Portal circulation carries blood from the forestomach, stomach, intestines, gallbladder, pancreas, and spleen to the liver. Portal blood contains nutrients absorbed through normal digestive processes but may also contain ingested toxins and orally administered medications. Comparing compounds in the portal circulation versus systemic circulation could allow for better understanding of first-pass hepatic metabolism of orally administered compounds. In the US, a recent survey revealed that 70.8% of feedlots administer antimicrobials in the feed; this is a common administration method in cattle.1 With oral administration, the small intestine is the primary site of absorption.2 From the small intestines, drugs will enter into portal circulation and then the liver, where the drug may undergo metabolism and/or excretion. Therefore, sampling the portal circulation would allow for better understanding of the absorption and hepatic metabolism of orally administered drugs. In addition, the liver may be a target site for some medications. It is critical to understand drug concentrations at target sites to develop accurate and precise dosing regimens. Therefore, sampling the portal circulation may be an alternative to collecting liver tissue by biopsy or at necropsy.
Historically, portal vein centesis was first done through laparotomies.3 This was not only time and labor intensive but also an animal welfare concern. The first ultrasound-guided portal vein centesis was reported in 1989, with other recent publications demonstrating that portal vein centesis was feasible and safe.3,4 The first reported attempt of placing an indwelling portal vein catheter (PVC) was in 1952.5 Different research teams have modified this method for serial sample collection of the portal vein.6,7 In these studies, catheterizing the portal vein was feasible, but adverse reactions were noted. In the study conducted by Braun et al,6 cattle were lethargic after placement, but minimal necropsy lesions were observed. Further, the equipment used in these studies is not readily accessible.
Our primary objective was to explore the feasibility of placing a PVC and obtaining serial portal vein blood sample collections in steers using commercially available supplies. We hypothesized that the portal vein catheterization would be a successful continuous sampling technique with minimal adverse effects noted in the enrolled steers.
Methods
Animals used
This study was conducted with 2 groups of steers. The first group (n = 2) was a pilot study. This study was conducted with the primary goal of assessing the feasibility of placing a PVC and determining the hepatic enzyme changes and postmortem hepatic lesions after catheter placement. The second group (n = 8) was an experimental group in which the PVC was placed and the steers underwent a pharmacokinetic study (not discussed in this manuscript). In the pilot study, 2 Jersey cross-steers (6 months of age, weighing 139 to 146 kg) were used. In the experimental group, 8 Jersey cross- and Holstein cross-steers (6 to 7 months old, 159 to 225 kg) were used. This study was approved by the North Carolina State University IACUC (protocol 23-372). Steers were fed free-choice timothy hay and provided a pelleted livestock grain (Purina Cattle Stocker Grower) twice a day with continuous access to water. This feeding regimen is appropriate for this age group of steers.
Portal vein catheterization
In both groups, the portal vein catheterization was performed in the same fashion. The steers were sedated with 0.05 mg/kg xylazine (Pivatel, Anased; 20 mg/mL, IM; Animal Health International). A transabdominal ultrasound (ExaPad Veterinary Ultrasound Scanner; IMV Technologies) was performed to identify the portal vein. Once the portal vein was identified, the hair was clipped, and the site was aseptically prepped; typically, either the 10th or 11th intercostal space was used. Lidocaine (2%; lidocaine hydrochloride; Patterson Veterinary) (2 mL) was administered through the skin and superficial muscle layers. After diffusion of lidocaine, the ultrasound was repeated to confirm the location of the portal vein. A #15 surgical blade was then used to make a stab incision through the skin and superficial muscle layers. Under ultrasound guidance, a 14-gauge catheter (short-term catheter; 13 cm in length; MILA International Inc) was advanced into the lumen of the portal vein (Figure 1). Once there was confirmation of portal vein centesis on the ultrasound, the stylet of the catheter was removed, and a guide wire (Amplatz Super Stiff; 260-cm length, 0.035 inches diameter; Boston Scientific) was inserted into the lumen of the catheter (Figure 1). The guide wire was fed about 30 cm into the portal vein, and then the catheter was removed. An 8 French tissue dilator (Cook Medical) was fed over the guide wire to create an SC tunnel. The tissue dilator was removed, and it was followed by a 10 French tissue dilator (Cook Medical) to ensure the SC tunnel would be large enough to accommodate the vascular catheter. The 10 French tissue dilator was removed, and a vascular balloon catheter (XXL Vascular Balloon Dilation Catheter; 12-mm or 14-mm balloon diameter, 6-cm balloon length, 75-cm long; Boston Scientific) was fed over the guide wire (Figure 1). The vascular catheter was advanced until it was visualized on ultrasound in the portal vein. Once in the portal vein, 5 to 7 mL of sterile water was used to inflate the balloon; the catheter was then pulled back to visualize the balloon on ultrasound to ensure it was located within the lumen of the portal vein. Finally, the guide wire was removed, and injection caps were placed on both the balloon port and sample collection port. The catheter site was closed with a purse string, and a finger trap (2-0 Nylon; Covidien) was placed. The catheter was coiled against the body wall. Tape and suture were used to secure the catheter (Figure 1). In the pilot study, the PVC was wrapped with elasticon. This was not performed in the experimental group.
Placement of a portal vein catheter (PVC). From top left to bottom right. A 14-guage catheter is advanced into the portal vein via ultrasound guidance (A). A guide wire is placed through the catheter and visualized in the portal vein on ultrasound (B). The vascular balloon catheter is fed over the guide wire and observed in the portal vein via ultrasound as the balloon is inflated (C). The catheter is sutured in place at the skin (D).
Citation: American Journal of Veterinary Research 85, 12; 10.2460/ajvr.24.06.0178
Blood collection
In the pilot study, blood was collected from the jugular vein prior to placement of the PVC and 3 days following portal vein catheterization. This was accomplished through jugular vein venipuncture with an 18-gauge needle. Blood was collected in a vacutainer red-top tube (serum tubes; Becton-Dickenson). Blood was submitted to North Carolina State University Clinical Pathology Lab for biochemistry analysis.
Portal vein sample collection
In the pilot group, blood was acquired from the portal vein through the catheter. The catheter was first flushed with 6 mL of heparinized saline. Then, 12 mL of blood was aspirated through the sample collection port and discarded. Then, 6 mL of blood was collected and placed into a red-top tube (serum tubes; Becton-Dickenson). The sample was submitted to North Carolina State University Clinical Pathology Laboratory for biochemistry analysis. Blood was collected from both the jugular vein and the portal vein in separate red top tubes. This was to compare hepatic enzyme values at the different collection sites.
Portal vein catheter maintenance and monitoring
The PVC was flushed with sodium heparin flush twice a day. The catheter site was monitored twice a day by visual inspection as well as palpation of the catheter site. The steers also had rectal temperatures taken daily, along with daily physical examinations by a veterinarian (JLH).
Portal vein catheter removal
In the experimental group, the PVCs were removed immediately prior to their euthanasia. The purse string and finger trap were cut and removed, and the balloon was deflated. The catheter was then removed from the portal vein by pulling with gentle tension.
Necropsy
The pilot steers were euthanized 4 days following portal vein catheterization with pentobarbital (Euthasol; 0.22 mL/kg; Virbac) IV through the jugular vein. These steers then had a necropsy performed to assess hepatic lesions. The experimental steers were also euthanized (although for a different study). The time frame of their euthanasia in reference to portal vein catheterization varied depending upon the follow-up study they were enrolled in. Regardless, their livers were examined at necropsy for any gross lesions.
Results
All steers, in both the pilot and experimental group, tolerated the placement of the PVC well. In the pilot study, the PVC remained functional for the entire study period (4 days). In the experimental group, the PVC remained functional for the entire study period (7 days). Steers did not have elevated rectal temperatures at any timepoint, and the only abnormality noted was a local skin reaction. This was characterized as a superficial skin infection, with slight purulent discharge and some tenderness around the catheter site (n = 3). This was monitored, and no treatments were necessary.
Functionality of the PVC
The PVCs remained functional throughout the length of the study period in both groups. The most common issue with the PVC was kinking of the catheter at the skin catheter site. This was easily remedied by straightening the catheter and/or applying slight tension to the catheter. Blood samples were collected with no issues whenever they were required. The steers showed no signs of pain during sample collection. There were no adverse reactions or signs of pain associated with PVC removal in the experimental group.
Blood biochemistry parameters
To ensure the placement of the PVC did not interfere with hepatic function, a baseline biochemistry profile and follow-up biochemistry profiles were obtained. For the follow-up biochemistry profile, blood was sampled from both the jugular vein and portal vein. There were no significant differences or changes in hepatic enzymes (Table 1).
Blood biochemistry values.
| Presentation: jugular vein | Four days post-PVC: jugular vein | Four days post-PVC: portal vein | |||||
|---|---|---|---|---|---|---|---|
| Parameter | Reference range | Steer 1 | Steer 2 | Steer 1 | Steer 2 | Steer 1 | Steer 2 |
| Glucose | 45–75 mg/dL | 82 | 83 | 95 | 85 | 95 | 85 |
| Urea nitrogen | 20–30 mg/dL | 9 | 10 | 13 | 12 | 13 | 12 |
| Creatinine | 1–2 mg/dL | 1.2 | 0.5 | 0.7 | 0.7 | 1 | 0.7 |
| Phosphorus | 5.6–6.5 mEq/L | 10.8 | 739 | 9.9 | 9.4 | 10.5 | 12.9 |
| Calcium | 9.7–12.4 mEq/L | 10.4 | 9.7 | 10 | 9.7 | 10.2 | 11.8 |
| Magnesium | 1.8–2.3 mEq/L | 2.2 | 2.3 | 2.1 | 2.1 | 2.2 | 2 |
| Total protein | 6.74–7.46 g/dL | 6.2 | 6.3 | 6.2 | 5.9 | 6.1 | 5.6 |
| Albumin | 3.03–3.55 g/dL | 3.6 | 3.5 | 3.7 | 3.3 | 3.7 | 3.1 |
| Globulin | 3.0–3.48 g/dL | 2.6 | 2.8 | 2.5 | 2.6 | 2.4 | 2.5 |
| Total bilirubin | 0.01–0.5 mg/dL | < 0.2 | < 0.2 | < 0.2 | < 0.2 | < 0.2 | < 0.2 |
| Alkaline phosphatase | 0–488 IU/L | 166 | 174 | 135 | 165 | 137 | 156 |
| AST | 78–132 IU/L | 105 | 76 | 76 | 105 | 87 | 99 |
| GGT | 15–39 IU/L | 11 | 12 | 13 | 5 | 10 | 4 |
| SDH | 4.3–15.3 IU/L | 22.9 | 21.4 | 15.2 | 21.6 | 10.6 | 18 |
| Creatine kinase | 44–211 IU/L | 1,453 | 334 | 408 | 375 | 436 | 359 |
| Sodium | 132–152 mEq/L | 140 | 138 | 142 | 138 | 142 | 133 |
| Potassium | 3.9–5.8 mEq/L | 5 | 4.4 | 4.6 | 4.9 | 4.7 | 6.2 |
| Chloride | 97–111 mEq/L | 97 | 98 | 97 | 98 | 96 | 95 |
| Bicarbonate | 17–29 mEq/L | 26 | 27 | 29 | 27 | 28 | 27 |
| Anion Gap | 14–20 mEq/L | 22.3 | 17.9 | 21 | 18.4 | 22.4 | 17.6 |
Biochemistries were performed on the day of presentation, prior to placement of the portal vein catheter (PVC) in the pilot steers (n = 2). Biochemistries were performed again 4 days after PVC placement, with samples obtained from both the jugular vein and the portal vein (through the PVC). Individual steer values are demonstrated here.
SDH = Sorbitol dehydrogenase.
Necropsy results
For the pilot steers, a necropsy was performed with the PVC in place to confirm the placement and observe any changes in the hepatic parenchyma. For steer 1, there was gross discoloration of the hepatic parenchyma surrounding the PVC (Figure 2); however, the portal vein itself appeared to be normal (Figure 2). For steer 2, there was no discoloration noted, and the portal vein appeared to be normal (Figure 3). The steers in the experimental group were euthanized following a different study, and their PVCs had been removed prior to euthanasia. However, their livers were still examined for any gross changes; minimal scarring was seen where the PVC was placed in a few steers (n = 3).
Necropsy findings in steer 1. A—Demonstrates the gross discoloration of the liver at the PVC entry site. B—Demonstrates an indwelling portal vein vascular catheter (denoted by the white arrow) in the portal vein. There are no apparent gross abnormalities to the portal vein.
Citation: American Journal of Veterinary Research 85, 12; 10.2460/ajvr.24.06.0178
Necropsy findings in steer 2. A—Demonstrates the PVC entry site with no visual gross discoloration. B—Demonstrates the portal vein with no apparent gross abnormalities.
Citation: American Journal of Veterinary Research 85, 12; 10.2460/ajvr.24.06.0178
Discussion
In this study, we were able to successfully catheterize the portal vein with little to no adverse reactions in steers. We were able to demonstrate that portal vein catheterization does not alter liver function, and there are minimal changes noted on necropsy. The PVC can be removed with ease, indicating cattle could have 1 placed and then removed.
There are several papers available in the scientific literature regarding portal vein centesis and catheterization.3–7 In a recent study conducted by Braun et al,3 the overall objective was to determine any changes associated with portal vein centesis in cattle.3 These cattle had a single portal vein centesis event (for blood pressure measurement and hematological parameter assessment) and were then slaughtered; insignificant changes were observed on bloodwork and no postmortem lesions were identified. In a follow-up study,6 the same group placed percutaneous PVCs using a modified technique. In this study, to facilitate the placement of a PVC, a biopsy transducer was used to guide a steel cannula into the portal vein; the cannula was then used as a guide to place a ballon-tipped catheter in the portal vein. In this study, the PVCs remained functional for 9 to 15 days. This group reported behavioral changes in a few of the cattle, such as lethargy and anorexia. We did not note any of these changes in our study. This group also reported postmortem lesions consisting of fibrin at the cannula site, and 2 cows had a localized peritonitis. While we did see discoloration of the liver grossly in 1 steer, we did not find any cattle with local peritonitis. In our study and the study conducted by Braun et al,3 both PVCs were placed aseptically, and animals were sedated to allow for efficient and safe placement. The differences in placement technique—cannula by Braun et al3 versus a guidewire in our study—may explain the differences in adverse events that were seen. A different study team also used ultrasound to guide a PVC placement.7 This team performed a similar technique, but they did not use a balloon-tipped catheter, and extravasation was their most common complication. This research team did not report any other adverse reactions in cattle. These studies, and ours, all show that the portal vein can be successfully catheterized and the catheter maintained for prolonged periods of time, allowing for serial sample collection; each method, though, has its own drawbacks, and these should be considered when deciding upon which technique to utilize.
In this study, there were several limitations. Although the catheters were functional throughout the entire study period, the most common limitation in terms of functionality was kinking of the vascular balloon catheter at the catheter site. This was easily fixed; however, it was time consuming. A different type of vascular balloon catheter may be explored to see if a stiffer catheter would decrease the degree of kinking but still allow for animal comfort. The vascular ballon catheter was also positional; at times, it required tension to allow for blood sample collection. Again, this was easily remedied but could be considered time consuming. Finally, in some steers, there were superficial abscesses noted at the catheter site. This was not reported in any of the other studies. We only inspected the catheter site daily and cleaned once an abscess was noted; in the future, daily cleaning and management of the catheter site may reduce the number of superficial abscesses present.
Currently, our research team has used portal vein catheterization for pharmacokinetic studies (in preparation). This technique could be utilized in the future for more pharmacokinetic studies, especially in drugs where the liver is considered the target site. In addition, portal vein catheterization can be used for blood sample collection to study nutrient metabolism or mineral analysis.
In conclusion, the described PVC technique in this study was feasible and practical and allowed for sample collection up to 7 days, and longer for some steers. The equipment used was accessible, and there were minimal adverse reactions noted in the steers. This is a novel sampling technique that can be utilized safely in future studies.
Acknowledgments
The authors would like to acknowledge North Carolina State University’s Clinical Pathology Lab for sample analysis. They would also like to acknowledge students Camryn Kline, Julio Mojica Perez, and Ivonne Miranda Martinez for their help with this project.
Disclosures
The authors have nothing to disclose. No AI-assisted technologies were used in the generation of this manuscript.
Funding
The authors have nothing to disclose.
References
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