Name: a modified method for intrathecal catheterization in rats
Uploaded: 2025-02-14T12:00:00.000+00:00
Duration: 11 min 15 s
Description: Intrathecal catheterization has been widely applied in animal experiments, especially those on neuropathic pain. However, the traditional methods still ...

This work was supported by the National Natural Science Foundation (No. 81971042) and the Key Support Specialist Projects of Shanghai Hongkou District Health Commission (No. HKZK2020A06).

Intrathecal catheterization was carried out in strict accordance with the recommendations in the Guidelines for the Care and Use of Laboratory Animals of the National Institutes of Health, and the protocol was approved by the Ethics Committee of Experimental Animals, China (No. TJBH15523201). Male Sprague-Dawley (SD) rats were used in the experiment. Care was taken to minimize the pain and discomfort of the animals.
1. Material and instrument preparation
NOTE: The preparation of materials and instruments is very important for successful intrathecal catheterization.
<ol>
	<li>Prepare a 15 cm long PE10 tube (the length is determined according to the distance between the rat head and the end of the tail), insert a 20 cm long stainless-steel wire (0.2 mm in diameter) with two polished ends into the PE10 tube as a support, and mark the tube 2 cm from one end to indicate the insertion depth (as marked by black crosses in Figure 1A,B).</li>
	<li>Cut the sharp tip of a 22 G needle and seal the distal end (Figure 1C).</li>
	<li>Cut an epidural catheter (1.0 mm in external diameter) into 1 cm fragments. Then, insert a fragment into a sharp-tip-free 22 G needle (Figure 1D), and seal the distal end of the fragment with a pair of heated straight forceps. This apparatus is called the tube sealing cap (Figure 1E) .</li>
	<li>Prepare a 0.3 cm &#215; 0.5 cm antiallergic band by cutting a silk tape (1.25 cm &#215; 9.1 m) with a pair of scissors (Figure 1F).</li>
</ol>
2. Preparation for surgery
<ol>
	<li>Prepare the instruments for intrathecal catheterization by sterilizing them before surgery. The instruments used for the surgery are toothed forceps, scissors, a gavage apparatus, a scalpel handle and #10 blades. (Figure 2).</li>
	<li>Immerse the PE10 tube and guide wire in 75% ethanol for sterilization for approximately 2 h.</li>
</ol>
3. Surgery
<ol>
	<li>Anesthetize the rat&#160;with 3% isoflurane at a flow rate of 3 L/min.</li>
	<li>Place the rat on the operating table and observe the withdrawal reflex when pinching the hind paw with forceps. The absence of hind paw movement in response to stimulation confirmed successful anesthesia. Administer adequate analgesia by intramuscular injection of 1 mg/kg meloxicam before intrathecal catheterization.</li>
	<li>Remove hair from the lumbar spine region of the back and the area between two ears with a shaver.</li>
	<li>Place a centrifuge tube (3 cm in diameter) under the abdomen of the&#160;rat at the waist-hip junction to increase the flexion in the lumbar spine, giving more room for the needle and the catheter to pass through.</li>
	<li>Sterilize the surgical sites (the area over the lumbar spine region and the area between two ears) with povidone-iodine solution&#160;and then with ethanol solution three times. Cover the rat with an aseptic dressing, and expose the surgical sites. Then wash the PE10 tube and guide wire with normal saline before surgery. 
	NOTE: The tail was not covered so that tail movement could be observed during intrathecal catheterization.</li>
	<li>Determine the location of the intervertebral space between L5 and L6 by locating the spinous process of L6 at the midpoint between the left and right bilateral iliac crests. Fix the skin with the left thumb and the left index finger of the operator, and then make a 3-4 cm long midline incision just above the spinous process between L4 and S1.</li>
	<li>Bluntly separate the subcutaneous tissues with a pair of scissors. Locate the intervertebral space between L5 and L6 again and make a small incision (0.3 - 0.5 cm) on both sides of the L5 and L6 dorsal processes.</li>
	<li>Clamp and lift the L5 dorsal process with a pair of toothed forceps to expand the intervertebral space. Then, bluntly separate the muscles around the vertebral body with a pair of scissors until the top of the L6 dorsal process is completely exposed. 
	NOTE: Removal of any part of the vertebral body and muscles should be avoided, with the aim of minimizing damage to surrounding tissues.</li>
	<li>When the L5 dorsal process is lifted with a pair of toothed forceps and the intervertebral space is expanded with another pair of forceps, clean the L5-6 intervertebral space with a cotton ball until the inverted &#34;V&#34; area is completely exposed.</li>
	<li>Puncture the spine with a 23 G needle in the inverted &#34;V&#34; area just under the top of the L6 dorsal process. 
	NOTE: A tail flick is observed, and/or colorless transparent fluid flows out from the subarachnoid space, indicating a successful puncture into the subarachnoid space.</li>
	<li>Carefully insert the PE10 tube containing stainless-steel wire into the spinal canal at the puncture site, being tilted 30&#176; toward the tail. Adjust the insertion angle until the PE10 tube can be successfully inserted without resistance (a tail flick was observed during this process).</li>
	<li>When the marked area of the PE10 tube reaches the posterior muscle, catheterization is stopped.</li>
	<li>Slowly remove the stainless wire from the PE10 tube. A tail flick may be observed. 
	NOTE: A tail flick may be observed, and after the wire is removed, transparent fluid (or light red fluid) may flow out of the tube.</li>
	<li>Then, connect the PE10 tube to a 1 mL syringe, through which 20 &#181;L of normal saline is injected. After the syringe is removed, saline will flow continuously out of the PE10 tube, indicating that it has been successfully inserted into the subarachnoid space.</li>
	<li>Once the PE10 tube is confirmed to be unobstructed, suture the muscles on one side of the vertebral body with a 4-0 suture and make a knot. Then, tie the suture around the PE10 tube and make another knot. Do not cut the suture; suture the muscles on the other side; tie the suture to the PE10 tube again, make a third knot, and cut the suture. 
	NOTE: This process fixes PE10 tube with a figure-8 suture to reduce the possibility of displacement and retraction of the tube.</li>
	<li>Make a 0.5 cm long incision 1 cm below the midpoint between the ears. Bluntly separate the subcutaneous tissues with scissors, and insert a metal gavage tube toward the tail until the tip is visible in the lumbar incision.</li>
	<li>Insert the distal end of the PE10 tube into the gavage tube until the PE10 tube exits the other end of the gavage tube; then gently withdraw the gavage.</li>
	<li>When the PE10 tube is confirmed to be unobstructed again, suture the remaining muscles around the lumbar incision with a 4-0 suture, tie the suture around the PE10 tube, and make another knot to fix the PE10 tube again.</li>
	<li>Suture the skin, avoiding damage to the PE10 tube. Then, suture the neck skin with a 4-0 suture, tie the suture around the PE10 tube, and make a knot to fix the PE10 tube.</li>
	<li>When the PE10 tube is confirmed to be unobstructed again, seal the extracorporeal end of the PE10 tube with a sealing cap.</li>
	<li>Dry the PE10 tube with a piece of tissue and then tie the antiallergic band around the PE10 tube several times to prevent retraction of the PE10 tube during rat movement.</li>
</ol>
4. Lidocaine validation experiment
<ol>
	<li>After the surgery, return the rat to its cage (one per cage) and closely monitor it during the recovery from anesthesia until the rat gains consciousness.</li>
	<li>After the rat is fully awake, remove the sealing cap and inject 20 &#181;L of 2% lidocaine into PE10 tube at a rate of 0.02 mL/s via a Hamilton syringe, followed by the injection of 10 &#181;L of normal saline.</li>
	<li>Seal the PE10 tube with the sealing cap.</li>
	<li>Place the rat on a table and observe carefully. The presence of hind limb paralysis after an intrathecal injection of lidocaine (from PE10 tube) indicates successful catheterization (Figure 3). Hind limb paralysis usually lasts approximately 30 min10. 
	NOTE: Allow the rat to recover for 5-7 days before the following experiments.</li>
	<li>Closely monitor the rat during the recovery period until complete recovery of limb function.</li>
</ol>

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The authors of this manuscript declare that there are no conflicts of interest.

There are several critical tips for this modified method to maximize the success rate of intrathecal catheterization. First, a 20 cm long stainless-steel wire with two polished ends should be prepared and inserted into PE10 tube as a support. Second, the operator should completely expose the inverted &#34;V&#34;&#160;area after cleaning the L5-6 intervertebral space with a cotton ball, and the intervertebral space should be expanded with another forceps while lifting the L5 dorsal process with toothed forceps. Third, PE10 tube should be fixed with a figure-8 suture four times. Finally, the extracorporeal end of PE10 tube should be tied with a band and sealed with a self-made cap.
The success rate of intrathecal catheterization and damage to tissues around vertebral bodies may significantly influence the reliability of the experimental results15. Thus, improving the success rate as much as possible and reducing the damage to the surrounding tissues are&#160;crucial in establishing animal models and relevant experiments1. In this modified method, a stainless-steel wire is inserted into PE10 tube for guidance, which increases the elasticity of the tube and improves the success rate of intrathecal catheterization. Moreover, the amount of space needed for operation is reduced with this modified method, and the damage to the tissues around the lumbar spine is minimized because the surrounding tissues are bluntly separated, but not cut. In comparison, in the previously reported method11, a 20 G guide cannula is used to reduce resistance during puncturing, and repeated puncturing is often needed, which may injure the tissues. In addition, in the previously reported method, to reduce the diameter of PE10 tube, it is immersed in warm water (60 &#176;C) and then stretched at one end to approximately 150% of the original length, which may not ensure the consistency of tube diameter and therefore may cause leakage of cerebrospinal fluid because the diameter of 20G guide cannula is approximately two times greater than or equal to that of the stretched PE10 tube. Moreover, in our method, lumbar function is preserved to the greatest extent, which avoids the influence of surgery on the results of subsequent experiments. These results are consistent with those reported by Xu et al2.
In previously reported method11, the length of PE10 tube is approximately 14 cm if the tube is fixed at the site around the puncture site, but the catheter indwelling time is often shorter than 7 days under these conditions (or the tube is removed from the body by the rat). The length of PE10 tube is approximately 28 cm if the tube is fixed at the back of the neck, which is significantly longer than the PE10 tube used in our method (15 cm). Although beads were formed following the protocol reported by St&#248;rkson et al.11, some tubes were removed from the body, and only approximately 65% of the tubes were still fixed in place at 7 days after surgery, which significantly affected the outcomes of subsequent experiments. In our method, PE10 tube is fixed with a figure-8 suture 4 times, and the extracorporeal end of PE10 tube is tied with a band to reduce the possibility of displacement and retraction. Following our method, approximately 85% of the tubes remained in place at 7 days after surgery, and approximately 80% of the tubes remained in place at 28 days after surgery.
In the previously reported method11, the extracorporeal tip of the intrathecal catheter should be cut off for each drug administration. However, the repeated intrathecal administration of drugs may shorten the catheter indwelling time, which makes the intrathecal administration of drugs inconvenient. Therefore, in our method, a self-made cap is used to seal PE10 tube, which is sterilized with ethanol once daily. This not only prevents the leakage of cerebrospinal fluid but also reduces the need for repeated cutting of PE10 tube for intrathecal administration of drugs, ensuring the effective delivery of the drugs.
The advantages and disadvantages of the modified method and previously reported method are summarized in Table 1. First, for the modified method, the use of stainless-steel wire in PE10 tube increases the elasticity of the tube and improves the success rate of intrathecal catheterization, the amount of space needed for the operation is reduced, and the damage to the tissues around the lumbar spine is minimized. In previously reported method, a 20G guide cannula is inserted until resistance is felt, and repeated puncture is often needed, which may result in damage to tissues. In addition, the PE10 tube at one end is stretched until its length reaches approximately 150% of original length, which may cause cerebrospinal fluid leakage because the diameter of the 20G guide cannula is 2 times greater than or equal to that of stretched PE10 tube. Second, in the modified method, the length of PE10 tube is determined before surgery, and the catheter indwelling time can be longer than one week. In previously reported method, the length of PE10 tube is approximately 14 cm if it is fixed at the puncture site, but the catheter indwelling time is often shorter than 7 days because the tube is susceptible to being pulled out of the body by the rat; the length of PE10 tube is approximately 28 cm if it is fixed at the back of the neck, which is significantly longer than the length of the tube used in our method. Third, in the modified method, the PE10 tube is fixed with a figure-8 suture 4 times to prevent tube movement and retraction; a self-made cap is used to seal PE10 tube, which not only prevents leakage of cerebrospinal fluid but also prevents the need for repeated cutting of PE10 tube. In the previously reported method, it is difficult to obtain beads with a consistent diameter, the displacement of PE10 tube is common when beads are formed, and repeated cutting of PE10 tube is often needed. Finally, in the modified method, the extracorporeal end of PE10 tube is tied with a band, which prevents the tube from retracting during movement. However, in the prior method, the beads cannot reliably prevent PE10 tube retraction because it is difficult to obtain beads with a consistent diameter.
Overall, this modified method for intrathecal catheterization has the following advantages. First, the use of stainless-steel wire in PE10 tube increases the elasticity of the tube and improves the success rate of intrathecal catheterization, the amount space needed for the operation is reduced, and damage to the tissues around the lumbar spine is minimized, which preserves the lumbar function to the greatest extent possible and avoids the influence of surgery on the results of subsequent experiments. Second, PE10 tube is fixed with a figure-8 suture 4 times, which prevents tube movement and retraction during movement. Third, a self-made sealing cap is used to seal PE10 tube, which not only prevents cerebrospinal fluid leakage but also prevents the need for repeated cutting of PE10 tube. Repeated cutting of the catheter may shorten the catheter, which makes the delivery of drugs inconvenient. Finally, the extracorporeal end of PE10 tube is tied with an antiallergic band, which prevents the tube from retracting during movement.
However, there are several limitations in this modified intrathecal catheterization technique. First, after surgery, the rats need to be housed separately (one per cage) to avoid damage to the extracorporeal end of PE10 tube. Second, recovery for 5-7 days after intrathecal injection of lidocaine is needed before subsequent experiments.
In conclusion, this modified method for intrathecal catheterization may serve as a useful tool for the repetitive intrathecal administration of drugs and represent&#160;a simple, convenient, and reliable way to shorten the duration of experiments.

Intrathecal catheterization (also known as subarachnoid catheterization) in rats is a method that involves inserting a catheter into the subarachnoid space through the intervertebral space1. Drugs are directly injected into the subarachnoid space through the catheter, which helps researchers investigate the effects of drugs on the spinal cord without considering the effects of drugs that penetrate the blood-brain barrier2,3. Moreover, cerebrospinal fluid can be collected following intrathecal catheterization to investigate the microenvironment of the central nervous system4,5. The currently used method for intrathecal catheterization was first established by Yaksh and Rudy6 in 1976, and since then, it has been widely applied in animal experiments in the fields of neuroscience, anesthesia and analgesia, spinal cord-mediated cardiovascular regulation, and especially neuropathic pain2,7. However, this method still has several limitations, such as a high incidence of spinal cord damage, subarachnoid hemorrhage, postoperative sensory and motor dysfunction, high postoperative mortality, and a high risk of neurological impairment4,5,8,9,10. In an attempt to overcome these limitations, catheterization of the subarachnoid space through lumbar interspaces was proposed by St&#248;rkson et al. in 199611, and a higher postsurgical success rate was reported. Notably, the fixation of indwelling catheter is still a challenge in this method, and catheter retraction is common due to animal movement, which makes intrathecal drug administration inconvenient.
Because of the above limitations, some investigators12,13,14,15 have attempted to improve the tools for puncturing, the methods of catheterization, and the methods of catheter fixation, but the available methods still need to be modified due to the difficulty&#160;in quantifying the diameter of the beads used, the need for repeated punctures and the short length of the catheter, etc.11
According to the lumbar approach for intrathecal catheterization1 and the Seldinger technique for central vein catheterization,16 we developed a method for intrathecal catheterization in rats that uses a stainless-steel wire, a self-made sealing cap, and an antiallergic band to simplify the existing method. Through this method, the catheter can be easily inserted into the subarachnoid space and stably fixed on the back of the rat, and the need for repeated punctures for repeated intrathecal drug administration is avoided.
Herein, we introduce a modified method that may improve the success rate of intrathecal catheterization in rats and represent&#160;a simple, convenient, and reliable approach for repetitive intrathecal administration of drugs.

<table><tbody><tr><td>1 cc syringe</td><td></td><td>Jiangxi Hongda Medical Equipment Co., Ltd</td><td>1 cc</td></tr><tr><td>22 gauge &amp;times; 1&amp;rdquo; needles</td><td></td><td>Jiangxi Hongda Medical Equipment Co., Ltd</td><td>22G</td></tr><tr><td>23 gauge &amp;times; 1&amp;rdquo; needles</td><td></td><td>Jiangxi Hongda Medical Equipment Co., Ltd</td><td>23G</td></tr><tr><td>25 &amp;mu;L Hamilton Syringes</td><td></td><td>Shanghai Bolige Co.,Ltd</td><td>0.31mm 25 &amp;mu;L&amp;nbsp;</td></tr><tr><td>4-O MERSILK NON-ABSORBABLE SUTURE</td><td></td><td>ETHICON</td><td>SA83G</td></tr><tr><td>50 mL corning centrifuge tubes 3 cm diameter</td><td>430820</td><td>CORNING</td><td></td></tr><tr><td>Epidural catheter and connector</td><td></td><td>Henan Tuoren Medical Device Co., Ltd</td><td>regular type</td></tr><tr><td>Gavage apparatus</td><td></td><td>Shanghai Bolige Co.,Ltd</td><td>8#</td></tr><tr><td>PE-10 Mirco Medical Tubing</td><td>BB31695-PE/1</td><td>Scientific Commodities, Inc</td><td></td></tr><tr><td>Scalpel handle and #10 blades</td><td></td><td>Jiangsu Songxin Medical Equipment Co., Ltd</td><td>125mm</td></tr><tr><td>Scissors</td><td></td><td>Jiangsu Songxin Medical Equipment Co., Ltd</td><td>100mm</td></tr><tr><td>Sprague-Dawley (SD) rats</td><td></td><td>Shanghai BK/KY Biotechnology Co., Ltd</td><td>Male</td></tr><tr><td>Stainless steel wire 0.2 mm diameter</td><td></td><td>Dongguan Jiazhi Metal Products Technology Co., Ltd.</td><td>0.2mm&amp;nbsp; &amp;times;&amp;nbsp; 1m</td></tr><tr><td>Toothed forceps</td><td></td><td>Jiangsu Songxin Medical Equipment Co., Ltd</td><td>18cm</td></tr><tr><td>URGO silk tape</td><td></td><td>URGO</td><td>1.25cm &amp;times; 9.1m</td></tr></tbody></table>

a modified method for intrathecal catheterization in rats

Intrathecal catheterization has been widely applied in animal experiments, especially those on neuropathic pain. However, the traditional methods still have&#160;several limitations. Although some investigators have attempted to improve the traditional methods, the available methods still need to be modified. Herein, we introduce a modified method for intrathecal catheterization in rats.
This method uses a 20 cm long stainless-steel wire (0.2 mm in diameter), a 15 cm long plastic PE10 tube, a self-made sealing cap, and a 0.3 cm &#215; 0.5 cm anti-allergic band. Our modified method for intrathecal catheterization has several advantages. First, introducing a stainless-steel wire to PE10 tube increases the elasticity of the tube, improves the success rate of intrathecal catheterization, reduces the amount of space required for the operation, and minimizes the damage to the tissues around the lumbar spine. Second, the length of PE10 tube is determined before the surgery, and catheter indwelling time can be longer than one week. Third, the PE10 tube is fixed by a figure-8 suture, 4 times, which prevents tube movement and retraction when the animal moves. Fourth, a&#160;self-made sealing cap is used to seal the PE10 tube, which not only prevents cerebrospinal fluid leakage but also reduces the need for repeated cutting of PE10 tube. Finally, the extracorporeal end of PE10 tube is tied with a band, which prevents tube retraction when the animal moves.
This method can increase the catheterization success rate in rats, as approximately 80% of PE10 tubes remained in place even 28 days after surgery. Thus, this modified method may represent a simple, convenient, and reliable approach for repetitive intrathecal drug administration.

For intrathecal injection, the extracorporeal tip of PE10 tube was cut off, and PE10 tube was sealed with a sealing cap between two drug injections. In our pilot study, the success rate of intrathecal catheterization was approximately 95% (19 out of 20 rats); success was indicated by a tail flick and/or the release of colorless transparent fluid during the procedure. Approximately 85% of the tubes remained in place 7&#160;days after surgery, and approximately 80% remained in place 28 days after surgery. The rats recovered soon after the operation, and no complications were observed within 7 days after surgery. The daily movement was normal, and behavioral abnormality was not observed. These results indicate our method is superior to those previously reported in terms of the success rate and long-indwelling rate.
Complete paralysis of the lower limbs upon lidocaine injection via catheters indicates successful intrathecal catheterization15. The success rate of intrathecal catheterization is calculated by dividing the total number of rats by the number of rats with successful catheterization. With our modified method, the success rate was 95%, which was greater than the rate achieved with the method reported by Hou et al. (88%)15. This is shown in Figure 4.
The intrathecal tube was monitored at 2, 5, and 7 days after intrathecal catheterization and the rate of successful indwelling catheter was calculated as number of rats with successful indwelling catheter/total number of rats &#215; 100%. At&#160;2, 5, and 7 days after intrathecal catheterization, the rate of successful indwelling catheter was 94%, 81%, and 65%, respectively, in the study of St&#216;rkson et al.11. The rate of successful indwelling catheter at 2, 5 and 7 days after intrathecal catheterization was 95%, 90% and 85%, respectively, with our technique (Figure 5).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66487/66487fig1.jpg" /> 
Figure 1. Materials and instruments used for intrathecal catheterization. (A)&#160;A 15-cm long PE10 tube was prepared, and the tube was marked 2 cm from one end to indicate the insertion depth. (B)&#160;A 20-cm long stainless-steel wire with two polished ends was inserted into the PE10 tube as a support. (C)&#160;The sharp tip of the 22G needle was cut with a pair of scissors, and the distal end was sealed with a pair of forceps. (D) An epidural catheter (1.0 mm in external diameter) was cut into 1-cm fragments, which were then inserted into the sharp-tip-free 22G needle. (E)&#160;The distal end of the epidural catheter was sealed with a pair of heated straight forceps; this apparatus was called the tube sealing cap. (F)&#160;A 0.3 cm &#215; 0.5 cm anti-allergic band (silk tape, 1.25 cm &#215; 9.1 m) was prepared with a pair of scissors. <a href="https://www.jove.com/files/ftp_upload/66487/66487fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66487/66487fig2.jpg" /> 
Figure 2. Preparation of instruments for intrathecal catheterization. Instruments (such as toothed forceps, scissors, a gavage apparatus, a scalpel handle, and #10 blades) were sterilized with ethanol for approximately 2 h and then washed with normal saline approximately 30 min before surgery. <a href="https://www.jove.com/files/ftp_upload/66487/66487fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66487/66487fig3.jpg" /> 
Figure 3. Results of the lidocaine validation experiment after intrathecal catheterization. After intrathecal injection of 20 &#181;L&#160;of 2% lidocaine followed by injection of 10 &#181;L of normal saline, the rat was&#160;temporarily paralyzed: lower limb paralysis occurred within 30 s and disappeared 30 min later, indicating successful intrathecal catheterization. <a href="https://www.jove.com/files/ftp_upload/66487/66487fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66487/66487fig4.jpg" /> 
Figure 4. Comparison of the rate of successful catheterization between our modified method and a previously reported method. <a href="https://www.jove.com/files/ftp_upload/66487/66487fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/66487/66487fig5.jpg" /> 
Figure 5. Comparison of the long-indwelling catheter rate between our modified method and a previously reported method. <a href="https://www.jove.com/files/ftp_upload/66487/66487fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>&#12288;</td>
			<td>Modified method</td>
			<td>Prior method</td>
			<td>Advantages of modified method</td>
			<td>Disadvantages of prior method</td>
		</tr>
		<tr>
			<td rowspan="4">Guiding method for insertion</td>
			<td rowspan="4">a stainless-steel wire</td>
			<td rowspan="4">Guide-canula (20G 0.9&#215; 38 mm)</td>
			<td>Increases the elasticity of the tube,</td>
			<td>The resistance is difficult to feel, increasing the difficulty of operation</td>
		</tr>
		<tr>
			<td>Improves the success rate of intrathecal catheterization</td>
			<td>Damage to the tissues due to repeated puncture</td>
		</tr>
		<tr>
			<td>Reduces the requirement of operative space</td>
			<td>One end of the tube is stretched to 1.5 times original length, making the diameter of both ends different</td>
		</tr>
		<tr>
			<td>Minimizes the damage to tissues around the lumbar spine</td>
			<td>Susceptibility to cerebrospinal fluid leakage because the diameter of 20G guide-canula is 2 times or more that of stretched PE10 tube</td>
		</tr>
		<tr>
			<td>Length of PE10 tube</td>
			<td>15 cm</td>
			<td>14 or 28 cm</td>
			<td>Easy to determine the length of PE10 tube regardless of the duration of catheter indwelling</td>
			<td>Duration of catheter indwelling is shorter for shorter PE10 tube; susceptibility to falling out of the body for long PE10 tube</td>
		</tr>
		<tr>
			<td>Fixation method</td>
			<td>&#34;8&#34; suture and 4 times</td>
			<td>1 or 2 beads</td>
			<td>Avoids the tube movement and retraction during the animal activities</td>
			<td>Difference in the diameter of the tube at both ends, and susceptibility to displacement of PE10 tube during the bead-making </td>
		</tr>
		<tr>
			<td rowspan="2">Method for tube sealing</td>
			<td rowspan="2">Self-made cap</td>
			<td rowspan="2">No</td>
			<td>Prevents the leakage of cerebrospinal fluid </td>
			<td rowspan="2">Requirement of repeated cutting of PE10 tube </td>
		</tr>
		<tr>
			<td>Avoids the repeated cutting of PE10 tube </td>
		</tr>
		<tr>
			<td>Method for prevention of retraction</td>
			<td>Anti-allergic band</td>
			<td>1 or 2 beads</td>
			<td>Prevents the tube from retraction during the animal activities </td>
			<td>Susceptibility to retraction</td>
		</tr>
	</tbody>
</table>
Table 1.&#160;Advantages and disadvantages of the modified method and a previously reported method.

Herein, we introduce a modified method for intrathecal catheterization in rats that represents a simple, convenient, and reliable approach for repetitive intrathecal drug administration.

A Modified Method for Intrathecal Catheterization in Rats

Intrathecal catheterization has been widely applied in animal experiments, especially those on neuropathic pain. However, the traditional methods still ...

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Research

JoVE Journal

Medicine

524 Views.  Tongji University. Herein, we introduce a modified method for intrathecal catheterization in rats that represents a simple, convenient, and reliable approach for repetitive intrathecal drug administration.

Video: A Modified Method for Intrathecal Catheterization in Rats

A General Method for Evaluating Deep Brain Stimulation Effects on Intravenous Methamphetamine Self-Administration

Substance use disorders, particularly to methamphetamine, are devastating, relapsing diseases that disproportionally affect young people. There is a need for novel, effective and practical treatment strategies that are validated in animal models. Neuromodulation, including deep brain stimulation (DBS) therapy, refers to the use of electricity to influence pathological neuronal activity and has shown promise for psychiatric disorders, including drug dependence. DBS in clinical practice involves the continuous delivery of stimulation into brain structures using an implantable pacemaker-like system that is programmed externally by a physician to alleviate symptoms. This treatment will be limited in methamphetamine users due to challenging psychosocial situations. Electrical treatments that can be delivered intermittently, non-invasively and remotely from the drug-use setting will be more realistic. This article describes the delivery of intracranial electrical stimulation that is temporally and spatially separate from the drug-use environment for the treatment of IV methamphetamine dependence. Methamphetamine dependence is rapidly developed in rodents using an operant paradigm of intravenous (IV) self-administration that incorporates a period of extended access to drug and demonstrates both escalation of use and high motivation to obtain drug.

This article describes the delivery of intracranial electrical stimulation that is temporally and spatially separate from the drug-use environment for the treatment of IV methamphetamine dependence.

Substance use disorders, particularly to methamphetamine, are devastating, relapsing diseases that disproportionally affect young people.  There is a need ...

Behavior

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 ...

Neuroscience

Rat Model of Widespread Cerebral Cortical Demyelination Induced by an Intracerebral Injection of Pro-Inflammatory Cytokines

Multiple sclerosis (MS) is the most common immune-mediated disease of the central nervous system (CNS) and progressively leads to physical disability and death, caused by white matter lesions in the spinal cord and cerebellum, as well as by demyelination in grey matter. Whilst conventional models of experimental allergic encephalomyelitis are suitable for the investigation of the cell-mediated inflammation in the spinal and cerebellar white matter, they fail to address grey matter pathologies. Here, we present the experimental protocol for a novel rat model of cortical demyelination allowing the investigation of the pathological and molecular mechanisms leading to cortical lesions. The demyelination is induced by an immunization with low-dose myelin oligodendrocyte glycoprotein (MOG) in an incomplete Freund&#39;s adjuvant followed by a catheter-mediated intracerebral delivery of pro-inflammatory cytokines. The catheter, moreover, enables multiple rounds of demyelination without causing injection-induced trauma, as well as the intracerebral delivery of potential therapeutic drugs undergoing a preclinical investigation. The method is also ethically favorable as animal pain and distress or disability are controlled and relatively minimal. The expected timeframe for the implementation of the entire protocol is around 8 - 10 weeks.

The protocol presented here allows the reproduction of a widespread grey matter demyelination of both cortical hemispheres in adult male Dark Agouti rats. The method comprises of intracerebral implantation of a catheter, subclinical immunization against myelin oligodendrocyte glycoprotein, and intracerebral injection of a pro-inflammatory cytokine mixture through the implanted catheter.

Multiple sclerosis (MS) is the most common immune-mediated disease of the central nervous system (CNS) and progressively leads to physical disability and ...

Intrathecal Delivery of Antisense Oligonucleotides in the Rat Central Nervous System

The blood brain barrier (BBB) is an important defense against the entrance of potentially toxic or pathogenic agents from the blood into the central nervous system (CNS). However, its existence also dramatically lowers the accessibility of systemically administered therapeutic agents to the CNS. One method to overcome this, is to inject those agents directly into the cerebrospinal fluid (CSF), thus bypassing the BBB. This can be done via implantation of a catheter for either continuous infusion using an osmotic pump, or for single bolus delivery. In this article, we describe a surgical protocol for delivery of CNS-targeting antisense oligonucleotides (ASOs) via a catheter implanted directly into the cauda equina space of the adult rat spine. As representative results, we show the efficacy of a single bolus ASO intrathecal (IT) injection via this catheterization system in knocking down the target RNA in different regions of the rat CNS. The procedure is safe, effective and does not require expensive equipment or surgical tools. The technique described here can be adapted to deliver drugs in other modalities as well.

Here, we describe a method for delivering drugs to the rat central nervous system by implanting a catheter into the lumbar intrathecal space of the spine. We focus on the delivery of antisense oligonucleotides, though this method is suitable for delivery of other therapeutic modalities as well.

The blood brain barrier (BBB) is an important defense against the entrance of potentially toxic or pathogenic agents from the blood into the central ...

In Vivo Intracellular Recording of Type-Identified Rat Spinal Motoneurons During Trans-Spinal Direct Current Stimulation

Intracellular recording of spinal motoneurons in vivo provides a &#8220;gold standard&#8221; for determining the cells&#8217; electrophysiological characteristics in the intact spinal network and holds significant advantages relative to classical in vitro or extracellular recording techniques. An advantage of in vivo intracellular recordings is that this method can be performed on adult animals with a fully mature nervous system, and therefore many observed physiological mechanisms can be translated to practical applications. In this methodological paper, we describe this procedure combined with externally applied constant current stimulation, which mimics polarization processes occurring within spinal neuronal networks. Trans-spinal direct current stimulation (tsDCS) is an innovative method increasingly used as a neuromodulatory intervention in rehabilitation after various neurological injuries as well as in sports. The influence of tsDCS on the nervous system remains poorly understood and the physiological mechanisms behind its actions are largely unknown. The application of the tsDCS simultaneously with intracellular recordings enables us to directly observe changes of motoneuron membrane properties and characteristics of rhythmic firing in response to the polarization of the spinal neuronal network, which is crucial for the understanding of tsDCS actions. Moreover, when the presented protocol includes the identification of the motoneuron with respect to an innervated muscle and its function (flexor versus extensor) as well as the physiological type (fast versus slow) it provides an opportunity to selectively investigate the influence of tsDCS on identified components of spinal circuitry, which seem to be differently affected by polarization. The presented procedure focuses on surgical preparation for intracellular recordings and stimulation with an emphasis on the steps which are necessary to achieve preparation stability and reproducibility of results. The details of the methodology of the anodal or cathodal tsDCS application are discussed while paying attention to practical and safety issues.

This protocol describes in vivo intracellular recording of rat lumbar motoneurons with simultaneous trans-spinal direct current stimulation. The method enables us to measure membrane properties and to record rhythmic firing of motoneurons before, during and after anodal or cathodal polarization of the spinal cord.

Intracellular recording of spinal motoneurons in vivo provides a &#8220;gold standard&#8221; for determining the cells&#8217; electrophysiological ...

Pre-Chiasmatic, Single Injection of Autologous Blood to Induce Experimental Subarachnoid Hemorrhage in a Rat Model

Despite advances in treatment over the last decades, subarachnoid hemorrhage (SAH) continues to carry a high burden of morbidity and mortality, largely afflicting a fairly young population. Several animal models of SAH have been developed to investigate the pathophysiological mechanisms behind SAH and to test pharmacological interventions. The pre-chiasmatic, single injection model in the rat presented in this article is an experimental model of SAH with a predetermined blood volume. Briefly, the animal is anesthetized, intubated, and kept under mechanical ventilation. Temperature is regulated with a heating pad. A catheter is placed in the tail artery, enabling continuous blood pressure measurement as well as blood sampling. The atlantooccipital membrane is incised and a catheter for pressure recording is placed in the cisterna magna to enable intracerebral pressure measurement. This catheter can also be used for intrathecal therapeutic interventions. The rat is placed in a stereotaxic frame, a burr hole is drilled anteriorly to the bregma, and a catheter is inserted through the burr hole and placed just anterior to the optic chiasm. Autologous blood (0.3 mL) is withdrawn from the tail catheter and manually injected. This results in a rise of intracerebral pressure and a decrease of cerebral blood flow. The animal is kept sedated for 30 min and given subcutaneous saline and analgesics. The animal is extubated and returned to its cage. The pre-chiasmatic model has a high reproducibility rate and limited variation between animals due to the pre-determined blood volume. It mimics SAH in humans making it a relevant model for SAH research.

Subarachnoid hemorrhage continues to carry a high burden of mortality and morbidity in man. To facilitate further research into the condition and its pathophysiology, a pre-chiasmatic, single injection model is presented.

Despite advances in treatment over the last decades, subarachnoid hemorrhage (SAH) continues to carry a high burden of morbidity and mortality, largely ...

Microsurgical Skills of Establishing Permanent Jugular Vein Cannulation in Rats for Serial Blood Sampling of Orally Administered Drug

Blood sampling in small laboratory animals is necessary for pharmaceutical lead optimization but can cause great harm and stress to experimental animals, which could potentially affect results. The jugular vein cannulation (JVC) in rats is a widely used model for repeated blood collection but requires adequate training of surgery skills and animal care. This article details the microsurgical procedures for establishing and maintaining a permanent JVC rat model with specific focus on the placement and sealing of the jugular cannula. The importance of monitoring physiological (e.g., body weight, food, and water intake) and hematological parameters, was highlighted with results presented for 6 days post-surgery during the rat&#39;s recovery. The drug-plasma concentration-time profile of orally administered natural phenol ellagic acid was determined in the JVC rat model.

Detailed microsurgical techniques are demonstrated to establish a longer-term jugular vein cannulation rat model for sequential blood collection in the same animal. Physiological and hematological parameters have been monitored during the rat's recovery phase. This model has been applied to study pharmacokinetics of orally administered polyphenol without inducing animal stress.

Blood sampling in small laboratory animals is necessary for pharmaceutical lead optimization but can cause great harm and stress to experimental animals, ...

Lumbar Puncture Delivery of AAV9 in Juvenile Rats for CNS Gene Therapy

Intrathecal Vector Delivery in Juvenile Rats via Lumbar Cistern Injection

Gene therapy is a powerful technology to deliver new genes to a patient for the treatment of disease, be it to introduce a functional gene, inactivate a toxic gene, or provide a gene whose product can modulate the biology of the disease. The delivery method for the therapeutic vector can take many forms, ranging from intravenous infusion for systemic delivery to direct injection into the target tissue. For neurodegenerative disorders, it is often desirable to skew transduction towards the brain and/or spinal cord. The least invasive approach to target the entire central nervous system involves injection into the cerebrospinal fluid (CSF), allowing the therapeutic to reach a large fraction of the central nervous system. The safest approach to deliver a vector into the CSF is the lumbar intrathecal injection, where a needle is introduced into the lumbar cistern of the spinal cord. This technique, also known as a lumbar puncture, has been widely used in neonatal and adult rodents and in large animal models. While the technique is similar across species and developmental stages, subtle differences in size, structure, and elasticity of tissues surrounding the intrathecal space require accommodations in the approach. This article describes a method for performing lumbar puncture in juvenile rats to deliver an adeno-associated serotype 9 vector. Here, 25-35 &#181;L of vector were injected into the lumbar cistern, and a green fluorescent protein (GFP) reporter was used to evaluate the transduction profile resulting from each injection. The benefits and challenges of this approach are discussed.

Author Spotlight: Assessing Intrathecal Gene Therapy Efficacy in Juvenile Rats

A surgical procedure is described to perform injections into the lumbar cistern of the juvenile rat. This approach has been used for the intrathecal delivery of gene therapy vectors, but it is anticipated that this approach can be used for a variety of therapeutics, including cells and drugs.

Gene therapy is a powerful technology to deliver new genes to a patient for the treatment of disease, be it to introduce a functional gene, inactivate a ...

Spinal Cord Surgery and Optical Shank Implantation in Rodents

Implantation of Optoelectronic Devices in the Rodent Spinal Cord

Neuromodulation can provide diagnostic, modulatory, and therapeutic applications. While extensive work has been conducted in the brain, modulation of the spinal cord remains relatively unexplored. The inherently delicate and mobile spinal cord tissue imposes constraints that make the precise implantation of neural probes challenging. Despite recent advances in neuromodulation devices, particularly flexible bioelectronics, opportunities to expand their use in the spinal cord have been limited by the surgical complexities of device implantation. Here, we provide a series of surgical protocols tailored specifically for the implantation of a custom-made optoelectronic device that interfaces with the spinal cord in rodents. The steps to place and anchor an optical shank on a specific segment of the spinal cord via two different surgical implantation methods are detailed here. These methods are optimized for a diverse range of devices and applications, which may or may not require direct contact with the spinal cord for optical stimulation. To elucidate the methodology, the vertebral anatomy is referenced first to identify prominent landmarks before making a skin incision. The surgical steps to secure an optical shank over the cervical spine in rodents are demonstrated. Procedures are then outlined for securing the optoelectronic device connected to the optical shank in a subcutaneous space away from the spinal cord, minimizing unnecessary direct contact. Behavioral studies comparing animals receiving the implants to those undergoing sham surgeries indicate that the optical shanks did not adversely affect hindlimb or forelimb function seven days post-implantation. The present work broadens the neuromodulation toolkit for use in future studies aimed at investigating various spinal cord interventions.

Author Spotlight: Innovative Methodology for Implanting and Securing Neural Probes in the Rodent Spinal Cord

This protocol details a surgical procedure for performing spinal cord surgery and for implanting and securing an optical shank over the spinal cord in rodents.

Neuromodulation can provide diagnostic, modulatory, and therapeutic applications. While extensive work has been conducted in the brain, modulation of the ...

Surgical Technique for Lumbar Spinal Catheter Insertion in Pigs Enabling Continuous Access to the Thecal Sac in a Terminal Setup

Pigs are increasingly used as a large animal model for pharmacologic CNS research due to the anatomical and physiological similarities between the porcine and human central nervous systems (CNS). However, accessing the cerebrospinal fluid (CSF) in larger pig breeds by conventional lumbar puncture techniques can be challenging due to an oblique orientation of the spinal spinous processes and a limited interlaminar space. Accordingly, an open surgical procedure for inserting a lumbar spinal catheter for continuous CSF sampling at the L4/L5 level in pigs is thoroughly described in this work. After positioning the pig and identifying the anatomical landmarks, a dorsal midline surgical incision is made to expose the spinous processes. By advancing the introducer needle, the spinal catheter is inserted inside the thecal sac of the spinal canal while leaving the bone structures of the spine intact. This method allows continuous infusion into or sampling from the porcine thecal sac with minimal bleeding or CSF leakage. The procedure is simple, time-efficient, and reproducible across different experimental setups, offering significant potential for various pre-clinical studies, including pharmacokinetic research, surgical training, and spinal cord injury models.

We present a technique for inserting a lumbar spinal catheter at the L4-L5 level in a 3-month-old Danish Landrace pig as part of a terminal research protocol, enabling continuous infusion or CSF sampling from the thecal sac.

Pigs are increasingly used as a large animal model for pharmacologic CNS research due to the anatomical and physiological similarities between the porcine ...