the intraperitoneal space, pull back on the syringe plunger. If any fluid or blood appears in the syringe, then the needle is not in the intraperitoneal space and should be removed. If no fluid is aspirated into the syringe, then inject the syringe contents with an even pressure over 1-5 seconds and release the mouse.</li>
	<li>Monitor the mouse for 5-10 minutes after the injection to ensure that the mouse returns to normal activity levels.</li>
</ol>
C. Oral Gavage
<ol>
	<li>Before beginning the injection, record the weight of the mouse. The maximum volume that can be delivered by oral gavage is 10mL per kilogram of body weight. For example, the maximum volume for a 20g mouse would be 200&mu;L. Attempting to inject larger volumes may result in reflux, which will cause incomplete drug delivery. If it is necessary to administer volumes greater than indicated above, as many as three doses may be administered over 24 hours.</li>
	<li>To avoid puncturing the esophagus, it is important to measure the length of the gavage needle for each mouse. Hold an 18g ball tip curved gavage needle at the last rib of the mouse, and then mark the length at the tip of the mouse's head using a permanent marker (see video). During the gavage, this mark will be a stopping point as you insert the needle into the mouth of the mouse.</li>
	<li>Restrain the mouse using the same hand grip as for an intraperitoneal injection. Insert the gavage needle in the mouth, over the tongue, and advance the needle through the pharynx. Do not insert the needle past the stopping mark. The needle should proceed smoothly without any pressure (see video). If you encounter pressure, stop and withdraw the needle to avoid injecting fluid into the lungs.</li>
	<li>With the needle in place, depress the plunger over 1-5 seconds and then remove the needle at the same angle that it was inserted. Monitor the mouse for 5-10 minutes, paying close attention to any signs of labored breathing which may indicate that fluid has entered the lungs.</li>
</ol>
2. Local Delivery
Acute Convection-Enhanced Delivery
A. Probe Construction
A reflux-resistant CED cannula for rodents is not yet commercially available. Here we will demonstrate a method for cannula construction that was adapted from a method first described by Krauze et al (Krauze 2005).
<ul>
	<li>The cannula has three parts (Figure 1): 100uM diameter silica tubing through which the infusate flows, a rigid metal needle for structural support, and flexible Teflon tubing for loading the infusate.</li>
	<li>To obtain the rigid metal needle, use an open flame to melt the plastic on a Surflo IV Catheter (24g stylet) and using tweezers remove the metal needle. The rest of the catheter can be discarded. Cut a length of silica tubing (OD 0.163mm), slightly longer than the metal needle, using a single-edge razor blade. Role the silica tubing in a small drop of cyanoacrylate based fast-acting adhesive (e.g. Krazy Glue), taking care not to get any glue on the ends of the tubing. Insert the silica tubing in the metal needle and let dry for 5 minutes.</li>
	<li>Once dry, the silica tubing should be firmly affixed to the needle. The ends of the silica should be trimmed so that 2mm of silica tubing protrudes from the pointed end of the needle and 3mm of silica tubing protrudes from the flat end (see video).</li>
	<li>Cut a section of Teflon tubing 20 cm in length. Roll the metal needle in a small drop of adhesive, again taking care not to get glue on the ends of the silica tubing as this will clog the cannula. Insert the needle into the Teflon tubing to a depth of 1cm. Let dry for 1 minute. Using a glue gun, apply a drop of hot glue to the joint between the metal needle and the Teflon tubing (see video). Ensure that the entire joint is covered on all sides and let dry for at least 1 hour. Cannulas can be made up to a week in advance and stored at room temperature.</li>
</ul>
B. Infusion Procedure
<ul>
	<li>To prepare the surgical area, all surfaces should be sprayed with a disinfectant, such as a 2% chlorhexidine solution. Sterile surgical gloves should be worn during the procedure. The surfaces are then covered with absorbent drapes. The following supplies should be placed in the surgical area:
 <ul>
 	<li>Heating pad to maintain mouse body temperature</li>
		<li>Two small Petri dishes; one containing 3% hydrogen peroxide, and one containing 2% chlorhexidine</li>
		<li>Sterile gauze and cotton swabs</li>
		<li>Sterile disposable scalpels (Number 21)</li>
		<li>Sterile 22g needles</li>
		<li>Mouse stereotaxic frame</li>
		<li>Controlled rate syringe pump</li>
		<li>Autoclaved mouse skin stapler, staples, and staple remover</li>
 </ul>
 </li>
	<li>To set-up the CED cannula, attach a 1mL syringe to the Teflon tubing using a set of plastic syringe adapters. The cannula should be affixed to the stereotaxic frame so that it is perpendicular to the surgical surface (see video). To disinfect the cannula, fill the 1mL syringe with 70% ethanol and depress the plunger to run the ethanol through the cannula. Repeat this process using sterile saline, and check for any leaks around the cannula joints. To disinfect the outside of the cannula, gently wipe with a 70% ethanol wipe.</li>
	<li>Fill the cannula with sterile saline and then pull back on syringe so that a small air bubble is drawn into the cannula. This air bubble will separate the infusate from the saline in the cannula. Then, backload your infusate (see video). Connect the syringe to the syringe pump and prime the pump by briefly running the cannula.</li>
	<li>Sedate the mouse using an injected anesthetic and prepare the skin by swabbing several times (for 5-10 seconds) with a piece of sterile gauze dipped in the 2% chlorhexidine solution. Ophthalmic ointment should be applied to the mouse to maintain adequate moisture during the procedure. Using a sterile scalpel, create a sagittal incision along the center of the skull, approximately 1.5cm long (see video). The skull surface is then cleaned using a cotton swab soaked in a 3% hydrogen peroxide solution. Take care to avoid getting hydrogen peroxide in the eyes of the mouse. The suture lines of the skull should be apparent at this point: if they are not visible, gently swab the skull with a fresh cotton swab soaked in 3% hydrogen peroxide solution.</li>
	<li>Identify the bregma (Figure 2) and then measure 2mm to the right and 1mm posterior of this structure to locate the infusion site. Using the sterile 22g needle, gently create a hole in the skull at this point by twisting the needle against the skull (see video). Avoid forcing the needle downward against the skull.</li>
	<li>At this point, the mouse should be placed in the stereotaxic frame and begin receiving inhaled anesthetic (see video). Administer the anesthetic at a low level (1%), and carefully monitor the mouse for changes in respiration rate, adjusting the anesthetic accordingly.</li>
	<li>Position the cannula over the skull hole and then lower 3mm below the skull surface. Begin the infusion, using the following rates and durations:
 <table border="0">
		<tbody>
			<tr>
				<td>0.1 &mu;L/min for 5 minutes</td>
				<td rowspan="5"><img src="/files/ftp_upload/1992/1992tablefigure.png" alt="Total Volume 10 ul" /></td>
 		</tr>
 		<tr>
				<td>0.2 &mu;L/min for 5 minutes</td>
			</tr>
 		<tr>
				<td>0.5 &mu;L/min for 5 minutes</td>
			</tr>
 <tr>
				<td>.8 &mu;L/min for 7.5 minutes</td>
			</tr>
			<tr>
 	<td>OFF for 1 minute</td>
			</tr>
	</tbody>
</table>
</li>
	<li>At the end of the infusion, slowly remove the cannula and swab the skull with 3% hydrogen peroxide. Apply sterile bone wax to the hole (see video). Using forceps, draw the skin together over the skull and staple to close. Buprenorphine should be administered for post-operative pain relief.</li>
	<li>Monitor the mouse post-operatively until it regains consciousness and mobility. Due to the length of the procedure, it may take up to an hour for the mouse to regain full activity. During this time place the mouse cage on a heating pad to avoid hypothermia, and do not house the recovering mouse with other active mice.</li>
 <li>Skin staples should be removed one week after surgery.</li>
</ul>
C. In Vivo Imaging
Fluorescently-labeled infusate can be imaged following CED administration, and can be monitored for changes in signal intensity as well as signal location. Typically, it is best to wait 2-3 hours after the infusion to image, so as to allow the mouse to recover from the infusion.
<ul>
	<li>For anesthesia during imaging, use a low level of an inhaled anesthetic. Position the mouse, dorsal side up, in an imaging station (e.g., IVIS Lumina, Caliper Life Sciences, Alameda, CA). Using an appropriate filter setting for the fluor that is being imaged, acquire an image. For CED, a successful infusion should show most of the material in the brain near the infusion site (Figure 3).</li>
</ul>
3. Representative Results
A lack of adverse reaction to administration of therapy is an important indicator of successful injection. For example, following tail vein injection there should be no change in appearance (e.g., size, color) of the tail. A bubble or blister following tail vein injection would indicate subcutaneous, rather than intravenous, delivery of therapy. For intraperitoneal injections, a bump on the skin or discoloration of the abdomen may indicate subcutaneous injection or damage to internal structures. In oral gavage, a mouse with labored breathing or coughing may indicate that fluid was injected into the lungs, rather than into the stomach.
For CED, neurologic function is important for assessing successful administration of therapy. A mouse that is exhibiting seizures or hemiparesis may have received an improper infusion. If a fluorescent infusate is used, in vivo imaging can be applied for assessing successful administration (Figure 3). If the infusate is not localized to the site of injection, the infusion was not successful.
<img src="/files/ftp_upload/1992/1992fig1.jpg" alt="Figure 1" /> Figure 1: CED Cannula and Surgical Set-up. The basic elements of the convection-enhanced delivery surgical set-up are shown. A microinfusion pump (A) is attached to the infusion cannula (B, shown enlarged at top). A stereotaxic frame (C) is used to position the probe. This image does not include heating and anesthetic equipment also used during the procedure. 
<img src="/files/ftp_upload/1992/1992fig2.jpg" alt="Figure 2" /> Figure 2: Mouse Skull Suture Lines. The CED infusion site (red star) can be located by identifying the intersection of the sagittal and coronal sutures (the bregma) and then measuring 2mm lateral and 1mm posterior of the bregma.
<img src="/files/ftp_upload/1992/1992fig3.jpg" alt="Figure 3" /> Figure 3: Representative Results from Successful CED. Liposomes labeled with a far-red fluorescent dye were infused into the mouse brain by CED and imaged both in vivo and ex vivo. A successful infusion shows a fluorescent signal localized to the site of infusion both in vivo (A) and ex vivo (B). The signal should be localized to the infused hemisphere without leakage into the contralateral hemisphere.
<img src="/files/ftp_upload/1992/1992fig4.jpg" alt="Figure 4" /> Figure 4: Representative Image of Successful CED into the Rat Brainstem. Fluorescently-labeled liposomes were infused into the rat brainstem. In rats, as much as 20&mu;L can be infused. The increased thickness of the rat skull and depth of injection precluded in vivo imaging, but correct infusion location could be verified by ex vivo imaging of the dissected brain.

<ol>
	<li>Bobo, R. H., Laske, D. W., Akbasak, A., Morrison, P. F., Dedrick, R. L., Oldfield, E. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Convection-enhanced+delivery+of+macromolecules+in+the+brain.">Convection-enhanced delivery of macromolecules in the brain.</a> Proc. Natl. Acad. Sci. 91, 2076-2080 (1994).</li><li>Degen, J. W., Walbridge, S., Vortmeyer, A. O., Oldfield, E. H., Lonser, R. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Safety+and+efficacy+of+convection-enhanced+delivery+of+gemcitabine+or+carboplatin+in+a+malignant+glioma+model+in+rats.">Safety and efficacy of convection-enhanced delivery of gemcitabine or carboplatin in a malignant glioma model in rats.</a> J Neurosurg. 99, 893-898 (2003).</li><li>Dickinson, P. J., LeCouteur, R. A., Higgins, R. J., Bringas, J. R., Roberts, B., Larson, R. F., Yamashita, Y., Krauze, M. T., Noble, C. O., Drummond, D. C., Kirpotin, D. B., Park, J. W., Berger, M. S., Bankiewicz, K. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Canine+model+of+convection-enhanced+delivery+of+liposomes+containing+CPT-11+monitored+with+real-time+magnentic+resonance+imaging:+laboratory+investigation.">Canine model of convection-enhanced delivery of liposomes containing CPT-11 monitored with real-time magnentic resonance imaging: laboratory investigation.</a> J Neurosurg. 108, 989-998 (2008).</li><li>Gill, S. S., Patel, N. K., Hotton, G. R., O'Sullivan, K., McCarter, R., Bunnage, M., Brooks, D. J., Svendsen, C. N., Heywood, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+brain+infusion+of+glial+cell+line-derived+neurotrophic+factor+in+Parkinson+disease.">Direct brain infusion of glial cell line-derived neurotrophic factor in Parkinson disease.</a> Nature Medicine. 9, 589-595 (2003).</li><li>Jain, R. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Delivery+of+novel+therapeutic+agents+in+tumors:+physiological+barriers+and+strategies.">Delivery of novel therapeutic agents in tumors: physiological barriers and strategies.</a> J Natl Cancer Inst. 81, 570-576 (1989).</li><li>Krauze, M. T., Saito, R., Noble, C. O., Tamas, M., Bringas, J., Park, J. W., Berger, M. S., Bankiewicz, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reflux-free+cannula+for+convection-enhanced+high-speed+delivery+of+therapeutic+agents.">Reflux-free cannula for convection-enhanced high-speed delivery of therapeutic agents.</a> J Neurosurg. 103, 923-929 (2005).</li><li>Krauze, M. T., Vandenberg, S. R., Yamashita, Y., Saito, R., Forsayeth, J., Noble, C. O., Park, J. W., Bankiewicz, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Safety+of+real-time+convection-enhanced+delivery+of+liposomes+to+primate+brain:+a+long-term+retrospective.">Safety of real-time convection-enhanced delivery of liposomes to primate brain: a long-term retrospective.</a> Exp Neurol. 210, 638-644 (2008).</li><li>Murad, G. J., Walbridge, S., Morrison, P. F., Garmestani, K., Degen, J. W., Brechbiel, M. W., Oldfield, E. H., Lonser, R. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Real-time,+image-guided,+convection-enhanced+delivery+of+interleukin+13+bound+to+pseudomonas+exotoxin.">Real-time, image-guided, convection-enhanced delivery of interleukin 13 bound to pseudomonas exotoxin.</a> Clin Cancer Res. 12, 3145-3151 .</li><li>Ozawa, T., James, C. D., Van Meir, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Human+Brain+Tumor+Cell+and+Tumor+Tissue+Transplantation+Models.">Human Brain Tumor Cell and Tumor Tissue Transplantation Models.</a> CNS Cancer: Models, Markers, Prognostic Factors, Targets, and Therapeutic Approaches. , 147-162 (2009).</li><li>Szerlip, N. J., Walbridge, S., Yang, L., Morrison, P. F., Degen, J. W., Jarrell, S. T., Kouri, J., Kerr, P. B., Kotin, R., Oldfield, E. H., Lonser, R. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Real-time+imaging+of+convection-enhanced+delivery+of+viruses+and+virus-sized+particles.">Real-time imaging of convection-enhanced delivery of viruses and virus-sized particles.</a> J Neurosurg. 107, 560-567 (2007).</li><li>Yamashita, Y., Krauze, M. T., Kawaguchi, T., Noble, C. O., Drummond, D. C., Park, J. W., Bankiewicz, K. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Convection-enhanced+delivery+of+a+topoisomerase+I+inhibitor+(nanoliposomal+topotecan)+and+a+topoisomerase+II+inhibitor+(pegylated+liposomal+doxorubicin)+in+intracranial+brain+tumor+xenografts.">Convection-enhanced delivery of a topoisomerase I inhibitor (nanoliposomal topotecan) and a topoisomerase II inhibitor (pegylated liposomal doxorubicin) in intracranial brain tumor xenografts.</a> Neuro Oncol. 9, 20-28 (2007).</li></ol>

All procedures demonstrated here were approved by the UCSF Institutional Animal Care and Use Committee.

Preclinical evaluation of the efficacy of any therapeutic agent must take into account pharmacological properties of the agent and the target tissue of interest. While systemic methods of administration are in general more facile and better tolerated by patients, the selectivity of the blood-brain barrier often necessitates local delivery of therapy for treating CNS disease. Here, we have demonstrated one method of direct delivery to the brain: CED. Other forms of local delivery to the brain include direct administration to the cerebrospinal fluid, intratumoral bolus injection, and ventricular injection. For these methods, diffusibility of therapeutic is critically important, with poor diffusion limiting the region of coverage to a few millimeters from the injection site (Jain 1989, Bobo 1994). In contrast, CED uses positive pressure to increase the area of drug distribution (Bobo 1994), while still enabling therapeutic targeting to specific neuroanatomical structures. In our laboratory, we have successfully performed CED into the brainstem as well as into the caudate putamen in rodents (Figure 4).
The ability to conduct CED in rodent models has become increasingly important in association with increasing interest in maximizing therapeutic efficacy when treating various types of CNS disease. CED can be used to deliver a variety of agents, including purified proteins, small molecule drugs, and viruses (Gill 2003, Degen 2003, Szerlip 2007). The range of diseases that can be treated with CED includes cancers of the CNS (Yamashita 2007) as well as neurodegenerative diseases such as Parkinson's disease(Gill 2003). As CED research continues to expand, the need for in vivo imaging of CED administered therapy is similarly increasing. While the fluorescent imaging demonstrated here would likely not be visible through the human skull, other methods employing co-infusion of MRI contrast agents (Dickinson 2008) are attracting interest for assessing applicability for monitoring infusates in patients with CNS disease.

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		<th>Comment</th>
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<td>2% Chlorhexidine</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>NC9756995</td>
<td>AKA &#x201C;Nolvasan&#x201D;</td>
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<td>20g Needle</td>
<td>&nbsp;</td>
<td>Becton Dickinson</td>
<td>305175</td>
<td>&nbsp;</td>
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<td>28g Needle (with syringe)</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>22-004-270</td>
<td>AKA Insulin Syringe</td>
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<td>3% Hydrogen Peroxide</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>H312P-4</td>
<td>Store away from light</td>
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<td>Cotton Swabs</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>23-400-100</td>
<td>Autoclave before use</td>
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<td>Feather</td>
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<td>No. 21</td>
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<td>Dunlap</td>
<td>HP950</td>
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<td>14-841-20</td>
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<td>&nbsp;</td>
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<td>Upchurch Scientific</td>
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<td>Fits on Luer Lock Syringes</td>
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<td>Silica Tubing</td>
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<td>Polymicro Technologies</td>
<td>2000020</td>
<td>&nbsp;</td>
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<td>Skin Stapler, Staples, Remover</td>
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<td>Stoelting</td>
<td>59020</td>
<td>&nbsp;</td>
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<td>Stereotaxic Frame</td>
<td>&nbsp;</td>
<td>Stoelting</td>
<td>51725</td>
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<td>Teflon Tubing</td>
<td>&nbsp;</td>
<td>Upchurch Scientific</td>
<td>1520</td>
<td>&nbsp;</td>
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<td>Xenogen Lumina</td>
<td>&nbsp;</td>
<td>Caliper Life Sciences</td>
<td>Contact for Quote</td>
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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.

Watch this Scientific Journal Video about Systemic and Local Drug Delivery for Treating Diseases of the Central Nervous System in Rodent Models at JoVE.com

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

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 ...

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Neuroscience

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 ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

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

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

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

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

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

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

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

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 ...

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal dysfunction and loss. Fluorescence-based imaging tools and technologies enable unprecedented analysis of subcellular neurobiological processes, yet there is still a need for unbiased, reproducible, and accessible approaches for extracting quantifiable data from imaging studies. We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-based imaging studies using Drosophila models of neurodegeneration. Specifically, we describe an easy-to-follow, semi-automated approach using Fiji/ImageJ to analyze two cellular processes: first, we quantify protein aggregate content and profile in the Drosophila optic lobe using fluorescent-tagged mutant huntingtin proteins; and second, we assess autophagy-lysosome flux in the Drosophila visual system with ratiometric-based quantification of a tandem fluorescent reporter of autophagy. Importantly, the protocol outlined here includes a semi-automated segmentation step to ensure all fluorescent structures are analyzed to minimize selection bias and to increase resolution of subtle comparisons. This approach can be extended for the analysis of other cell biological structures and processes implicated in neurodegeneration, such as proteinaceous puncta (stress granules and synaptic complexes), as well as membrane-bound compartments (mitochondria and membrane trafficking vesicles). This method provides a standardized, yet adaptable reference point for image analysis and quantification, and could facilitate reliability and reproducibility across the field, and ultimately enhance mechanistic understanding of neurodegeneration.

Quantitative Cell Biology of Neurodegeneration in Drosophila Through Unbiased Analysis of Fluorescently Tagged Proteins Using ImageJ

We have developed a simple and adaptable workflow to extract quantitative data from fluorescence-imaging-based cell biological studies of protein aggregation and autophagic flux in the central nervous system of Drosophila models of neurodegeneration.

With the rising prevalence of neurodegenerative diseases, it is increasingly important to understand the underlying pathophysiology that leads to neuronal ...