This work was supported by NIH 1R01HL132325 and R21 DK119740-01 (VCC) and AHA Cardio-oncology SFRN CAT-HD Center grant 857078 (VCC and SL).

For this study, eight female C57BL/6J mice, 8-12 weeks in age, and an average weight of 20 g were used. The mice were housed under standard conditions and were fed with chow and water ad libitum. This study was performed with the approval of the Institutional Animal Care and Use Committee (IACUC), Boston University Medical Center (AN-1549). The procedures described here were performed under sterile conditions.
1. Anesthetize the mouse in an Isoflurane chamber, and inject the analgesic subcutaneously
<ol>
	<li>Hold the animal from the base of the tail. Keep the animal on the dorsum surface of the non-dominant hand.</li>
	<li>Transfer the animal to the continuous anesthetic induction chamber filled with 3%-4% isoflurane. Confirm adequate general anesthesia by the absence of toe-pinch reflex in the right and left hind limbs. Keep the maintenance of the general anesthesia with Isoflurane 1%-3%.</li>
	<li>Apply ophthalmic ointment on both eyes.</li>
	<li>Administer a subcutaneous injection of Buprenorphine.
	<ol>
		<li>Dissolve the stock of Buprenorphine at a concentration of 0.3 mg/mL in sodium chloride (NaCl) 0.9% to achieve the final concentration of 0.03 mg/mL.</li>
		<li>Inject a dosage of 0.05-0.1 mg/kg of 0.03 mg/mL Buprenorphine, together with 500 &#181;L of sterile NaCl 0.9%, 20 min before surgery in a 20 g mouse (2 &#181;g or 66 &#181;L of 0.03 mg/mL Buprenorphine per mouse).</li>
	</ol>
	</li>
</ol>
2. Skin preparation
<ol>
	<li>Place the fully anesthetized mouse in a left lateral position, exposing its right flank to the heating blanket. Shave the right side of the abdomen, just close to the midline to the paraspinal area, and down to the animal&#39;s tail.</li>
	<li>Disinfect the shaved area three times using a cotton swab with the alternate application of the antiseptic solution or scrub and either 70% alcohol or sterile saline in a circular motion, starting at the surgical incision site and moving outward. Discard the cotton swab after each use. Be careful not to excessively wet non-surgical areas of the animal with alcohol or antiseptic as this can worsen hypothermia. 
	NOTE: It is important to properly dilute antiseptic solutions and not leave surgical scrubs on the skin during surgery, as they can be irritating and need to be rinsed off. Frequently check the temperature of the heating blanket during the procedure to ensure that the temperature does not fall.</li>
</ol>
3. Measure the catheter length and mark the insertion point within the abdomen and the tube tract over the prepared skin
<ol>
	<li>Assign the access port pocket 1 cm above the animal&#39;s tail. Hold the installation segment with the non-dominant index and thumb finger over the assigned area near the tail.</li>
	<li>Place the catheter above the skin and estimate the place for the catheter&#39;s tube insertion within the abdominal cavity. Mark the assigned place for tube insertion, respecting the minimal bending of the tube near the anterior midline. 
	NOTE: All the procedures must be performed with sterile gloves, and the catheter should be kept sterile during the measurement. Surgical tools must be autoclaved at 121 &#176;C before use. Refer to Supplemental Figure S1 for the instruments required for the procedure.</li>
</ol>
4. Customize the peritoneal catheter reservoir section
<ol>
	<li>Punch a side hole over the frame of the reservoir section with the mouse ear tagger (Figure 1 and Figure 2). It should be noted that the ear punch is a surgical tool, and it needs to be sterile.</li>
</ol>
5. Place the instillation port
<ol>
	<li>Make a horizontal 1 cm wide skin incision 1 cm above the tail. Bluntly dissect the subcutaneous plane from the underlying muscular layer to make a pouch for the catheter placement to ensure the instillation port resides in the ideal port pocket freely.</li>
	<li>Keep the iris scissors&#39; tip toward the midline to make an oblique tunnel for the tube placement (Figure 3A).</li>
	<li>Pass the 3.0 suture from the customized side hole. Fix the access port to the muscular bed by tightening the passed suture, keeping the tubing course cephalad.</li>
</ol>
6. Make the catheter tip insertion site incision
<ol>
	<li>Make a 1 cm incision over the formerly marked area near the midline. Confirm the well-developed tract by passing scissors through the tract.</li>
	<li>Pick the catheter tip gently with forceps to place the catheter in a retrograde course. 
	NOTE: Avoid pinching the side holes of the tube.</li>
	<li>Pass the catheter tube through the prepared tract (Figure 3B). Make a 1 cm incision over the muscular layer close to the right midline.</li>
</ol>
7. Confirm the functioning of the catheter
<ol>
	<li>Before closing all the incisions, make sure the placed catheter is functional. Check the function with a 1 mL syringe attached to the specific Huber needle for the port.</li>
	<li>Inject 200 &#181;L of normal saline into the instillation port. Look for a smooth flow with a zero tolerance for resistance.</li>
	<li>Flush the port and catheter with 10% heparin to maintain patency.</li>
</ol>
8. Close the skin incisions
<ol>
	<li>Close the skin incisions around the port reservoir (Figure 3C) with 3-0 absorbable sutures.</li>
</ol>
9. Fix the catheter tip inside the abdominal cavity
<ol>
	<li>Place a loose purse-string suture with 4-0 round absorbable suture around the incised abdominal wall muscle. Pass the proximal felt of the catheter inside the incision.</li>
	<li>Tighten the prepared purse-string suture around the tube while keeping the second felt outside the purse string, over the muscular layer (Figure 3D), and close the skin with 3-0 absorbable sutures (Figure 2).</li>
</ol>
10. Monitor the animals postoperatively and daily, administer postoperative analgesia and fluids, and maintain daily postoperative records for a minimum of 7 days and until complete recovery
<ol>
	<li>Keep the catheter functional with a daily injection of 200 &#181;L of normal saline through the catheter.</li>
</ol>
11. Fluid injections
<ol>
	<li>Confirm the uneventful postprocedural process by carefully inspecting the skin incision.</li>
	<li>Prepare LPS 2 mg/kg body weight for intraperitoneal injections (i.p.) by diluting 40 &#181;g of the LPS with sterile phosphate-buffered saline (PBS) to the working concentration of 0.2 &#181;g/&#181;L (in essence, 10 &#181;L for 2 &#181;g/g body weight and 200 &#181;L of LPS for 20 g mice).</li>
	<li>Start the injections in the second week following the catheter implantation.
	<ol>
		<li>Hold the animal gently with the non-dominant hand and restrain the instillation port while moving the index and thumb fingers in the cephalad direction.</li>
		<li>Disinfect the skin overlying the reservoir with 70% isopropyl alcohol. Use the syringe attached to the Huber needle to inject the LPS.
		<ol>
			<li>After entering the port with the Huber needle, inject 100 &#181;L of normal saline into the port to confirm the patent course.</li>
			<li>Inject the prepared 200 &#181;L of LPS, followed by the 100 &#181;L of normal saline for tube irrigation, and make sure there is no resistance.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
12. Anesthetize the mice before harvesting the peritoneum and collect the peritoneal fluid
<ol>
	<li>Following 7 days of LPS injections and 2 weeks of catheter implantation, plan for the peritoneal biopsy.</li>
	<li>Plan for general anesthesia.
	<ol>
		<li>Anesthetize the mouse in an isoflurane chamber and inject the analgesic subcutaneously.</li>
		<li>Hold the animal from the base of the tail, and keep the animal on the dorsum surface of the hand.</li>
		<li>Transfer the animal to the continuous anesthetic induction chamber filled with 3%-4% isoflurane. Confirm adequate general anesthesia by the absence of toe-pinch reflex in the right and left hind limbs. Keep the maintenance of the general anesthesia with isoflurane 1%-3%.</li>
	</ol>
	</li>
</ol>
13. Peritoneal biopsy
<ol>
	<li>Place the animal on the heated blanket in the supine position. Make a midline skin incision from the sub-xiphoid to the bladder.</li>
	<li>Perfuse the subfascial plane with cold PBS (Figure 3E).</li>
	<li>Make sure the plane is completely dissected without disturbing the integrity of the peritoneum. Start dissecting the peritoneum from the lateral peritoneal reflection at the left lower quadrant, starting from the hilum to the left flank, and bladder in the lower aspect to keep samples consistent between animals (Figure 3F).</li>
	<li>Following the peritoneal harvest, euthanize the animal by cervical dislocation.</li>
</ol>

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The authors have no conflicts of interest to disclose.

Three murine models of PD are described. This includes a blind puncture of the peritoneal surface, an open-permanent system, and a closed system10. The blind puncture of the peritoneal surface involves direct peritoneal access similar to intraperitoneal injections but does not allow drainage of dialysate. Being a blinded procedure, this method can injure the abdominal visceral organs. The open-permanent system model keeps the dialysis catheter and instillation port outside the body. However, this ...

An erratum was issued for:&#160;<a href="https://www.jove.com/t/63689">A Retrograde Implantation Approach for Peritoneal Dialysis Catheter Placement in Mice</a>. The Authors section was updated from:
Saran Lotfollahzadeh1 
Mengwei Zhang1 
Marc Arthur Napoleon1 
Wenqing Yin1 
Josephine Orrick1 
Nagla Elzind1 
Austin Morrissey1 
Isaac E. Sellinger1 
Lauren D. Stern1 
Mostafa Belghasem2 
Jean M. Francis1 
Vipul C. Chitalia1,3,4 
1Renal Section, Department of Medicine, Boston University School of Medicine 
2Department of Biomedical Science, Kaiser Permanente Bernard J. Tyson School of Medicine 
3Veterans Affairs Boston Healthcare System 
4Institute of Medical Engineering and Sciences, Massachusetts Institute of Technology
to:
Saran Lotfollahzadeh1 
Mengwei Zhang1 
Marc Arthur Napoleon1 
Wenqing Yin1 
Josephine Orrick1 
Nagla Elzind1 
Austin Morrissey1 
Isaac E. Sellinger1 
Lauren D. Stern1 
Mostafa Belghasem2 
Jean M. Francis1 
Vipul C. Chitalia1,3,4 
1Renal Section, Department of Medicine, Boston University Aram V. Chobanian &#38; Edward Avedisian School of Medicine 
2Department of Biomedical Science, Kaiser Permanente Bernard J. Tyson School of Medicine 
3Veterans Affairs Boston Healthcare System 
4Institute of Medical Engineering and Sciences, Massachusetts Institute of Technology

An erratum was issued for:&#160;<a href="https://www.jove.com/t/63689">A Retrograde Implantation Approach for Peritoneal Dialysis Catheter Placement in Mice</a>. The Authors section was updated.

Erratum: A Retrograde Implantation Approach for Peritoneal Dialysis Catheter Placement in Mice

End-stage renal disease burden 
Chronic kidney disease (CKD) is a worldwide health problem1. Current estimates suggest that more than 850 million people worldwide have kidney disease. The prevalence of kidney disease almost doubles the number of people with diabetes (422 million) and is more than 20 times the prevalence of cancer (42 million) or HIV/AIDS (36.7 million) patients worldwide2. Approximately one in seven Americans have CKD, and two in 1,000 Americans have ESKD requiring a kidney transplant or dialysis support3. Considering the escalating burden of ESKD worldwide, optimizing dialysis technology is crucial3.
Peritoneal dialysis 
PD is a significantly underutilized modality for the treatment of ESKD in the United States. According to the United States Renal Data System (USRDS), the percentage of prevalent PD patients was only 11% in 20204,5. PD confers several advantages over in-center hemodialysis (HD), including a better quality of life, fewer clinic visits, and a decrease in Medicare expenditures6,7. Additionally, PD is a home-based therapy and is associated with a much lower risk of severe infections such as bacteremia and endocarditis that are often related to hemodialysis catheters. Furthermore, PD can be initiated rapidly with an urgent start protocol, decreasing the need for dialysis initiation with indwelling vascular catheters8. PD is considered the preferred method of dialysis in the pediatric ESKD population9.
Peritoneal impairment induced by peritoneal dialysis 
PD entails introducing PD fluid (dialysate) into the peritoneum, which results in inflammation and remodeling of the peritoneal membrane over time. Peritoneal inflammation triggers fibrosis, culminating in the potential loss of ultrafiltration capabilities of the membrane over time. Preservation of the peritoneal membrane is a significant challenge in PD, and further research is critically important to ensure that best clinical practices are available to practitioners. There are well-established murine models to help further the understanding of pathophysiological mechanisms of peritoneal infection and inflammation, solute, water transport kinetics, and membrane failure; still, technical issues with the catheter often limit these models10.
Analyzing the peritoneal membrane changes 
In ESKD patients, dialysate is traditionally introduced in the peritoneal cavity through a Tenkhoff catheter with a deep and superficial cuff. The patients can potentially experience catheter-related complications, including catheter migration, infusion pain, and poor drainage of the dialysate11,12,13. Two major types of peritoneal catheters have been introduced for humans, coiled or straight, to minimize these complications12. Several modifications, including an extra cuff to the conventional two-cuffed catheters, have been added to the original catheters to prolong PD catheter survival11. The insertion technique varies according to several factors by preventing catheter migration to be added after survival, including the availability of the resources and the level of expertise14.
In contrast, the murine models of peritoneal dialysis have fundamental differences in techniques and purpose compared to human peritoneal catheters. For example, peritoneal catheters in murine models are used primarily to study membrane alterations and are less intended for bidirectional drainage functions. The current technique suffers from potential port dislodgement and catheter migration due to the handling of the animals. In the conventional murine models, the access ports were not fixed to the skin. This aspect created an unstable access port, which in awake animals might get dislodged, resulting in catheter migration. Given the importance of murine models in peritoneal membrane research, it is imperative to create effective surgical techniques to generate reliable models. Therefore,&#160;we set out to optimize the conventional model of PD catheter placement. It is important to note that the catheter itself causes histopathologic alterations in the peritoneal membrane, and, thus, any conclusions regarding the effect of PD solutions in animal studies must be interpreted in the context of the PD catheter as a foreign body15,16,17.
Peritoneal membrane histopathology 
PD failure is mainly related to fibrosis and excess angiogenesis resulting in the loss of an osmolar concentration gradient. In addition, the peritoneal membrane filtration capacity might be affected by peritonitis. In addition, infectious peritonitis is a well-established cause for change in the dialysis modality from peritoneal dialysis to hemodialysis.18.

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	<tbody>
		<tr>
			<td>10% heparin&#160;</td>
			<td>Canada Inc., Boucherville, QC, Canada)</td>
			<td>Pharmaceutical product</td>
			<td></td>
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			<td>&#160;&#160;&#160;&#160; Buprenorphine 0.3 mg/mL</td>
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			<td>C57BL/6J mice</td>
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			<td>Dumont Vessel Cannulation Forceps</td>
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			<td>Fine Scissors - Large Loops</td>
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			<td>Fisherbrand&#160;Animal Ear-Punch</td>
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			<td>13-812-201</td>
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			<td>Hill Hemostat</td>
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			<td>13111-12</td>
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			<td>Huber point needle</td>
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			<td>Needle for injections</td>
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		<tr>
			<td>&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; Isoflurane, USP</td>
			<td>&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; Covetrus</td>
			<td>&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; NDC 11695-6777-2</td>
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			<td>&#160;&#160;&#160;&#160;&#160;&#160; Lipopolysaccharide from E.coli</td>
			<td>&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; SIGMA</td>
			<td>&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; L4391</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope</td>
			<td>Nikon Eclipse Inverted Microscope</td>
			<td>TE2000</td>
			<td></td>
		</tr>
		<tr>
			<td>Minute Mouse Port 4French with retention beads and cross holes</td>
			<td>&#160;&#160;&#160; Access&#160; technologies&#160;</td>
			<td>&#160;&#160;&#160;&#160;&#160;&#160; MMP-4S-061108A</td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;Posi-Grip Huber point needles 25 G x 1/2&#180;&#180;</td>
			<td>&#160;&#160; Access&#160; technologies</td>
			<td>&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; PG25-500</td>
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			<td>&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; Scissors</td>
			<td>&#160;&#160;&#160;&#160; Fine Science Tools</td>
			<td>&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160;&#160; 14079-10</td>
			<td></td>
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		<tr>
			<td>Vicryl Suture</td>
			<td>AD-Surgical</td>
			<td>#L-G330R24</td>
			<td></td>
		</tr>
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a retrograde implantation approach for peritoneal dialysis catheter placement in mice

Murine models are employed to probe various aspects of peritoneal dialysis (PD), such as peritoneal inflammation and fibrosis. These events drive peritoneal membrane failure in humans, which remains an area of intense investigation due to its profound clinical implications in managing patients with end-stage kidney disease (ESKD). Despite the clinical importance of PD and its related complications, current experimental murine models suffer from key technical challenges that compromise the models&#39; performance. These include PD catheter migration and kinking and usually warrant earlier catheter removal. These limitations also drive the need for a greater number of animals to complete a study. Addressing these drawbacks, this study introduces technical improvements and surgical nuances to prevent commonly observed PD catheter complications in a murine model. Moreover, this modified model is validated by inducing peritoneal inflammation and fibrosis using lipopolysaccharide injections. In essence, this paper describes an improved method to create an experimental model of PD.

All the implanted catheters were functional till the end of the study, and catheter dislodgement or kinking did not complicate any of the implanted catheters. The current, modified technique was further validated with a peritonitis-induced model using LPS. The control mice received 200 &#181;L of daily normal saline injections, while the experimental mice were injected with 200 &#181;L of LPS, as discussed in protocol step 11, for a total of 7 days following catheter implantation.
The peritone...

Watch this Scientific Journal Video about A Retrograde Implantation Approach for Peritoneal Dialysis Catheter Placement in Mice at JoVE.com

A Retrograde Implantation Approach for Peritoneal Dialysis Catheter Placement in Mice

This article describes modifications of a procedure to implant a peritoneal dialysis catheter in a murine model to avoid major technical issues observed with the conventional techniques.

Murine models are employed to probe various aspects of peritoneal dialysis (PD), such as peritoneal inflammation and fibrosis. These events drive ...

a-retrograde-implantation-approach-for-peritoneal-dialysis-catheter

Research

JoVE Journal

Medicine

2.4K Views.  Boston University Aram V. Chobanian & Edward Avedisian School of Medicine. This article describes modifications of a procedure to implant a peritoneal dialysis catheter in a murine model to avoid major technical issues observed with the conventional techniques.

Video: A Retrograde Implantation Approach for Peritoneal Dialysis Catheter Placement in Mice

Implantation of a Carotid Cuff for Triggering Shear-stress Induced Atherosclerosis in Mice

It is widely accepted that alterations in vascular shear stress trigger the expression of inflammatory genes in endothelial cells and thereby induce atherosclerosis (reviewed in 1 and 2). The role of shear stress has been extensively studied in vitro investigating the influence of flow dynamics on cultured endothelial cells 1,3,4 and in vivo in larger animals and humans 1,5,6,7,8. However, highly reproducible small animal models allowing systematic investigation of the influence of shear stress on plaque development are rare. Recently, Nam et al. 9 introduced a mouse model in which the ligation of branches of the carotid artery creates a region of low and oscillatory flow. Although this model causes endothelial dysfunction and rapid formation of atherosclerotic lesions in hyperlipidemic mice, it cannot be excluded that the observed inflammatory response is, at least in part, a consequence of endothelial and/or vessel damage due to ligation.
In order to avoid such limitations, a shear stress modifying cuff has been developed based upon calculated fluid dynamics, whose cone shaped inner lumen was selected to create defined regions of low, high and oscillatory shear stress within the common carotid artery 10. By applying this model in Apolipoprotein E (ApoE) knockout mice fed a high cholesterol western type diet, vascular lesions develop upstream and downstream from the cuff. Their phenotype is correlated with the regional flow dynamics 11 as confirmed by in vivo Magnetic Resonance Imaging (MRI) 12: Low and laminar shear stress upstream of the cuff causes the formation of extensive plaques of a more vulnerable phenotype, whereas oscillatory shear stress downstream of the cuff induces stable atherosclerotic lesions 11. In those regions of high shear stress and high laminar flow within the cuff, typically no atherosclerotic plaques are observed.
 
In conclusion, the shear stress-modifying cuff procedure is a reliable surgical approach to produce phenotypically different atherosclerotic lesions in ApoE-deficient mice.

The constricting cuff presented in this article is designed to induce atherosclerosis in the murine common carotid artery. Due to the conical shape of its inner lumen the implanted cuff generates well-defined regions of low, high and oscillatory shear stress triggering the development of atherosclerotic lesions of different inflammatory phenotypes.

It is widely accepted that alterations in vascular shear stress trigger the expression of inflammatory genes in endothelial cells and thereby induce ...

Carotid Artery Infusions for Pharmacokinetic and Pharmacodynamic Analysis of Taxanes in Mice

When proposing the use of a drug, drug combination, or drug delivery into a novel system, one must assess the pharmacokinetics of the drug in the study model. As the use of mouse models are often a vital step in preclinical drug discovery and drug development1-8, it is necessary to design a system to introduce drugs into mice in a uniform, reproducible manner. Ideally, the system should permit the collection of blood samples at regular intervals over a set time course. The ability to measure drug concentrations by mass-spectrometry, has allowed investigators to follow the changes in plasma drug levels over time in individual mice1, 9, 10. In this study, paclitaxel was introduced into transgenic mice as a continuous arterial infusion over three hours, while blood samples were simultaneously taken by retro-orbital bleeds at set time points. Carotid artery infusions are a potential alternative to jugular vein infusions, when factors such as mammary tumors or other obstructions make jugular infusions impractical. Using this technique, paclitaxel concentrations in plasma and tissue achieved similar levels as compared to jugular infusion. In this tutorial, we will demonstrate how to successfully catheterize the carotid artery by preparing an optimized catheter for the individual mouse model, then show how to insert and secure the catheter into the mouse carotid artery, thread the end of the catheter out through the back of the mouse&#8217;s neck, and hook the mouse to a pump to deliver a controlled rate of drug influx. Multiple low volume retro-orbital bleeds allow for analysis of plasma drug concentrations over time.

This method was developed with the goal of delivering a steady drug solution via the carotid artery, to assess the pharmacokinetics of novel drugs in mouse models.

When proposing the use of a drug, drug combination, or drug delivery into a novel system, one must assess the pharmacokinetics of the drug in the study ...

The Supraclavicular Fossa Ultrasound View for Central Venous Catheter Placement and Catheter Change Over Guidewire

The supraclavicular fossa ultrasound view can be useful for central venous catheter (CVC) placement. Venipuncture of the internal jugular veins (IJV) or subclavian veins is performed with a micro-convex ultrasound probe, using a neonatal abdominal preset with a probe frequency of 10 Mhz at a depth of 10-12 cm. Following insertion of the guidewire into the vein, the probe is shifted to the right supraclavicular fossa to obtain a view of the superior vena cava (SVC), right pulmonary artery and ascending aorta. Under real-time ultrasound view, the guidewire and its J-tip is visualized and pushed forward to the lower SVC. Insertion depth is read from guidewire marks using central venous catheter. CVC is then inserted following skin and venous dilation. The supraclavicular fossa view is most suitable for right IJV CVC insertion. If other insertion sites are chosen the right supraclavicular fossa should be within the sterile field. Scanning of the IJVs, brachiocephalic veins and SVC can reveal significant thrombosis before venipuncture. Misplaced CVCs can be corrected with a change over guidewire technique under real-time ultrasound guidance. In conjunction with a diagnostic lung ultrasound scan, this technique has a potential to replace chest radiograph for confirmation of CVC tip position and exclusion of pneumothorax. Moreover, this view is of advantage in patients with a non-p-wave cardiac rhythm were an intra-cardiac electrocardiography (ECG) is not feasible for CVC tip position confirmation. Limitations of the method are lack of availability of a micro-convex probe and the need for training.

The overall goal is to enable clinicians to insert a central venous catheter under real-time ultrasound guidance via the right supraclavicular fossa view of the lower part of the superior vena cava. This view is useful for different catheter insertion sites, thrombosis detection before insertion and ultrasound guided correction of misplaced catheters.

The supraclavicular fossa ultrasound view can be useful for central venous catheter (CVC) placement. Venipuncture of the internal jugular veins (IJV) or ...

Non-fluoroscopic Catheter Tracking for Fluoroscopy Reduction in Interventional Electrophysiology

A technological platform (MediGuide) has been recently introduced for non-fluoroscopic catheter tracking. In several studies, we have demonstrated that the application of this non-fluoroscopic catheter visualization system (NFCV) reduces fluoroscopy time and dose by 90-95% in a variety of electrophysiology (EP) procedures. This can be of relevance not only to the patients, but also to the nurses and physicians working in the EP lab. Furthermore, in a subset of indications such as supraventricular tachycardias, NFCV enables a fully non-fluoroscopic procedure and allows the lab staff to work without wearing lead aprons. With this protocol, we demonstrate that even complex procedures such as ablations of atrial fibrillation, that are typically associated with fluoroscopy times of &#62;30 min in conventional settings, can safely be performed with a reduction of &#62;90% in fluoroscopy exposure by the additional use of NFCV.

Radiation exposure is an underestimated risk in complex ablation procedures. Here, we describe a protocol to significantly decrease fluoroscopy time and dosage for both the patient and the lab staff by using a novel non-fluoroscopic catheter visualization system.

A technological platform (MediGuide) has been recently introduced for non-fluoroscopic catheter tracking. In several studies, we have demonstrated that ...

Quantitation of Intra-peritoneal Ovarian Cancer Metastasis

Epithelial ovarian cancer (EOC) is the leading cause of death from gynecologic malignancy in the United States. Mortality is due to diagnosis of 75% of women with late stage disease, when metastasis is already present. EOC is characterized by diffuse and widely disseminated intra-peritoneal metastasis. Cells shed from the primary tumor anchor in the mesothelium that lines the peritoneal cavity as well as in the omentum, resulting in multi-focal metastasis, often in the presence of peritoneal ascites. Efforts in our laboratory are directed at a more detailed understanding of factors that regulate EOC metastatic success. However, quantifying metastatic tumor burden represents a significant technical challenge due to the large number, small size and broad distribution of lesions throughout the peritoneum. Herein we describe a method for analysis of EOC metastasis using cells labeled with red fluorescent protein (RFP) coupled with in vivo multispectral imaging. Following intra-peritoneal injection of RFP-labelled tumor cells, mice are imaged weekly until time of sacrifice. At this time, the peritoneal cavity is surgically exposed and organs are imaged in situ. Dissected organs are then placed on a labeled transparent template and imaged ex vivo. Removal of tissue auto-fluorescence during image processing using multispectral unmixing enables accurate quantitation of relative tumor burden. This method has utility in a variety of applications including therapeutic studies to evaluate compounds that may inhibit metastasis and thereby improve overall survival.

Ovarian cancer metastasis is characterized by numerous diffuse intra-peritoneal lesions, such that accurate visual quantitation of tumor burden is challenging. Herein we describe a method for in situ and ex vivo quantitation of metastatic tumor burden using red fluorescent protein (RFP)-labeled tumor cells and optical imaging.

Epithelial ovarian cancer (EOC) is the leading cause of death from gynecologic malignancy in the United States.  Mortality is due to diagnosis of 75% of ...

Surgical Approach for Middle Cerebral Artery Occlusion and Reperfusion Induced Stroke in Mice

Stroke is a leading cause of death worldwide and continues to be one of the major causes of long-term adult disabilities. About 87% of strokes are ischemic in origin and occur in the territory of the middle cerebral artery (MCA). Currently the only Food and Drug Administration (FDA) approved drug for the treatment of this devastating disease is tissue plasminogen activator (tPA). However, tPA has a small therapeutic window for administration (3 - 6 hr), and is only effective in 4% of the patients who actually receive it. Current research focuses on understanding the pathophysiology of stroke in order to find potential therapeutic targets. Thus, reliable models are crucial, and the MCA occlusion (MCAo) model (also termed the intraluminal filament or suture model) is deemed to be the most clinically relevant surgical model of ischemic stroke, and is fairly non-invasive and easily reproducible. Typically the MCAo model is used with rodents, especially with mice due to all the genetic variations available for this species. Here we describe (and present in the video) how to successfully perform the MCAo model (with reperfusion) in mice to generate reliable and reproducible data.

In order to understand the pathophysiology of stroke, it is important to use reliable models. This paper will describe one of the most frequently used stroke models in mice, termed the middle cerebral artery occlusion (MCAo) model (also termed the intraluminal filament or suture model) with reperfusion.

Stroke is a leading cause of death worldwide and continues to be one of the major causes of long-term adult disabilities. About 87% of strokes are ...

Implantation of Combined Telemetric ECG and Blood Pressure Transmitters to Determine Spontaneous Baroreflex Sensitivity in Conscious Mice

Blood pressure (BP) and heart rate (HR) are both controlled by the autonomic nervous system (ANS) and are closely intertwined due to reflex mechanisms. The baroreflex is a key homeostatic mechanism to counteract acute, short-term changes in arterial BP and to maintain BP in a relatively narrow physiological range. BP is sensed by baroreceptors located in the aortic arch and carotid sinus. When BP changes, signals are transmitted to the central nervous system and are then communicated to the parasympathetic and sympathetic branches of the autonomic nervous system to adjust HR. A rise in BP causes a reflex decrease in HR, a drop in BP causes a reflex increase in HR.
Baroreflex sensitivity (BRS) is the quantitative relationship between changes in arterial BP and corresponding changes in HR. Cardiovascular diseases are often associated with impaired baroreflex function. In various studies reduced BRS has been reported in e.g., heart failure, myocardial infarction, or coronary artery disease. 
Determination of BRS requires information from both BP and HR, which can be recorded simultaneously using telemetric devices. The surgical procedure is described beginning with the insertion of the pressure sensor into the left carotid artery and positioning of its tip in the aortic arch to monitor arterial pressure followed by the subcutaneous placement of the transmitter and ECG electrodes. We also describe postoperative intensive care and analgesic management. After a two-week period of post-surgery recovery long-term ECG and BP recordings are performed in conscious and unrestrained mice. Finally, we include examples of high-quality recordings and the analysis of spontaneous baroreceptor sensitivity using the sequence method.

The baroreflex is a heart-rate regulation mechanism by the autonomic nervous system in response to blood-pressure changes. We describe a surgical technique to implant telemetry transmitters for continuous and simultaneous measurement of electrocardiogram and blood pressure in mice. This can determine spontaneous baroreflex sensitivity, an important prognostic marker for cardiovascular disease.

Blood pressure (BP) and heart rate (HR) are both controlled by the autonomic nervous system (ANS) and are closely intertwined due to reflex mechanisms. ...

Dynamic Navigation for Dental Implant Placement

In modern implantology, the application of surgical navigation systems is becoming increasingly important. In addition to static surgical navigation methods, a guide-independent dynamic navigation implant placement procedure is becoming more widespread. The procedure is based on computer-guided dental implant placement utilizing optical control. This work aims to demonstrate the technical steps of a new dynamic computer-aided implant surgery (DCAIS)&#160;system (design, calibration, surgery) and check the accuracy of the results. Based on cone-beam computed tomography (CBCT) scans, the exact positions of implants are determined with dedicated software. The first step of the operation is the calibration of the navigation system, which can be performed in two ways: 1) based on CBCT images taken with a marker or 2) based on CBCT images without markers. Implants are inserted with the aid of real-time navigation according to the preoperative plans. The accuracy of the interventions can be evaluated based on postoperative CBCT images. The preoperative images containing the planned positions of the implants and postoperative CBCT images were compared based on the angulation (degree), platform, and apical deviation (mm) of the implants. To evaluate the data, we calculated the standard deviation (SD), mean, and standard error of the mean (SEM) of deviations within planned and performed implant positions. Differences between the two calibration methods were compared based on this data. Based on the interventions performed so far, the use of DCAIS allows for high-precision implant placement. A calibration system that does not require labeled CBCT recording allows for surgical intervention with similar accuracy as a system that uses labeling. The accuracy of the intervention can be improved by training.

Dynamic computer-aided implant surgery (DCAIS) is a controlled implant surgical placement method performed without a surgical template using optical control. The real-time intraoperative control of movement and position of the surgical device simplifies the procedure and gives more freedom to the surgeon, providing similar precision as static navigation methods.

In modern implantology, the application of surgical navigation systems is becoming increasingly important. In addition to static surgical navigation ...

A Mice Model of Chlorhexidine Gluconate-Induced Peritoneal Damage

Peritoneal fibrosis is an important complication of peritoneal dialysis (PD). To investigate and address this problem, an appropriate animal model of PD is required. The present protocol establishes a chlorhexidine gluconate (CG) induced peritoneal fibrosis model that mimics the condition of a patient with PD. Peritoneal fibrosis was induced by intraperitoneal injection of 0.1% of CG in 15% ethanol for 3 weeks (administered every other day), for a total of nine times in male C57BL/6 mice. Peritoneal functional tests were then performed on day 22. After the mice were sacrificed, the parietal peritoneum of the abdominal wall and the visceral peritoneum of the liver were harvested. They were thicker and more fibrotic when analyzed microscopically after Masson's trichrome staining. The ultrafiltration rate decreased, and glucose mass transport indicated a CG-induced increase in peritoneal permeability. The PD model thus established may have applications in improving PD technology, dialysis efficacy, and prolonging patient survival.

The present protocol establishes a peritoneal dialysis (PD) mouse model of chlorhexidine gluconate (CG)-induced peritoneal fibrosis. The current model is simple and easy to use compared to other PD animal models.

Peritoneal fibrosis is an important complication of peritoneal dialysis (PD). To investigate and address this problem, an appropriate animal model of PD ...

Hickman Catheter for Long-Term Vascular Access in Swine

Hickman Catheter Use for Long-Term Vascular Access in a Preclinical Swine Model

Central venous catheters (CVCs) are invaluable devices in large animal research as they facilitate a wide range of medical applications, including blood monitoring and reliable intravenous fluid and drug administration. Specifically, the tunneled multi-lumen Hickman catheter (HC) is commonly used in swine models due to its lower extrication and complication rates. Despite fewer complications relative to other CVCs, HC-related morbidity presents a significant challenge, as it can significantly delay or otherwise negatively impact ongoing studies. The proper insertion and maintenance of HCs is paramount in preventing these complications, but there is no consensus on best practices. The purpose of this protocol is to comprehensively describe an approach for the insertion and maintenance of a tunneled HC in swine that mitigates HC-related complications and morbidity. The use of these techniques in &gt;100 swine has resulted in complication-free patent lines up to 8 months and no catheter-related mortality or infection of the ventral surgical site. This protocol offers a method to optimize the lifespan of the HC and guidance for approaching issues during use.

Author Spotlight: Hickman Catheter Use for Long-Term Vascular Access in a Preclinical Swine Model  

A reliable and reproducible approach for the insertion and maintenance of a tunneled Hickman catheter for long-term vascular access in swine is described. Placement of a central venous catheter allows for convenient daily sampling of whole blood from awake animals and intravenous administration of medication and fluids.

Central venous catheters (CVCs) are invaluable devices in large animal research as they facilitate a wide range of medical applications, including blood ...