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 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 >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.
The indispensable role of central venous catheters (CVCs) in patient care is owed to their convenience, favorable safety profile, and versatility1. Functions of a CVC include reliable access for total parenteral nutrition, hematopoietic stem cell transplantation, plasmapheresis/apheresis, and efficient fluid, blood, or co-drug administration2. In veterinary medicine, CVCs also minimize animal discomfort via the rapid dilution of irritant drugs and blood sampling without repeated venipuncture3. Despite their broad applications, the use of CVCs in large animal research still presents several considerable challenges4.
Percutaneous CVC placement via a guidewire or introducer catheter can be difficult for non-veterinary researchers, particularly in animals with deep venous structures5. An improper CVC installation technique may result in inadvertent placement into nearby structures, necessitating ultrasound-guided placement or a postprocedure radiography of the positioning6. However, compared to human operating rooms, ultrasounds are not readily available in many large animal research laboratories. Further, long-term use of indwelling catheters can result in line kinking, puncture, infection, or extrication by animals, with the possible disruption of timely treatment, clinical monitoring, and research outcomes4,7. Replacement of the CVC requires additional resources, including material procurement, surgical scheduling, fasting time, and radiographic access. CVC-related complications can therefore create significant technical and financial barriers or a disruption to productive translational research, particularly in swine. Contamination by food or feces, scratching against cage walls, and kicking sites of irritation may compromise a CVC, and the risk of CVC-related complications is amplified by long-term use. Thus, safe and uncomplicated maintenance of a CVC in swine requires careful consideration of CVC choice, placement, securement, protection, sanitation, and surveillance.
The Hickman catheter (HC) used in this protocol is a tunneled CVC with a polyester cuff and one to three lumens, that is commonly used for long-term intravenous access in humans and animals1,4,8,9. The tunneled catheter approach has been associated with lower complication rates and maintenance costs relative to non-tunneled variations10,11,12. The cuff reduces HC extrication by incorporating into the subcutaneous tissues surrounding the skin exit site. The multi-lumen design also enables the separation of medication administration and blood draws, thereby minimizing blood sample contamination and inaccuracy. Despite this, HC use is not without challenges, the most common of which include fracture, migration, occlusion, and infection13,14,15,16. Proper installation and maintenance of a HC are therefore indispensable skills when used in translational research. However, the current literature offers little guidance for best-practices for HC use in swine during long-term trials5,6,17.
The purpose of this study is to outline an optimized approach for HC insertion into the internal jugular vein (IJV), skin securement, and durable protection that minimizes long-term catheter-related complications and discomfort in swine. A discussion of the important considerations for HC use, potential challenges that may be encountered, and modifications that may improve the quality of this approach is included.
All animal procedures were conducted in accordance with an animal protocol approved by the Johns Hopkins University Institutional Animal Care and Use Committee (IACUC). Strains of male and female swine undergoing HC placement include miniature swine from the Massachusetts General Hospital (MGH) swine colony, Yucatan swine, and Yorkshire-crossed swine from an agricultural vendor (20-40 kg). Swine ranged from 3-10 months of age when the HC was placed. The HC may be placed anytime relative to the animal's experimental procedure. However, it is recommended to place it beforehand to permit the collection of baseline blood values. It is also recommended to give the swine at least a 1 week acclimation period before undergoing any experimental manipulation.
1. Preoperative planning
2. Intraoperative monitoring
3. Surgical preparation
4. Internal jugular vein identification and preparation
5. Catheter exit site preparation
6. Introduction and tunneling of the catheter
7. Insertion of the catheter
8. Securing the catheter
9. Postoperative care
10. Catheter maintenance
Over 100 swine have undergone successful HC insertion in our lab. The HC can be safely and correctly placed and secured in under 1 h with a surgeon, assistant, circulator, and anesthetist. The catheter pouch takes roughly 15-20 min to make. The technique is straightforward and easy to teach and has been performed by veterinarians, surgical residents, and medical students following supervised instructions.
HCs have remained in place without complications or revision for up to 8 months. In a recent representative cohort of 32 swine with endpoints ranging from 8 to 132 days, 78.13% of HCs remained patent until the experimental endpoint (Figure 11). Of the swine that had clinical indications that required euthanasia and diagnostic necropsy, the HC was documented with correct placement in the IJV without thrombi, debris, or signs of infection. Complication rates requiring intervention were modest: 9.38% of HCs required removal or replacement before 30 days and 12.5% required removal or replacement on or after 30 days. Additionally, 9.38% of HCs required small repairs to the original HC under sedation (Figure 11). Reasons for the line compromises were not always elucidated, but etiologies included displacement, puncture, and internal blockages (Table 1). Timely line repair and replacement have demonstrated a 100% functional success rate without significant interference with study data collection. In cases of suspected central line infection, swine underwent prompt appropriate medical treatment without further complication. There have been no HC-related mortality in swine.
Figure 1: Animal positioned in ventral recumbency. Please click here to view a larger version of this figure.
Figure 2: Ventral incision through skin and platysma to expose the internal jugular vein. Please click here to view a larger version of this figure.
Figure 3: Accessing the internal jugular vein. (A) Incision of the platysma, revealing the sternocleidomastoid. (B) Internal jugular vein medial to the sternocleidomastoid and lateral to the sternohyoid. (C) Coated and braided non-absorbable suture ties are positioned at the caudal and cranial ends of the isolated portion of the IJV. (D) Wet gauze is placed in the incision to keep the vessel protected and maintain coated and braided non-absorbable suture positioning while repositioning the swine. Abbreviations: SCM = sternocleidomastoid; IJV = internal jugular vein; SH = sternohyoid. Please click here to view a larger version of this figure.
Figure 4: Animal positioned in lateral tilt to allow access to both the dorsal catheter exit site and the ventral incision. Please click here to view a larger version of this figure.
Figure 5: Dorsal puncture through the skin to serve as the catheter exit site. Please click here to view a larger version of this figure.
Figure 6: Placement of the catheter. (A) Puncture of the skin at the catheter exit site on the dorsolateral neck ipsilateral to the target IJV. (B) Catheter entering the surgical site lateral to the IJV. (C) Insertion of the catheter into the distal end of the IJV with the aid of a vein pick. (D) The vein is secured around the catheter distally and the proximal segment is occluded with a coated and braided non-absorbable suture tie. (E) Closure of the platysma. (F) Closure of the skin. Abbreviation: IJV = internal jugular vein. Of note, deep dorsal sutures that are extensively knotted for security may protrude through the closure; this should not interfere with wound healing. Please click here to view a larger version of this figure.
Figure 7: Lateral radiograph of Hickman catheter placement. The arrow demonstrates its correct location above the right atrium. Please click here to view a larger version of this figure.
Figure 8: Securement of the catheter and bandage collar. (A) The catheter is sandwiched between layers of 1 in medical tape and at the fork where the two lumens divide. (B) Cotton roll is wrapped around the neck, alternating between cranial and caudal to the lumens. (C) The elastic adhesive bandage is wrapped around the neck with slits cut to secure the lumens centrally. (D) The two lumens are strung through the hole in the catheter pouch and the pouch is secured to the elastic adhesive bandage. Please click here to view a larger version of this figure.
Figure 9: Catheter pouch. (A) Top of the pouch. (B) Top flap (Flap 3) lifted to show middle flap (Flap 2). The deep portion of the pocket shows the cut edge of Flap 2. (C) Bottom of the pouch with a hole in the bottom flap (Flap 1) to reveal Flap 2. Please click here to view a larger version of this figure.
Figure 10: Design and assembly of the catheter pouch. (A) A long strip of elastic adhesive bandage tape is folded to create three flaps of equal length, with a residual loose tail. (B) The tail is folded over the end of the pouch to secure the exposed sticky folds. (C) With the tail on top, the flaps are numbered 1-3 from top to bottom. A strip of the long edge of the middle flap (Flap 2) is cut to create an inner pocket. (D) Sutures are used to join the flaps according to the numbers and suture positions around the pouch. (E) A hole is cut in the middle of Flap 3. (F) Top of the pouch. Please click here to view a larger version of this figure.
Figure 11: Hickman catheter outcomes in swine. N = 32. Experimental endpoints ranged from 8 to 132 days. Please click here to view a larger version of this figure.
Table 1: Hickman catheter outcomes and complications in swine. Abbreviation: HC = Hickman catheter. Please click here to download this Table.
While CVCs serve a spectrum of functions in large animal research, current literature lacks a consensus approach for safe and sustainable use in long-term trials over 30 days. This protocol's stepwise procedure for HC insertion, skin securement, and storage in a handmade pouch has undergone significant adjustments for quality improvement. As such, this protocol presents a technique for HC use that permits efficient and effective intravenous access while ensuring animal welfare and minimizing complications.
Clinical and research applications of this protocol
Swine are used in many areas of biomedical research, and their use as preclinical models has increased dramatically since the early 1980s18. Notably, many anatomical and physiological characteristics of swine closely resemble humans, making them a suitable model for many diseases and a predominant laboratory animal species for surgical and interventional protocols19. The cardiovascular, integumentary, urinary, digestive, and renal systems are the most common models in swine due to the comparative overlaps with humans. Swine are one of the primary models for surgical training in laparoscopic and endoscopic techniques, organ transplantation, toxicology, pharmaceutics, and biomedical device assessment18,20,21,22. The versatility of the HC makes it an invaluable tool in these clinical and research settings. They allow reliable access for serial bloodwork and are an efficient means of intravenous fluid and medication administration4, which avoid multiple sedations that can introduce confounding variables into studies and compromise subject health. Multiple studies have also shown their utility in regularly monitoring blood drug levels, such as tacrolimus values in immunosuppression protocols23. Some of the swine included in this protocol had HCs for immunosuppressive drug level monitoring and intravenous administration of immunosuppressive and/or immunomodulatory drugs in pigs undergoing vascularized composite allotransplantation (VCA) procedures, including a heterotopic hind limb transplant model24, a swine hemiface graft dissection and transplantation model, and a renal autotransplantation model.
Critical steps in the protocol
Appropriate line length and reliable line care are essential to extend the life of the HC. The catheter must be cut where it reaches one-third of the length of the sternum (protocol step 7.4), as shorter lines are at risk of displacement and longer lines are at risk of hitting the vessel wall. To maintain patency and prevent blockages, both lines must be flushed daily with normal and heparinized saline (protocol steps 9.2-9.3). The lines should also be systematically identified by color before each use to avoid switching the roles of the red (blood draw) and white (medication administration) lines to avoid damage. For example, high negative pressure during blood draw can collapse the white line and cause clotting, whereas drug administration through the red line can cause residual drug to persist within the line or the clave, thereby artificially raising the measured drug content of drawn blood. Despite the secure dressing technique detailed in this manuscript, swine are occasionally able to gain access to the lines and cause mechanical damage, such as chewing or scratching. If one of the lines becomes unusable, the dysfunctional line should be securely occluded by tying a double knot in the line after the fork (to retain access to the remaining patent line) to prevent accidental use or dislodgement of the blockage into circulation. The remaining patent line may be used to fulfill both line roles temporarily, but the dysfunctional line should be replaced as soon as possible via a Hickman Repair Kit. It is also important to protect the lines by securely storing them in the pouch (protocol step 8.3.5) after each use. To ensure proper fixation of the pouch, the collar must be secured to the skin at both the cranial and caudal end (protocol step 8.3.4), and the pouch secured to the collar along each edge (protocol step 8.3.8). Broken sutures should be replaced expeditiously.
Protocol challenges and solutions
HC-related complications include migration, displacement, and central line infection. If it is suspected that the internal tip of the catheter has migrated distally to press against the right atrium wall, plain radiographs can help locate the catheter tip. This complication is managed with removal and replacement of the HC using the contralateral IJV. In the swine that have HCs placed according to this protocol, the use of the contralateral IJV has not resulted in any complications related to animal health or research outcomes. This is likely because swine have a robust intracranial and facial venous network that permits drainage of the head via bilateral external jugular veins, whereas the IJV has a comparatively minor contribution to head drainage25,26. In other cases where the HC is displaced from its surgical position due to loosening of the catheter or collar attachments to the skin, broken sutures should be immediately replaced. If the subcutaneous cuff exits the skin, the HC should be replaced. Regarding central line infection, signs and symptoms in swine may include lethargy, new onset of cough, decreased appetite, pyrexia, leukocytosis, and raised or unraised petechiae. Any signs of infection should be addressed immediately with antibiotics under the direction of a veterinarian, and culture of any abscess fluid should be considered if the animal is on immunosuppression treatment. Risks of central line infection are reduced by creating secure catheter pouches, regularly sanitizing and flushing the lines, employing sterile practices, and being vigilant in identifying subtle or early signs of HC dysfunction. These include line leaking, bubbles during blood draw, and increasing difficulty flushing lines.
Comparison to other CVC protocols in swine
Percutaneous methods of gaining central venous access in the external jugular vein (EJV) via palpable landmarks have been described. This provides the benefit of a reduction in soft tissue disruption and postoperative pain but can lead to complications, such as inadvertent carotid artery puncture and hematoma formation27. In contrast to percutaneous modalities, the protocol detailed in this manuscript allows for direct visualization of the target structures, which can help reduce damage to nearby tissues. Additionally, while open procedures for cannulating the EJV have been reported28, access to deeper structures such as the IJV and detailed guidance on placement and troubleshooting are limited. Another study utilized a similar approach of a paratracheal incision to access the jugular trunk, but instead used a laparoscopic suctioning device to create a subcutaneous tunnel to pass the HC and protected the external portion with a fitted jacket29. The results from this study showed a higher infection rate and thromboembolic complications in swine with tunneled HCs as compared to another group with a subcutaneous vascular access port. While the source of these complications is likely multifactorial, the protocol described in this manuscript has shown few infectious complications and helps mitigate potential causes by securing the external catheter at multiple points, creating a multilayer collar, and using a catheter pouch for line protection.
Limitations
This study presents with some limitations. While the use of three different swine strains demonstrates HC procedure success in a diverse cohort, swine have little anatomic variability and low rates of anomalous vasculature19. As such, the use of the sternocleidomastoid and sternum as surgical landmarks provided a consistent technique for incision and catheter length, respectively. The development of an optimized approach for HC placement, skin securement, and protective storage has occurred over several years in parallel with protocol-based studies in swine. Sequential modifications have been made in response to animal observation and creative troubleshooting. Therefore, a detailed report on failed modifications, or the process by which the proposed technique was established, was not included. Moreover, the data analysis does not include a control comparison group, such as swine that underwent venipuncture under sedation. Additionally, as with any procedure, this technique requires surgical experience, practice, and appropriate training of less experienced team members. The assembly of the catheter pouch may also be subject to user error. Detailed diagrams are included that aim to effectively illustrate this technique. Further, as this technique involves prophylactic antibiotics, it may not be appropriate for studies that would be impacted by antibiotic administration. Finally, this approach to HC placement and outcomes is limited to use in swine. This technique may not demonstrate the same efficacy in other large animals with varying anatomies. While HCs have been used in other species, further research is needed to adapt this technique to other animals.
Conclusions
The HC is an effective method of regular blood monitoring and intravenous drug administration in swine. This study details our best practices for HC insertion, skin securement, and durable protection that minimizes HC-related complications and animal discomfort. Through years of technique modification and troubleshooting, this protocol details an optimized approach to HC use in swine, with high reproducibility and minimal complications. Finally, guidance is offered to prevent and resolve problems that may arise during the lifetime of the HC.
The authors have nothing to disclose.
We would like to acknowledge the support of the Army, Navy NIH, Air Force, VA, and Health Affairs regarding the AFIRM II effort under award CTA05: W81XWH-13-2-0052 and CTA06: W81XWH-13-2-0053. The U.S. Army Medical Research Acquisition Activity, 820 Chandler Street, Fort Detrick MD 21702-5014, is the awarding and administering acquisition office. Opinions, interpretations, conclusions and recommendations are those of the author and are not necessarily endorsed by the Department of Defense. In addition, we would like to acknowledge support from the Department of Defense Congressionally Directed Medical Research Programs (CDMRP), Reconstructive Transplantation Research Program (RTRP), through awards W81XWH-17-1-0280, W81XWH-17-1-0624, W81XWH-17-1-0287, and W81XWH18-1-0795. We would also like to acknowledge the Department of Plastic and Reconstructive Surgery and the Johns Hopkins University School of Medicine. Additionally, we would like to acknowledge the entire veterinary staff, including Melanie Adams, Karen Goss, Haley Smoot, Kayla Schonvisky, and Victoria Manahan.
#10 blade | Medline | MDS15110 | |
0.9% Sterile Sodium Chloride | Baxter | 2F7123 | |
0-0 Coated and Braided Nonabsorbable Suture | Covidien | S-196 | |
0-0 Synthetic, Monofilament, Nonabsorbable Polypropylene Suture | Ethicon | 8690H | |
1 inch Medical Tape | 3M | 1548S-1 | |
10 USP units/mL Heparin flush | Becton, Dickinson and Company | 306424 | |
3-0 Braided Absorbable Suture | Covidien | SL-636 (cutting needle), GL-122 (taper needle) | |
3-0 Monofilament Absorbable Suture | Covidien | SM-922 (cutting needle), CM-882 (taper needle) | |
4-0 Coated and Braided Non-absorbable Suture Ties | Ethicon | A303H | |
70% Ethanol | Vedco | VINV-IPA7 | |
Adson tissue forceps | MPM Medical Supply | 132-508 | |
Adson-Brown forceps | MPM Medical Supply | 106-2572 | |
Air warming blanket and pad | 3M Bair Hugger | UPC 00608223595770 | |
Backhaus towel clamp | MPM Medical Supply | 117-5508 | |
Brown needle holder | MPM Medical Supply | 110-1513 | |
Buprenorphine | PAR Pharmaceutical | 3003408B | |
Cefazolin | Hikma Farmacuetica (Portugal) | PLB 133-WES/1 | |
Chlorhexidine | Vet One | 501027 | |
Clave | Baxter | 7N8399 | |
Cotton Padding | Medline | NON6027 | |
Debakey forceps | MPM Medical Supply | 106-5015 | |
Elastic Adhesive Bandage Tape | 3M | XH002016489 | |
Halstead mosquito forceps | MPM Medical Supply | 115-4612 | |
Hickman Catheter | Bard Access Systems | 603710 | |
Hickman Catheter Repair Kit, 7Fr, Red and White Connectors | Bard Access Systems | 0601690 (red), 0601680 (white), 502017 | |
Kelly hemostatic forceps | MPM Medical Supply | 115-7014 | |
Ketamine | Vet One | 383010-03 | |
Lactated Ringers | Baxter | 2B2324X | |
Maropitant Citrate | Zoetis | 106 | |
Mayo scissors | MPM Medical Supply | 103-5014 | |
Metzenbaum scissors | MPM Medical Supply | 132-711 | |
Pantoprazole | JH Pharmacy | NDC 0143-9284-10 | |
Scalpel blade handle | Medline | MDS10801 | |
Vein Pick | SAI infusion technologies | VP-10 | |
Veterinary Ophthalmic Ointment | Dechra | IS4398 | |
Xylazine | Vet One | 510004 |